Regulation of epithelial cell migration and

macropinocytosis by septin GTPases

A Thesis

Submitted to the Faculty

of

Drexel University

by

Lee Dolat

in partial fulfillment of the

requirements for the degree

of

Doctor of Philosophy

June 2016

© Copyright 2015 Lee Dolat. All Rights Reserved.

ii

Acknowledgements

I thank my adviser, Dr Elias Spiliotis, for instilling in me a deep appreciation for cell

biology. Equally important, he provided me with countless opportunities to develop as a

scientist, explore my own ideas and interests, and accomplish everything I set out to do.

My labmates – Jon Bowen, Xiaobo Bai, Eva Karasmanis, and Jimmy Angelis -

have been exceptional colleagues and friends.

I thank my thesis committee for their critical comments on my work. Drs

Saunders, Stanford and Anandan also shared with me invaluable career advice, and I

am especially grateful for their support during the search for my next position.

Finally, I thank my family and friends for their help and support along the way.

iii

Table of Contents

List of Figures ................................................................................................................ v

List of Tables ............................................................................................................... vii

Thesis Abstract .......................................................................................................... viii

Chapter 1: Introduction ................................................................................................. 1

Septin structure, assembly and dynamics ................................................................... 1

Septins and the membrane skeleton ........................................................................... 6

Septins and the actin cytoskeleton .............................................................................. 8

Septins and the microtubule cytoskeleton ................................................................. 12

Chapter 2: Tissue-specific expression of septins in health and disease ................ 21

Septin expression and roles in embryogenesis ......................................................... 21

Cardiovascular system .............................................................................................. 23

Immune system ......................................................................................................... 25

Nervous system ........................................................................................................ 27

Reproductive system ................................................................................................. 31

Urinary and digestive system .................................................................................... 33

Respiratory system ................................................................................................... 35

Endocrine system...................................................................................................... 36

Integumentary system ............................................................................................... 37

Chapter 3: Septins in Cancer ...................................................................................... 40

Septins in hematological malignancies ...................................................................... 40

Altered septin expression in solid tumors .................................................................. 41

Septins in the tumor microenvironment ..................................................................... 44

Chapter 4: Septin overexpression: implications in tumor cell migration, trafficking and metabolism ........................................................................................................... 46

Septins functions in tumor cell migration ................................................................... 47

Septins in membrane traffic and metabolism ............................................................. 50

Chapter 5: Septins promote stress-fiber mediated maturation of focal adhesions and renal epithelial motility ........................................................................................ 57

Abstract ..................................................................................................................... 57

Introduction ............................................................................................................... 57

Materials and Methods .............................................................................................. 59

iv

Results and Discussion ............................................................................................. 68

Figures ...................................................................................................................... 74

Chapter 6: Septins 9 exhibits polymorphic binding to F-actin and inhibits myosin and cofilin activity ....................................................................................................... 88

Abstract ..................................................................................................................... 88

Introduction ............................................................................................................... 88

Materials and Methods .............................................................................................. 90

Results ...................................................................................................................... 96

Discussion............................................................................................................... 103

Figures .................................................................................................................... 106

Chapter 7: Septins promote macropinosome maturation and traffic to lysosomes by facilitating membrane fusion ............................................................................... 120

Abstract ................................................................................................................... 120

Introduction ............................................................................................................. 120

Materials and Methods ............................................................................................ 132

Results and Discussion ........................................................................................... 139

Figures .................................................................................................................... 140

Chapter 8: Conclusions and future directions ........................................................ 155

List of References ..................................................................................................... 162

Appendix .................................................................................................................... 190

Vita ............................................................................................................................. 192

v

List of Figures Septin filament assembly ............................................................................................... 16

Septins regulate membrane organization, shape and dynamics .................................... 18

Septins interact with the actin and microtubule cytoskeleton ......................................... 19

Septin functions in organ systems and their connection to human disease .................... 39

Modes of single tumor cell migration ............................................................................. 55

Macropinosome formation, maturation and traffic to endolysosomes ............................. 56

Septin filaments interface with RSF and TA stress fibers in the leading lamella of renal

epithelia ......................................................................................................................... 74

Septins regulate the organization of the lamellar actin network and are required for the

stabilization of nascent FAs ........................................................................................... 76

SEPT9 functions as an actin cross-linking protein in FA maturation .............................. 79

SEPT9 promotes the motility of renal epithelia during EMT ........................................... 81

Lamellar septin fibers are composed of SEPT2/6/7/9 .................................................... 83

Quantification of septin knockdown and overexpression in MDCK cells ......................... 85

Septin depletion does not affect the levels of pSer19 myosin II RLC; myosin II N93K

rescues FA phenotype in septin-depleted cells .............................................................. 86

SEPT9 bundles F-actin through the B-domain of its NTD ............................................ 106

The NTD and B-domain of SERPT9 bind to three sites on the surface of F-actin ........ 108

SEPT9 inhibits the activity and interaction of myosin with actin ................................... 109

SEPT9 reduces the extent of cofilin-driven F-actin depolymerization ........................... 111

Overall 3D reconstruction of F-actin decorated with the NTD suggest that sites of SEPT9

binding to the actin filament ......................................................................................... 113

Convergence of structural classes validates model-based crosscorrelation sorting ..... 114

Reference free 2D averages of occupied classes ........................................................ 115

vi

Comparison of the frequencies of structural classes upon cross-correlation sorting of F-

actin segments in the presence or absence of the A-domain of SEPT9 ....................... 116

Full length SEPT9 and SEPT9 B-domain do not significantly affect F-actin bundling over

concentration ranges that inhibit myosin activity .......................................................... 117

Cosedimentation assay of full length SEPT9 with F-actin ............................................ 118

Resolution determination ............................................................................................. 119

Septins localize to macropinocytic vacuoles and tubule-vesicular endosomes, and their

contact/fusion sites ...................................................................................................... 139

Septins associate preferentially with maturing macropinosomes in a PI(3,5)P2-

dependent manner ...................................................................................................... 141

Septin depletion decreases macropinocytic fusion events and hinders macropinosome

maturation and turnover .............................................................................................. 143

Septins are required for the lysosomal delivery of fluid phase cargo ............................ 145

Septins are required for fusion of macropinosomes/endosomes with lysosomes, and

promote fusion directly in an in vitro reconstitution assay ............................................ 147

PM-mCherry and SEPT2 localization with respect to Rab5 and Rab7 ......................... 149

SEPT2 knock-down increases macropinosome size and number, but does not disrupt

colocalization of macropinocytic cargo with Rab5 and Rab7 ........................................ 151

SEPT2 does not colocalize with Sec6/8 and Exo70 at the contact sites of

macropinosomes/endosomes ...................................................................................... 153

vii

List of Tables

Table 1.1 ....................................................................................................................................... 17

Table 2.1 ....................................................................................................................................... 38

viii

ABSTRACT

Regulation of epithelial cell motility and macropinocytosis

by septin GTPases

Lee Dolat

Supervisor: Elias T. Spiliotis, PhD

Epithelial cancers, or carcinomas, account for approximately 90% of all cancers.

Unregulated cellular proliferation drives tumor growth and expansion, while metastatic

behavior, or the motility of tumor cells to distant sites, underlies the majority of cancer-

related deaths. Thus, better understanding of epithelial cell biology will provide novel

anti-cancer therapeutics.

Septin GTPases are a novel component of the mammalian cytoskeleton. Human

septins are comprised of thirteen genes that are divided into four groups based on

sequence similarity. Septins bind and hydrolyze GTP in order to assemble into

heteropolymers and higher-order structures such as bundles, ring and patches. Septins

bind directly to the actin and microtubule cytoskeleton and cell membranes. Through

these interactions, septins are posited to regulate their organization and dynamics by

acting as scaffolds and diffusion barriers. Importantly, septins are frequently

overexpressed in carcinomas, but their role in tumorigenesis and metastasis is unknown.

In the first part of this thesis, I investigated how septins function in renal epithelial

cell migration. Epithelial cell migration is driven by the actin cytoskeleton, which

assembles into a network of stress fibers that exert contractile forces at focal adhesions

with the extracellular matrix (ECM). Here, I discovered that septin 9 (SEPT9) crosslinks

directly actin filaments and controls focal adhesions stability, which is essential for

productive epithelial migration. Importantly, I found that SEPT9 expression is

ix

upregulated during epithelial-to-mesenchymal transition (EMT), a critical step during

tumor cell invasion, and SEPT9 overexpression promotes renal carcinoma and renal

epithelial cell migration. These results are among the first evidence that septins are bona

fide actin crosslinking and suggest that septin overexpression may promote tumor cell

invasion.

In the second part of this thesis, I investigated the molecular interaction between

SEPT9 and actin filaments. I identified that a basic region in the SEPT9 N-terminus is

sufficient to crosslink actin filaments. In addition, SEPT9 binds to a common site on the

actin filament and interferes with the binding of the myosin motor domain and the actin-

severing protein cofilin. By inhibiting myosin binding and cofilin-mediated severing,

SEPT9 may protect nascent actin filaments from depolymerization. These results

provide the first evidence that septins regulate spatially the binding of actin-binding

proteins along the actin filament.

In the last part of this thesis, I identified a novel role of septins in epithelial

macropinocytosis, a form of clathrin-independent endocytosis that is frequently

upregulated in cancer and provides essential molecular intermediates for carbon

metabolism during tumor cell proliferation. Here, I found that septins localize to

macropinosome contact sites with endosomes in a phosphoinositide-specific manner.

Significantly, septins regulate the delivery of macropinocytic cargo to the lysosome by

promoting membrane fusion. These results provide the first evidence of a role for septins

in a clathrin-independent endocytic pathway and endocytic vesicle fusion.

Collectively, these findings advance our knowledge of septin cell biology and

provide new evidence for how septin overexpression may promote tumor growth and

metastasis through two different cellular pathways.

1

CHAPTER 1. Introduction

Similar to microtubules, actin microfilaments and intermediate filaments, septins

constitute a network of filamentous polymers, which are essential for the development

and physiological functions of eukaryotic organisms. Septin filaments associate with cell

membranes and the actin and microtubule cytoskeleton, exhibiting a structural and

functional interdependence that distinguishes them apart from the conventional

cytoskeleton. Septin filaments function as diffusion barriers and scaffolds that affect the

compartmentalization of cell membranes and the spatial organization and functions of

the microtubules and actin cytoskeleton (Caudron and Barral, 2009; Spiliotis and

Gladfelter, 2012).

Septins are guanosin-5’-triphosphate (GTP)-binding proteins that self assemble

into non-polar oligomers and polymers (Weirich et al., 2008; Zhang et al., 2000). Similar

to actin and microtubules, septin assembly is coupled to nucleotide-binding and

hydrolysis (Gasper et al., 2009). Resembling the hetero-polymeric nature of intermediate

filaments, septin filaments consist of various subunits, which in mammalian organisms

are encoded by 13-14 different genes with alternative splicing and translation initiation

sites that give rise to multiple septin isoforms (Caudron and Barral, 2009).

I. Septin structure, assembly and dynamics Classified under the P-loop superfamily of nucleotide-binding and hydrolyzing

proteins, which includes the Ras-like GTPases and the kinesin and myosin motors,

septins possess a highly conserved GTP-binding domain (G domain), which is

structurally characterized by α-helices and β-sheets that alternate between flexible loops

(Leipe et al., 2002). Septin G domains contain the G1 Walker A (GxxxxGKS/T), G3

Walker B (DxxG) and G4 (NKxD) motifs, which interact directly with atoms of the

2 guanine nucleotide. In addition to these signature motifs, the N- and C-termini of the

septin G domains are respectively characterized by a polybasic region, which binds

phosphoinositides (Casamayor and Snyder, 2003; Zhang et al., 1999), and a sequence

of variable length and unknown function termed the septin unique element (SUE)

(Versele and Thorner, 2005). Beyond these homology domains, septins differ

significantly in the length and sequence of their N- and C-terminal extensions, which

flank their G domains and consist of proline-rich and α-helical domains, respectively

(Luedeke et al., 2005).

Based on amino acid similarity, mammalian septin paralogs are categorized into

four groups: SEPT2, SEPT6, SEPT7 and SEPT9 (Dobbelaere and Barral, 2004;

Luedeke et al., 2005). The majority of septins fall under the SEPT2 group, which

includes SEPT1, SEPT2, SEPT4 and SEPT5, and the SEPT6 group, which consists of

SEPT6, SEPT8, SEPT10, SEPT11 and SEPT14 (Figure 1.1). Lacking a critical threonine

residue from their G domains, septins of the SEPT6 group are hydrolytically inactive and

always bound to GTP (Gulesserian et al., 2007; Sirajuddin et al., 2009). The remaining

septins are classified under the SEPT9 group, which comprises SEPT3, SEPT9 and

SEPT12, and the SEPT7 group, which contains SEPT7 isoforms (Figure 1.1). SEPT2,

SEPT7 and SEPT9 are nearly ubiquitously expressed, while SEPT1, SEPT3, SEPT12

and SEPT14 are tissue-specific . The remaining septins are expressed widely, but not

present in every tissue (Dolat et al., 2014a).

Septin polymerization is driven by the propensity of their G domains to homo-

and hetero-dimerize (Sirajuddin et al., 2007). In the presence of guanosine nucleotide,

septin G domains dimerize by interacting directly through their GTP-binding pockets (G

interface), which are oriented opposite to one another, or, alternatively, through their N-

and C-terminal helices (NC interface). Using the G and NC dimerization interfaces in

3 tandem, septin monomers assemble linearly into non-polar oligomers (Sirajuddin et al.,

2007) (Figure 1.1). In mammalian systems, the basic septin oligomer is a hetero-

octamer composed of septins from each of the SEPT2, SEPT6, SEPT7 and SEPT9

groups at a 2:2:2:2 stoichiometry (SEPT9-SEPT7-SEPT6-SEPT2-SEPT2-SEPT6-

SEPT7-SEPT9) (Kim et al., 2011; Sellin et al., 2011b). This mode of hetero-octameric

assembly is conserved between mammals and yeast (Bertin et al., 2008; Farkasovsky et

al., 2005), septin filaments could also consist of hetero-hexameric (SEPT7-SEPT6-

SEPT2-SEPT2-SEPT6-SEPT7) and –tetrameric (SEPT6-SEPT7-SEPT7-SEPT6) units

(Engidawork et al., 2003a; Kinoshita, 2003). In addition, “non-canonical” hetero-

oligomers that consist of multiple subunits of the same septin group may also exist

(Beites et al., 1999; Blaser et al., 2002; Hsu et al., 1998; Martinez et al., 2004; Shinoda

et al., 2010). Overall, septin hetero-oligomers could vary in size and composition

depending on cell and tissue type. Irrespective of subunit identity, presence of GDP and

thus GTP hydrolysis favors oligomerization by stabilizing the G interface, while GTP-

binding appears to destabilize the NC interface (Gulesserian et al., 2007). With the

exception of the SEPT6 group septins, which cannot hydrolyze GTP, oligomeric septins

are GDP-bound and septin polymers contain an excess of GDP relative to GTP

(Farkasovsky et al., 2005; Sirajuddin et al., 2007). Therefore, in contrast to microtubules

and actin microfilaments whose polymerization is favored by a non-hydrolyzed

nucleotide, septin polymerization is favored by GTP hydrolysis.

Septin hetero-octamers polymerize into higher order structures such as linear

and curved filaments, rings and gauze-like meshworks (Bertin et al., 2008; Garcia et al.,

2011; Kinoshita, 2002) (Figure 1.1). End-to-end binding of septin hetero-octamers

results in filaments, which are 5-10 nm wide and up to several microns long (Bertin et al.,

2008; Bertin et al., 2010; Kinoshita, 2002). Septin filaments pair with one another by

4 interacting laterally through the C- (coiled coil domains) and possibly N-terminal

extensions that stretch outwards from the plane of oligomerization (Bai et al., 2013;

Bertin et al., 2008; de Almeida Marques et al., 2012; John et al., 2007). Thus, septin

bundles form by lateral stacking of individual septin filaments and adaptor proteins

further enhance septin bundling (Sadian et al., 2013). Owing to the flexibility of the

SEPT2 NC interface, which behaves like a hinge, septin oligomers can bend within a 25-

30o angle of one another generating a curvature, which allows for the formation ring-like

structures with 0.5 – 0.7 µm diameter (Sirajuddin et al., 2007). Septin subunits that

enhance the flexibility of NC interface favor the formation of septin rings and post-

translational modifications can promote the assembly of a gauze-like meshwork of

orthogonally arranged filaments (Garcia et al., 2011). The filamentous pattern of septins

is further influenced by their association with membrane bilayers and the actin and

microtubule cytoskeleton. Giant unilamellar vesicle membranes enriched with

phosphatidylinositol-4,5-bisphosphate can promote the formation of tightly paired septin

filaments (Tanaka-Takiguchi et al., 2009) . Conversely, actin-templated assembly of

septins favors the formation of linear filaments as opposed to septin rings, which

become more prevalent upon actin depolymerization (Gulesserian et al., 2003;

Kinoshita, 2002). Similarly, the disk-like organization of septins in non-adherent cells

depends on microtubules (Engidawork et al., 2003b; Sellin et al., 2011a).

Septin filaments are less dynamic than microtubules and F-actin (Table 1.1).

Half-times of subunit exchange and turnover could vary from 10s of seconds for

cytoplasmic septin filaments in mammalian cells to 3 minutes for membrane-associated

septin polymers in yeast organisms (Dobbelaere et al., 2003; Engidawork and Lubec,

2003; Gulesserian et al., 2003; Hagiwara et al., 2011; Lubec and Engidawork, 2002).

Recent studies show that membrane-bound septin filaments grow by end-to-end

5 annealing (Bridges et al., 2014). Rod-shaped septin oligomers diffuse, collide and

anneal to the free ends of septin filaments, which do not exchange any subunits along

their length (Bridges et al., 2014). In contrast to membrane-bound septin filaments,

which incorporate new subunits after fragmentation, cytoplasmic septin filaments are

more dynamic, exchanging subunits along their length (Bridges et al., 2014; Engidawork

and Lubec, 2003; Gulesserian et al., 2003). Septin filament dynamics are affected by

post-translational modifications such as phosphorylation and acetylation, and vary

depending on the phase of the cell cycle (Hernandez-Rodriguez and Momany, 2012).

For example, in the budding yeast Saccharomyces cerevisiae, membrane-bound septin

filaments are dynamic in G1, but highly stable during the S, G2 and M phases of the cell

cycle (Dobbelaere et al., 2003). These changes in septin dynamics are mediated by

septin-dependent kinases and phosphatases that are recruited to septin filaments in the

beginning and end of the cell cycle, respectively (Dobbelaere et al., 2003).

The molecular mechanisms that regulate septin assembly and dynamics are not

fully understood. Recent advances indicate that regulation of GTP hydrolysis, septin

synthesis and degradation as well as post-translational modifications and protein-protein

interactions influence the filamentous organization and dynamics of septins. Septin

folding is mediated by the cytosolic chaperonin containing TCP-1 (CCT), which also

controls the folding of actin and tubulin, and septin degradation involves E3 ubiquitin

ligases including parkin, the von Hippel-Lindau (VHL) tumor suppressor and the Ring

Finger protein 8 (RNF8) (Chahwan et al., 2013; Craven et al., 2006a; Dekker et al.,

2008; Zhang et al., 2000). While these factors could regulate the availability of septin

subunits for polymerization, proteins that promote the GTPase activity of septins (e.g.,

Orc6) and cross-link septin filaments (e.g., Gic1) can directly enhance septin assembly

(Huijbregts et al., 2009; Sadian et al., 2013). In contrast, Cdc42, a signaling molecule of

6 the Rho family of GTPases, induces the unbundling and dissociation of septin filaments

(Sadian et al., 2013). Post-translational modifications such as phosphorylation,

acetylation and sumoylation further modulate septin assembly and dynamics, but many

of the details remain unknown (Hernandez-Rodriguez and Momany, 2012).

II. Septins and the membrane skeleton

Septin filaments assemble on the cytoplasmic leaflet of membrane bilayers

forming a meshwork that resembles the spectrin-actin membrane skeleton. In yeast

cells, septins form filamentous rings and gauze-like patches in actin-free areas of the cell

cortex (Engidawork et al., 2001a; Engidawork and Lubec, 2001), while in mammalian

cells septin filaments associate with the actin membrane skeleton (Hagiwara et al.,

2011). These membrane-bound septin filaments restrict protein diffusion and affect the

shape and rigidity of cell membranes (Caudron and Barral, 2009; Spiliotis and Gladfelter,

2012). Septins and are posited to interact with phosphoinositides, yet their specificity for

different phosphoinositides, which regulate the intracellular localization of membrane-

bound proteins, is not fully known.

In the budding yeast Saccharomyces cerevisiae, a membrane-bound septin ring

forms at the base of the emerging bud early in the cell cycle and evolves to a collar that

underlies the mother-bud neck (Engidawork et al., 2001a; Engidawork and Lubec, 2001).

Septin filaments are positioned between the plasma membrane and the smooth

endoplasmic reticulum (ER), impeding the lateral diffusion of cortical and ER membrane

proteins during polarized bud growth (Luedeke et al., 2005) (Figure 1.2 A). In mitosis,

splitting of the septin collar into two rings results in the formation of a membrane

compartment, which enables the accumulation of membrane-associated factors (e.g.,

vesicle tethering complexes) without diffusing into the mother and bud cell membranes

7 (Barral, 2000). During mammalian mitosis, a septin diffusion barrier is posited to similarly

limit the diffusion of inner leaflet membrane proteins across the equatorial plane of a

dividing cell (Schmidt and Nichols, 2004a).

Septins form membrane-bound rings at the base of primary and motile cilia and

the dendritic spines of neurons (Fliegauf et al., 2014; Gulesserian et al., 2001; Hu et al.,

2010; Tada et al., 2007). At the interface of the plasma and ciliary membrane, septins

control the localization of ciliary membrane proteins and thus, they are required for the

formation and maintenance of the primary cilium (Engidawork et al., 2001e; Hu et al.,

2010). Septins could similarly impede the lateral diffusion of post-synaptic proteins from

the dendritic spines to the shaft, aiding to the compartmentalization of synaptic receptors

and channels (Caudron and Barral, 2009). At the annulus of spermatozoa, septins form

a barrier between the principal and mid pieces of the sperm, and are required for the

proper localization of a protein that is essential for spermiogenesis (Engidawork et al.,

2001c; Ihara et al., 2005; Kissel et al., 2005). The mechanism by which septins restrict

lateral diffusion in cell membranes is unknown, but recent evidence indicates that

septins influence the organization of lipid domains and interact with the cytoplasmic tails

of membrane transporters (Hagiwara et al., 2011; Kinoshita et al., 2004; Sharma et al.,

2013b). Protein-lipid compartmentalization and domain identity could be further

reinforced by septin-mediated inhibition of vesicle delivery; septins interact with the

exocyst, a vesicle tethering complex, and components of the SNARE machinery of

vesicle fusion, and may introduce a physical barrier that interferes with vesicle delivery

to the plasma membrane (Engidawork et al., 2001b; Engidawork et al., 2001d;

Gulesserian et al., 2000; Yoo et al., 2001) (Figure 1.2 B). Hence, septins are an integral

component of the structure of cell membranes, contributing to the organization of

8 membrane domains by inhibiting selectively the lateral diffusion and delivery of

membrane proteins.

In addition to their diffusion barrier properties, membrane-bound septins affect

the shape and physical properties of lipid bilayers. Similar to the membrane-remodeling

functions of the Bin-Amphiphysin-Rvs (BAR)-domain proteins, septins have been shown

to induce the tubulation of spherical liposomes made of phosphatidylcholine and

phosphoinositides (Tanaka-Takiguchi et al., 2009) (Figure 1.2 C). These membrane

tubules have the diameter of septins rings (~0.5 µm) and are circumferentially braced by

septin filaments (Tanaka-Takiguchi et al., 2009). It is unknown, however, whether

septins affect membrane curvature and induce membrane tubules in vivo (Figure 1.2 D).

Conversely, several studies have shown that septin depletion decreases the rigidity and

stiffness of cell membranes (Mostowy et al., 2011; Tooley et al., 2009) (Figure 1.2 E). In

T lymphocytes, septins suppress membrane blebbing and protrusive activity is spatially

restricted to septin-free areas of the cell cortex (Gilden et al., 2012; Tooley et al., 2009).

Measurements of membrane elasticity and viscosity show a marked decrease in the

rigidity of cell membranes after septin depletion, which alters cell shape and morphology

(Mostowy et al., 2011). Thus, like the spectrin-actin membrane skeleton, septin filaments

contribute to the organization and mechanochemical properties of cell membranes.

III. Septins and the Actin Cytoskeleton

From yeast to mammals, septins are implicated in the organization and functions

of the actomyosin cytoskeleton. Septins colocalize with actin filaments and interact with

actin-binding proteins and signaling effectors that regulate the organization of the actin

cytoskeleton (Kinoshita, 2002; Mavrakis et al., 2014; Nagata and Inagaki, 2005) (Figure

9 1.3 A). Notably, septins interact directly with myosin II, regulating its localization and

activation in interphase and mitotic cells (Joo et al., 2007).

In the budding yeast S. cerevisiae, septins are essential for the recruitment of

proteins involved in actin cable assembly during bud growth (Figure 1.3 D). At the site of

bud formation, linear actin cables, which enable the polarized transport of vesicles, are

polymerized by the formins Bni1 and Bnr1 (Bretscher, 2013). In the absence of the

nonessential septin Shs1, Bnr1 is mislocalized and actin cables fail to assemble (Buttery

et al., 2012; Gao et al., 2010). The actin assembly activity of Bnr1 appears to depend on

Shs1 (Buttery et al., 2012) and the septin-interacting partner Bni5 is required for the

recruitment of myosin II, which drives actin cable retrograde flow (Nam, 2007). Thus,

septins scaffold and may activate the assembly of actin cables during bud growth.

Interestingly, mammalian septins scaffold the recruitment of cortactin, an activator of

Arp2/3, to actin patches in the axons of sensory neurons (Hu et al., 2012). Septin 6

(SEPT6) localizes to actin patches, which consist of branched actin filaments, and

SEPT6 depletion reduces the recruitment of cortactin to actin patches and their transition

to filopodia (Hu et al., 2012) (Figure 1.3 B and D). Reconstitution of branched actin

assembly in vitro and in the presence of Arp2/3 shows that SEPT6 localizes

preferentially to actin branch points (Hu et al., 2012). In agreement with this observation,

overexpression of SEPT6 increases the recruitment of cortactin to the lamellipodium, a

branched actin network at the leading edge of motile cells (Hu et al., 2012). Additional

septins (e.g., SEPT1, SEPT4) are enriched in the lamellipodia of squamous carcinoma

cells (Mizutani et al., 2013). Septins, therefore, have an evolutionarily conserved role in

scaffolding proteins involved in the formation of linear and branched actin filaments.

More recently, septins were found to cross-link and bundle actin filaments

directly, contributing to the assembly of contractile actin rings in Drosophila embryos

10 (Mavrakis et al., 2014). In developing Drosophila embryos, the blastoderm contains

thousands of nuclei, which become individual cells. Cellularization requires the assembly

of contractile rings of actomyosin filaments that furrow the plasma membrane resulting in

membrane canals that envelope the nuclei of the syncytial embryo. Embryos expressing

a septin mutant are characterized by a loss of circular actin filaments, which are less

bundled and contract at slower rates (Mavrakis et al., 2014). Drosophila septins bundle

actin filaments and copolymerization of actin with septins results in curved and circular

actin bundles (Mavrakis et al., 2014) (Figure 1.3 C and F). Mammalian septins also

bundle actin filaments in vitro and septin depletion results in loss of actin stress fibers

(Kinoshita, 2002; Kremer et al., 2007). These findings suggest that septins are actin-

binding and bundling proteins that regulate the organization of actin microfilaments.

However, the function of septins during actomyosin assembly is not fully understood.

In addition to their direct roles in actin organization, septins influence actin

organization and contractility by interacting with signaling factors and myosin II.

Assembly of stress fibers is under the control of Rho GTPases, which activate actin

nucleation factors and promote the contraction of myosin II (Chrzanowska-Wodnicka

and Burridge, 1996). Several studies have indicated that septins are involved in the Rho

signaling pathways that control stress fiber assembly. Septin 9 (SEPT9), which is

positioned at the terminal ends of septin hetero-octamers, interacts directly with a Rho

guanine exchange factor (GEF), termed the septin-associated Rho GEF (SA-Rho GEF),

and Rhotekin, a secondary effector of Rho signaling (Ito et al., 2005; Nagata and

Inagaki, 2005). Both SA-Rho GEF and Rhotekin regulate septin assembly and

localization to stress fibers. Interestingly, SEPT9 appears to negatively regulate Rho

activation by inhibiting GTP loading by the SA-Rho GEF (Nagata and Inagaki, 2005).

Septins also interact with Binders of Rho GTPases (Borgs), which are effectors of Cdc42

11 (Joberty et al., 2001). New studies show that Gic1, a yeast homologue of the human

Borg, binds and cross-links septin filaments in vitro in a Cdc42-dependent manner

(Sadian et al., 2013). Binding of the active Cdc42-GTP to Gic1 results in the dissociation

of Gic1 from septin filaments. The inactive Cdc42-GDP, however, also inhibits the

assembly of septin filaments by competing with Gic1 for binding to septins (Sadian et al.,

2013). In mammalian cells, Borgs have been shown to affect both septin and stress fiber

assembly (Joberty et al., 1999; Joberty et al., 2001).

While more work is needed to elucidate how the interactions between septins

and signaling molecules affect actin organization, structural and functional analysis of

the septin-myosin II interaction has provided a novel mechanism for the activation of the

non-muscle myosin II. The mammalian SEPT2 has been shown to interact directly with

the heavy chain of the non-muscle myosin II and yeast two-hybrid data indicate that

Myo1, the S. cerevisiae type II myosin, binds the sporulation-specific septin Spr3 (Drees

et al., 2001; Joo et al., 2007). Disruption of the SEPT2-myosin II interaction results in

loss of stress fibers in interphase cells and incomplete ingression of the cleavage furrow

during mitosis (Joo et al., 2007). These defects are accompanied by significant reduction

in the phosphorylation of the myosin light chain, which stimulates myosin II contractility.

Importantly, SEPT2 appears to scaffold the interaction of myosin II with the citron and

Rho-activated myosin kinases, which in the absence of septins dissociate from the

cleavage furrow (Joo et al., 2007). Thus, septins may control spatially and temporally the

localization and activation of myosin II. Importantly, this function might be evolutionarily

conserved as myosin activation and actin organization is also disrupted in Xenopus

embryos after Sept7 depletion, decreasing planar tension and protrusive activity during

the collective cell migration that drives body axis elongation (convergent extension)

(Shindo, 2014).

12

In summary, septins are integral components of the actomyosin cytoskeleton.

Recent studies have revealed that septins interact directly with actin filaments and

myosin II, but many of the molecular and functional details remain unknown. Future

studies will address the mechanisms by which septins affect the formation of linear and

branched actin filaments and myosin II-dependent contractility, and how septins

interface with signaling pathways that affect actin organization and contractility.

IV. Septins and the Microtubule Cytoskeleton

Septin filaments colocalize with microtubules in a diversity of eukaryotic

organisms and cells (Spiliotis, 2010). The extent of septin-microtubule colocalization

varies between septin paralogs and isoforms, and depends on cell type, phase of the

cell cycle and tubulin isoform expression and post-translational modifications. Recent

studies have shown that septins interact directly with microtubules, microtubule-

associated proteins (MAPs) and motors as well as enzymes that modify tubulin’s post-

translational modification. Thus, septins have emerged as novel regulators of

microtubule organization and dynamics, and may play a key role in the spatial regulation

of microtubule-dependent transport.

Mammalian SEPT2/6/7/9 complexes interact directly with microtubules through

their SEPT9, SEPT7 and SEPT6 subunits (Figure 1.3 I); SEPT2 does not bind

microtubules (Bai et al., 2013). It is unknown how SEPT6 and SEPT7 associate with

microtubules, but SEPT9 interacts with the acidic tails of βII-tubulin via its N-terminal

extension, which contains multiple repeats of the K/R-R/x-x-D/E motif (Bai et al., 2013).

Similar to the microtubule-binding sequence repeats of filamentous MAPs (e.g., tau,

MAP1), the SEPT9 repeat motifs consist of positively and negatively charged residues,

which interact electrostatically with one another and the acidic C-terminal tails of α-

13 tubulin (Bai et al., 2013). Thus, the N-termini of SEPT9 mediate septin-septin and septin-

microtubule interactions that cross-link microtubules into bundles. By forming

filamentous cross-bridges that bind microtubules in a staggered arrangement, septins

could increase the thickness and length of microtubule bundles as observed in vitro (Bai

et al., 2013). Consistent with this function, septins colocalize preferentially with

elongated microtubule bundles in cultured kidney epithelia (Bowen et al., 2011) (Figure

1.3 H). Moreover, expression of SEPT9 isoforms that lack N-terminal repeat motifs

reduces both microtubule bundling and septin-microtubule colocalization (Bai et al.,

2013). Hence, through the properties of SEPT9, mammalian septins function as bona

fide MAPs that facilitate the bundling of microtubules.

Like most MAPs, septins affect not only microtubule bundling, but also

microtubule dynamics. In kidney epithelia, SEPT2 knock-down increases microtubule

catastrophe and reduces the rate of microtubule growth, indicating that septins are

required for persistent microtubule growth (Bowen et al., 2011). Interestingly, loss of

SEPT2 also affects microtubule-microtubule interactions and the overall directionality of

microtubule growth (Bowen et al., 2011). In agreement with a septin requirement for

microtubule growth and bundling, SEPT9 knock-down decreased the density of the

microtubule network in mammary epithelia (Bai et al., 2013). Depletion of SEPT7,

however, has the opposite effect in HeLa cells, increasing microtubule density and

acetylation, a marker of microtubule stability (Kremer et al., 2005). A direct interaction

between SEPT2 and MAP4 indicates that cytoplasmic SEPT2/6/7 complexes inhibit

MAP4 from binding and bundling microtubules, but SEPT2/6/7 and MAP4 also compete

for binding to microtubules and a new study shows that SEPT7 mediates the interaction

of α-tubulin with the deacetylase HDAC6 (Ageta-Ishihara et al., 2013; Spiliotis et al.,

2008). Surprisingly, SEPT7 is required for α-tubulin deacetylation and SEPT7-depleted

14 neurons have increased levels of acetylated tubulin and reduced rates of microtubule

growth (Ageta-Ishihara et al., 2013). While the cause-and-effect relationship between

microtubule “stability” and acetylation is unclear, septins appear to affect microtubule

bundling, dynamics and post-translational modifications including acetylation and

polyglutamylation. Given that septins interact with α-tubulin, MAP4 and HDAC6 in a

paralog- and isoform-specific manner, it is possible that septins have distinctly different

roles, targeting different aspects of microtubule organization and dynamics.

Microtubule post-translational modifications and MAPs affect the binding and

processive movement of motors, generating a code for the spatial regulation of

intracellular transport (Verhey and Gaertig, 2007). Presence of septins on select

microtubule tracks (e.g., perinuclear microtubule bundles) raises the possibility that

septins regulate microtubule-dependent transport (Spiliotis et al., 2008). In kidney

epithelial cells, tubular vesicular carriers egress from the trans-Golgi network on septin-

coated microtubule tracks (Spiliotis et al., 2008). Microinjection of septin function-

blocking antibodies or SEPT2 depletion results in stalling of vesicle transport en route to

the plasma membrane (Spiliotis et al., 2008). Competition between SEPT2 and MAP4

for binding to microtubules suggests that septins could block the microtubule binding of

MAP4, which inhibits kinesin-dependent transport (Spiliotis et al., 2008). It is unknown

whether septins affect directly the binding and/or processive movement of microtubule

motors, but SEPT7 interacts directly with the mitotic motor CENP-E (kinesin 7) (Zhu et

al., 2008). Interestingly, septins are required for the proper localization of CENP-E and

consequently, for the stable capture and biorientation of the kinetochores of congressing

chromosomes (Spiliotis et al., 2005). Because SEPT7 interacts with the C-terminal tail of

CENP-E, which binds and inhibits the motor domain of CENP-E, SEPT7 could promote

the activation and motility of CENP-E by relieving its autoinhibition. Alternatively, septins

15 may scaffold the interaction of CENP-E with the Aurora A and B kinases or the protein

phosphatase 1 (PP1), which regulate CENP-E binding to microtubules; Aurora B has

been reported to interact directly with SEPT1 (Qi et al., 2005).

16

Figure 1.1. Septin filament assembly. Mammalian septins are classified in four groups

(SEPT2, SEPT6, SEPT7 and SEPT9) and characterized by highly conserved domains

such as a phosphoinositide-binding polybasic domain (PB), a GTP-binding domain, a

septin unique element (SUE) and C-terminal α-helical coiled-coil (CC) domains. Septins

form non-polar hetero-oligomeric complexes by dimerizing in tandem using their GTP-

binding domains, which interact with one another via their N- and C-terminal helices (NC

interface) or their GTP-binding pockets (G interface). GTP hydrolysis favors

oligomerization by stabilizing the G interface, but septins of the SEPT6 group do not

hydrolyze GTP. End-to-end assembly of septin hetero-oligomers results in the formation

of non-polar filaments, which interact laterally through the N- and C-terminal extensions

of their septin subunits. Owing to the flexibility of the NC interface, paired septin

polymers can bend into curved and ring-like filaments. In addition to septin bundles and

rings, membrane-associated septins form gauze-like structures, which consist of

orthogonally arranged filaments.

17 Table 1.1. Septin filaments in comparison to actin microfilaments, microtubules and intermediate filaments.

18

Figure 1.2. Septins regulate membrane organization, shape and dynamics.

(A) Septins form barriers to lateral diffusion of membrane lipids (orange) and proteins

(blue)

(B) Septins localize to membrane-membrane contact sites and regulate vesicle

docking and fusion.

(C) Paired septin filaments induce the tubulation of giant unilamellar vesicles in vitro.

(D) Septins can sense membrane curvature.

(E) Septins rigidify cell membranes by localizing adjacent to membrane protrusions.

19

20

Figure 1.3. Septins interact with the actin and microtubule cytoskeleton. (A – E)

Fluorescence images show purified recombinant septins tagged with the green

fluorescent protein decorating linear (A), branched (B) and circular (C) actin filaments.

Recent evidence indicates that septins crosslink actin filaments into curved and circular

bundles (Mavrakis et al, 2014) and scaffold the recruitment of cortactin, an actin

nucleation promoting factor, at Arp2/3-mediated branch points (Hu et al, 2012). In the

budding yeast Saccharomyces cerevisiae, septins are required for the localization and

activity of formins, which stimulate the formation of actin cables (D). In the axons of

sensory neurons, septins accumulate to actin patches (E), which consist of Arp2/3-

nucleated branched filaments, and in Drosophila embryos, septins are essential for the

assembly of the contractile rings of actomyosin (F), which drives membrane invagination

and furrow ingression during cellularization. Scale bars, 1 µm. (G – L) Fluorescent

images show colocalization of septin filaments with actin stress fibers (G) and

microtubules (H) in Madin-Darby canine kidney cells, and in vitro decoration of pre-

polymerized microtubules with purified recombinant SEPT7. Septins bundle

microtubules and inhibit their interaction with other microtubule-associated proteins (J).

Post-translational modifications (PTM) of tubulin such as acetylation and

polyglutamylation are influenced by septins, which can scaffold the interaction of tubulin

with the deacetylase HDAC6 (K). In addition, septins have been shown to guide spatially

microtubule-microtubule interactions and may influence the localization of microtubule

motors (L). Scale bars, 10 µm.

21

CHAPTER 2. Tissue-specific expression of septins in health and disease

Insights into the tissue expression and functions of mammalian septins have

emerged from DNA microarray analyses and septin knock-out mice. Serial analysis of

gene expression in a variety of human tissues shows unique expression of individual

septins (Table 2.1). SEPT2, SEPT7 and SEPT9 are ubiquitously expressed (Cao et al.,

2007; Connolly et al., 2011a; Hall et al., 2005a). While absent from some tissues, the

septins SEPT4, SEPT8, SEPT10 and SEPT11 are expressed widely (Cao et al., 2007;

Hall et al., 2005b). In contrast, expression of SEPT1, SEPT3, SEPT12 and SEPT14 is

limited to specific tissues. SEPT1 and SEPT3 are expressed in lymphoid and nervous

tissues, respectively (Hall et al., 2005a), and SEPT12 is a testis-specific septin (Steels et

al., 2007; Xue et al., 2004). SEPT14 was originally identified as a novel testis-specific

protein, but is also expressed in the nervous system (Peterson et al., 2007; Shinoda et

al., 2010). Little is known about how septin expression varies throughout development

and adult life, but a recent study showed that the levels of ten different septin proteins

change dramatically during brain development (Tsang et al., 2011).

I. Septin expression and roles in embryogenesis

To date, knockout mice have been generated for seven of the thirteen septins.

Genetic ablation of the ubiquitous Sept7, Sept9 and Sept11 resulted in embryonic

lethality. Sept7-/- mice were never born, arresting early in development possibly due to

mitotic failure (Kinoshita, 2008). Development of Sept11-/- mice ceased at day 11.5 in

utero and the mice died by day 13.5 (Roseler et al., 2011). Despite the development of a

healthy yolk sac, heartbeat and blood vessels, Sept9-/- embryos died by day 10 of

gestation, exhibiting mesenchymal tissue degeneration and extensive cell death

22 (Fuchtbauer et al., 2011). Surprisingly, loss of the neuron-specific Sept3 did not have

any discernible effects on the physiology and function of Sept3-/- mice (Tsang et al.,

2008). Functional redundancy between Sept3 and Sept9 or compensation of Sept3 loss

by Sept9 isoforms could account for the lack of phenotype. Similarly, Sept5-/- and Sept6-/-

mice were reported to be normal, but newer behavioral assays indicate changes in the

social interaction, rewarded goal approach and anxiety-related behavior of Sept5-/- mice

(Ono et al., 2005a; Suzuki et al., 2009). From all the septin knock-out mice, Sept4-/- are

the most well-studied and characterized. Male Sept4-/- mice are sterile due to a

morphologically altered and immotile sperm (Ihara et al., 2005; Kissel et al., 2005).

Neurologically, the cerebellum of Sept4-/- mice is mildly maldeveloped and hypo-

dopaminergic phenotypes such as an increase in the inhibition of the startling response

by weaker pre-stimuli have been reported (Kinoshita, 2008). Loss of Sept4 from mice

that express the alpha synuclein mutant A53T, which is common among familial forms of

Parkinson’s disease, increased amyloid deposition and neurodegenerative pathology,

leading to shorter lifespans (Ihara et al., 2007). Moreover, Sept4-/- mice are more

susceptible to tumor growth and liver fibrosis, and contain high levels of hematopoietic

and hair follicle stem cells (Fuchs et al., 2013; Garcia-Fernandez et al., 2010; Iwaisako

et al., 2008).

The embryonic lethality phenotype of the Sept7-/-, Sept9-/-, Sept11-/- mice

suggests that septins play an essential role in embryogenesis. It is unknown how septins

are involved in the embryonic development of mammals, but studies in model organisms

such as the fruit fly Drosophila melanogaster, the roundworm Caenorhabditis elegans

and the frog Xenopus lavis have shed some light on the developmental functions of

septins. In early Drosophila development, multiple rounds of mitotic nuclear divisions

lead to the formation of a multinucleated single cell embryo. The nuclei of this syncytial

23 blastoderm are partitioned into individual cells by a process termed cellularization. In

septin-deficient embryos, cellularization is incomplete resulting in multinucleated cells,

less imaginal discs (precursors of epithelial tissues) and larval death (Adam et al., 2000;

Neufeld and Rubin, 1994). In C. elegans, septins localize similarly to the cytokinetic

furrows, but are not essential for embryonic development (Nguyen et al., 2000). Septin-

depleted embryos rarely fail to complete cytokinesis, but the asymmetric geometry of

furrow ingression is disrupted and the contractile apparatus is less robust and more

prone to stochastic errors (Maddox et al., 2007). Post-embryonic development of C.

elegans tissues is more severely affected by septin mutations, which disrupt

gonadogenesis and the formation of a functional sensory and motor nervous system

(Finger et al., 2003; Nguyen et al., 2000). Recently, a study in Xenopus showed that

septins are involved in the planar cell polarity (PCP) signaling pathway, which directs the

collective cell movements of embryogenesis during gastrulation, axial elongation and

organogenesis. The PCP signaling protein Fritz, which interacts directly with Sept2, was

shown to control septin localization to the cortical membrane of gastrulating cells (Kim et

al., 2010). Moreover, Fritz and septins synergized toward the formation of cilia, which

are critical for the transduction of the Sonic Hedgehog signals that regulate

organogenesis (Kim et al., 2010). Thus, septins are essential components of the

molecular and cellular mechanisms that give rise to complex organ and tissue systems.

II. Cardiovascular System

The heart is the first organ to develop during embryogenesis. Terminal

differentiation of cardiomyocytes occurs near birth and is characterized by the cessation

of cell division and the development of contractile multinucleated cells. In mouse

embryonic cardiac cells, the levels of septin 2, 6, 7 and 9 levels are the highest in early

24 development and decrease at birth and adulthood (Ahuja et al., 2006). In embryonic

cardiomyocytes, septins localize to the cytokinetic ring and midbody of dividing cells and

are absent from sarcomeric actomyosin (Ahuja et al., 2006). Thus, septins could function

in early cardiac development by interacting with key components of the cytokinetic

machinery (e.g., myosin, anillin). Moreover, septin downregulation could trigger mitotic

arrest and the formation of terminally differentiated multinucleated cells. Interestingly,

SEPT5 is located within the 22qII locus that is commonly deleted in the DiGeorge/velo-

cardial-facial syndrome (DGS), which is characterized by cardial malformations and

lesions (Judith M. McKie, 1997). Studies in animal models suggest that 22qII

hemizygosity, which occurs in 90% of DGS cases, results in developmental impairment

of the right ventricle and outflow tract (Scambler, 2000). It is unknown if partial loss of

SEPT5 contributes to the pathology of DGS pathology, but decreased septin (SEPT8)

expression is also implicated in the toxic effects that anti-inflammatory drugs have on the

development of embryonic cardiomyocytes (Baek et al., 2010).

Septin expression and functions in the vascular network have been identified in

platelets, which are specialized secretory cells that regulate haemostasis, thrombosis

and injury repair. SEPT5 (CDCRel-1) localizes around platelet α-granules, storage

vesicles that contain growth factors and effector molecules involved in adhesion and clot

formation (Dent et al., 2002). Additionally, SEPT5 co-precipitates with syntaxin-4, a

component of the SNARE-SNAP complex that mediates vesicle fusion with the plasma

membrane (Dent et al., 2002). Platelets isolated from Sept5 null mice secrete serotonin

more readily upon stimulation, suggesting that Sept5 negatively regulates α-granule

fusion with the plasma membrane (Dent et al., 2002).

Abnormal SEPT5 expression has been linked to the Bernard-Soulier syndrome

(BSS), a rare autosomal recessive blood disorder characterized by excessive bleeding

25 and abnormal platelet morphology, count and secretion (Lopez et al., 1998). BSS

patients harbor mutations in the quaternary platelet glycoprotein (GP) Ibβ-IX receptor

subunit, which regulates platelet adhesion, activation and aggregation. A mouse model

lacking the IBβ gene recapitulates BSS pathology and isolated platelets contain

abnormally large α-granules and elevated levels of SEPT5 (Kato et al., 2004). More

recently, genotypic analysis of a juvenile patient with a rather severe case of BBS

showed homozygous deletions of the Ibβ and SEPT5 genes, which are positioned

directly next to one another (Bartsch et al., 2011; Zieger et al., 1997).

III. Immune System

The mammalian innate and adaptive immune systems encompass several

tissues and specialized cell types that defend against infectious agents. Despite that the

first mammalian septin (SEPT1) was cloned from lymphocytes twenty five years ago

(Carol., 1990), it was not until recently that researchers began to investigate the

immunological functions of septins.

Cells of the innate immunity (e.g. dendritic cells, macrophages) recognize and

engulf foreign bodies, and elicit a larger response through antigen presentation. SEPT10

was initially identified in dendritic cells, where it is abundantly expressed (Sui et al.,

2003). In contrast, SEPT10 is weakly expressed in the spleen, thymus and peripheral

blood leukocytes. In dendritic cells SEPT10 localizes to the cytoplasm and the nucleus,

and is transcriptionally upregulated upon stimulation with lipopolysaccharide (Sui et al.,

2003). Septins 2, 6, 8, 10 and 11 are abundantly expressed in macrophages, and

SEPT2 and SEPT11 localize to actin-based structures at the base and periphery of the

phagocytic cup (Huang et al., 2008). Disrupting septin assembly by expressing the

septin-binding domain of BORG3 or targeted knock down of SEPT2 or SEPT11 reduced

26 phagosome formation and uptake of IgG-coated beads (Huang et al., 2008). Further

studies are necessary to understand the precise functions of septins in dendritic and

macrophage cell biology.

T lymphocytes proliferate and mature in lymphoid tissues (e.g., thymus, spleen,

lymph nodes) and migrate to peripheral tissues, searching for antigen presenting cells.

Septins are essential for both T cell development and migration. During mouse T-cell

development, the transition from a double negative (CD4-CD8-) to a double positive

(CD4+CD8+) stage is accompanied by a 5- to 10-fold increase in Sept9 expression

(Lassen et al., 2013). Sept9 deletion affects T-cell development in the thymus leading to

an increase in stage 3 double negative cells (CD4-CD8-CD25+CD44-) at the expense of

stage 4 double negative cells (CD4-CD8-CD25-CD44-). Peripheral CD8+ and CD4+ T-

cells are reduced concomitantly and in vitro proliferation of Sept9-deleted CD8+ T-cells is

impaired; the effects of Sept9 deletion in vivo are more severe on CD8+ than CD4+ T-

cells (Lassen et al., 2013). These studies are the first to implicate septins in T-cell

development, but more work is needed to determine the mechanism by which SEPT9

regulates T-cell maturation and selection.

Lymphocytes migrate by forming a leading pseudopod and a trailing uropod, both

of which require polarized assembly of filamentous actin and contraction of the cortical

membrane. In T-cells, septins localize to the middle of the cell cortex and are distinctly

absent from the protruding pseudopod and uropod (Tooley et al., 2009). SEPT7-

depleted lymphocytes have elongated uropods and show poor persistent migration

despite maintaining normal levels of actin, microtubules and phosphorylated (active)

myosin (Tooley et al., 2009). In addition, SEPT7 depletion results in excessive blebbing

and reduces the rate of cortical retraction in a hydrostatic volume change assay, which

measures the ability of cells to expand and retract their cell membrane (Gilden et al.,

27 2012). In migrating lymphocytes, septins (SEPT6 and SEPT7) are absent from blebs

and filopodia, but are recruited to plasma membrane sites adjacent to these protrusive

structures (Gilden et al., 2012; Sellin et al., 2011a). Overall, the plasma membrane

organization and dynamics of migrating T-cells appear to depend on septins, which

affect persistent migration.

In non-immune cells, septins are essential for host defense against infectious

bacteria. The invasive bacterium Listeria monocytogenes enters mammalian cells

through the interaction of its surface proteins internalin A and B (InIA, InIB) with E-

cadherin and the Met receptor, respectively (Cossart, 2011). Targeted depletion of

SEPT2 inhibits bacterial invasion and impairs the InIB/Met signaling pathway (Mostowy

et al., 2009). Septins also thwart the intracellular mobility of the Shigella flexneri

bacterium, which propels itself through the cytoplasm by recruiting and activating the

Arp2/3 complex, which induces the polymerization of an actin tail (Egile et al., 1999).

Septin filaments are recruited to cytoplasmic Shigella and assemble cage-like structures

that inhibit bacterial mobility and facilitate the formation of an autophagosome for

bacterial degradation (Mostowy et al., 2010).

IV. Nervous system

Development of a functional nervous system involves the migration of embryonic

stem cells and neural precursors, and their differentiation into neurons and glial cells.

Neuronal morphogenesis requires the formation of axons, dendrites and synapses.

Recent studies have shown that septins play essential roles in neuronal migration,

axonal and dendritic arborization, and synaptic activity. In developing mouse embryos,

SEPT4 and SEPT14 are highly expressed in the cortical plate of the developing cortex,

which is formed by post-mitotic neurons that migrate tangentially from the subventricular

28 zone to the outer layers of the neocortex (Shinoda et al., 2010). In vivo knock down of

SEPT14 or SEPT4 expression and inhibition of their interaction perturb neuronal

migration to the cortical plate; neurons fail to extend leading processes and stall at the

ventricular and intermediate zones without reaching the cortical plate (Shinoda et al.,

2010). The sensory and motor neurons of the C. elegans septin mutants unc-59 and

unc-61 exhibit similar migration defects during larval development (Finger et al., 2003).

In cultured hippocampal neurons, septins are required for the formation of

dendritic spines (Cho et al., 2011; Tada et al., 2007; Xie et al., 2007). SEPT5/7/11

complexes localize to dendritic branch points and at the base of dendritic spines (Xie et

al., 2007). Targeted depletion of SEPT7, SEPT11 or SEPT6 reduces dendritic

arborization and alters the length, density and head morphology of dendritic spines (Cho

et al., 2011; Tada et al., 2007; Xie et al., 2007). In contrast, SEPT7 overexpression

increases dendritic protrusions and branchpoints (Tada et al., 2007; Xie et al., 2007).

Because dendritic spines are enriched with membrane proteins (e.g., neurotransmitter

receptors) that localize specifically at the post-synaptic cleft, septins are posited to form

a diffusion barrier at the base of dendritic spines blocking free diffusion of membrane

proteins in and out of the dendritic shaft (Caudron and Barral, 2009). In addition, septins

may also synergize with the actin and microtubule cytoskeleton for the formation of

dendritic spines.

Septin-mediated reorganization of the actin and microtubule cytoskeleton is

required for the collateral branching of axons. In sensory neurons isolated from chicken

dorsal root ganglia, SEPT6 and SEPT7 localize at the base of axonal filopodia and axon

branch points (Hu et al., 2012). This localization is reminiscent of septin complexes at

the base of dendritic spines, but in axons septins control the initiation and maturation of

collateral branches (Hu et al., 2012). SEPT6 enhances the transition of axonal patches

29 of F-actin to filopodia by enhancing the recruitment of the actin-binding protein cortactin,

an activator of Arp2/3-mediated actin polymerization (Hu et al., 2012). In contrast,

SEPT7 is involved in the entry of axonal microtubules into nascent filopodia, enabling

the formation of collateral branches (Hu et al., 2012). These regulatory roles of septins

are likely to influence not only the outgrowth and differentiation of neurites into axons

and dendrites, but also axon extension and growth cone motility. Indeed, septins have

been reported to affect axon length (Vega and Hsu, 2003), but more studies are needed

to understand their precise roles in axonal differentiation and growth.

Neurotransmission and organ innervation rely on the formation and function of

chemical synapses, which release and receive neurotransmitter molecules through their

pre- and post-synaptic clefts, respectively. Neurotransmitter secretion and uptake are

coupled to ion channels that sense and generate membrane potential. Initial studies

indicated that septins inhibit the release of synaptic vesicles (Beites et al., 2001), but

new evidence suggests that septins could also regulate the localization of ion channels

in membrane microdomains and their endocytosis from the cell membrane. The septins

3, 5, 6, 7 and 11 localize to the presynaptic cleft and their filamentous organization

coincides with the filament-like strands, which have been observed by electron

microscopy to connect synaptic vesicles with each other and the plasma membrane

(Hirokawa et al., 1989; Kinoshita et al., 2000; Tsang et al., 2011). SEPT5 and SEPT8

have been reported to interact with components of the molecular machinery of synaptic

vesicle fusion (Beites, 2005; Beites, 1999; Ito et al., 2009). SEPT5 and SEPT7 are

posited to inhibit vesicle fusion by inhibiting the formation of productive SNARE

complexes (Beites, 2005; Wasik et al., 2012). Interestingly, however, synaptic

transmission and morphology were not altered in Sept5-/- and Sept3-/- mice (Tsang et al.,

2008). Further analysis of the developing calyx of Held, which is a large glutamatergic

30 synapse of the auditory nervous system, showed that Sept5 filaments play a key role in

the positioning of pre-docked synaptic vesicles relative to the voltage gated calcium

channels (VGCCs) of the active zone (Yang et al., 2010). In Sept5-/- synapses, synaptic

vesicles are more tightly docked to the VGCCs. In addition, synaptic vesicles are more

readily and cooperatively released requiring a lower number of calcium channels to

trigger single fusion events (Yang et al., 2010). Thus, SEPT5 introduces a physical

barrier between synaptic vesicles and VGCCs regulating the efficiency of synaptic

transmission. A new study, however, suggests that SEPT5 and SEPT4 may modulate

synaptic activity by controlling the membrane organization and clustering of calcium and

possibly other ion channels (Sharma et al., 2013b). Although this hypothesis has yet to

be tested in neurons, SEPT4 is required for the organization of phosphatidyl-inositol-

(4,5)-bisphosphate-rich domains in the plasma membrane of HeLa cells and the

clustering of the calcium channel ORAI1, which triggers calcium entry and signaling

when calcium concentration is low in the endoplasmic reticulum (Sharma et al., 2013b).

Interestingly, the astrocyte glutamate transporter GLAST has been shown to interact

directly with SEPT2 (Kinoshita et al., 2004). SEPT2 mutations increase GLAST

internalization and decrease glutamate uptake (Kinoshita et al., 2004). Interaction of

septins (SEPT5, SEPT11) with dynamin, which mediates scission of vesicles from the

plasma membrane, raises the possibility that septins regulate endocytosis at synaptic

terminals, affecting the recycling of synaptic vesicles and ion transporters

(Maimaitiyiming et al., 2013; McNiven, 1998).

In summary, septins have fundamental roles in the development, activity and

connectivity of the nervous system. Thus, abnormalities in septin expression are likely to

compromise the function of the nervous system, contributing the pathology of

neurodevelopmental and neurodegenerative disorders. Hereditary neuralgic

31 amyotrophy, a disorder characterized by shoulder and arm pain and muscle atrophy, is

genetically linked to missense mutations and duplications in the N-terminal sequence of

SEPT9 (Chance and Windebank, 1996; Kuhlenbaumer et al., 2005). These

abnormalities may cause the axonal degeneration and focal demyelination observed in

the brachial plexus nerves of HNA patients (Ueda et al., 2010). In the maldeveloped

brain of fetuses with Down Syndrome, expression of SEPT6 and SEPT7 is markedly

reduced and SEPT4 might be hyper-phosphorylated due to over-expression of the

DYRK1A kinase (Sitz et al., 2008). Decreases in septin 7 and 11 expression were also

observed in Schizophrenia and bipolar disorders, and SEPT9 was abnormally

overexpressed in a mouse model of demyelinating neuropathy (Patzig et al., 2011;

Pennington et al., 2008). Histological analysis of brains from Alzheimer’s patients

identified SEPT1/2/4 in neurofibrillary tangles (Kinoshita et al., 1998), which are

intracellular aggregates of the hyper-phosphorylated tau protein. In addition, allelic

variations in SEPT3 isoforms have been observed in Alzheimer’s tissue samples

indicating a potential role or risk factor in AD pathogenesis (Takehashi et al., 2004). In

the brains of patients with Parkinson’s disease, SEPT4 is found in the Lewy body

aggregates of α-synuclein (Ihara et al., 2003). Interestingly, SEPT4 interacts with α-

synuclein directly, and loss of SEPT4 expression in the A53T familial Parkinson’s mouse

model increases amyloid-like aggregation and neurodegeneration (Ihara et al., 2007).

V. Reproductive system

Septins have been genetically linked to male infertility and ovarian cancer. The

importance of septins for sperm development was discovered by two independent

studies of SEPT4 knock-out mice. Male Sept4-/- mice are sterile and their sperm is

immotile and morphologically bent with an abnormal L-shape (Ihara et al., 2005; Kissel

32 et al., 2005). This phenotype correlates with Sept4 localization to the annulus, a cortical

ring separating the middle and principle pieces of spermatozoa. In Sept4-/- sperm, the

annular structure is abolished (Ihara et al., 2005; Kissel et al., 2005). Moreover, Sept4-/-

sperm mitochondria have abnormal size, cristiae and membrane morphology, and an

excess of cytoplasmic droplets is observed in the head and neck regions of the sperm

(Kissel et al., 2005). While these data suggest that Sept4 functions in mitochondrial

division and caspase-mediated removal of the cytoplasm during spermiogenesis, follow-

up studies explored the hypothesis that Sept4 maintains a cortical diffusion barrier at the

annulus. Hunnicutt and colleagues tracked the localization of the diffusing membrane

protein basigin, which localizes to the principle piece during spermatogenesis and shifts

to the middle piece during epididymal maturation (Kwitny et al., 2010). In the sperm of

Sept4-/- mice, basigin fails to localize to either the principle or middle pieces, indicating

that Sept4 is critical for maintaining the proper localization of sperm proteins through the

establishment of an annular diffusion barrier (Kwitny et al., 2010). Because a defective

annulus is common among patients with asthenospermia, screening for Sept4

localization is now used as a diagnostic tool for sperm malformation (Sugino et al.,

2008).

In addition to Sept4, septins 1, 6, 7 and 12 also localize to the annulus of the

sperm (Toure et al., 2011). Notably, Sept12 is expressed only in the testis (Steels et al.,

2007) and similar to the Sept4-/- mice, Sept12+/- chimeric mice are sterile and their sperm

is immotile with nuclear defects and distorted shape (Lin et al., 2009). In vitro fertilization

of mouse oocytes with Sept12+/- sperm has low success rates and fertilized oocytes do

not progress past the morula stage (Lin et al., 2011). Interestingly, genetic analysis of

infertile men revealed two missense SEPT12 mutations, T89M and D197N, which map

to the switch I and GTP recognition motifs of Sept12, respectively (Kuo et al., 2012).

33 Biochemical analysis showed that the SEPT12-T89M mutant had reduced rates of GTP

hydrolysis and SEPT12-D197N did not bind GTP (Kuo et al., 2012). Both mutants failed

to form filaments in tissue culture cells and SEPT12 was absent from the sperm annulus

of SEPT2-D197N patients (Kuo et al., 2012). Thus, mutations in the testis-specific

SEPT12 are genetically linked to male infertility.

Septin expression in ovarian tissue was first identified in Drosophila where sep1

localizes to follicle cells during embryonic development (Fares et al., 1995). The precise

function of septins in ovarian physiology is unknown. Misregulation of septin expression

has been identified in ovarian tumors and is discussed in the next chapter.

VI. Urinary and digestive systems

The tissue and organ surfaces of the urinary and digestive systems are

composed of epithelial cells. The epithelial plasma membrane is biochemically

differentiated into the apical and basolateral membrane domains. The directional

transport of water, ions and solutes depends on the proper localization of specific

channels (e.g., aquaporins, Na/K pump) and transporters (e.g., p-glycoprotein) to the

epithelial membrane domains. Establishment and maintenance of the polarized

distribution of these proteins is key for the development and function of organs such as

the kidney, intestine and lung.

In Madin-Darby canine kidney cells (MDCKs), a well-established model for

studying renal epithelial morphogenesis in 2D and 3D cultures, SEPT2 is essential for

the organization of the microtubule cytoskeleton and the transport of membrane vesicles

from the Golgi to the plasma membrane (Bowen et al., 2011). During the establishment

of columnar epithelial morphology, SEPT2 guides microtubule growth and microtubule-

microtubule interactions for the establishment of the subapical microtubule network.

34 Moreover, SEPT2 localization to a subset of perinuclear, microtubule bundles enables

the egress of Golgi-derived vesicles by hindering the binding of the microtubule-

associated protein 4 (MAP4), which impedes microtubule motor-driven transport .

Without proper microtubule architecture and post-Golgi membrane traffic, SEPT2-

depleted epithelia fail to assume a columnar shape and most apical and basolateral

membrane proteins accumulate in the cytoplasm. New studies suggest that septins are

also important for the two-dimensional expansion of the renal epithelium, which occurs

by planar division and asymmetric ingression of the cleavage furrow from the basal to

the apical membrane. In Drosophila epithelia, septins enable the closure of the

cytokinetic furrow by strengthening the cytokinetic actomyosin ring, which must

overcome the tensile forces exerted by the apical adherens junctions (Founounou et al.,

2013; Guillot and Lecuit, 2013).

Renal epithelia not only maintain water and ion homeostasis, but also monitor the

flow of fluid in the lumen of the nephron, which in turn can affect cell morphology and

proliferation. Fluid flow is sensed by the primary cilium, a microtubule-based antenna-

like organelle that projects from the epithelial apical membrane into the lumen of the

nephron (Pazour and Witman, 2003). Septins are integral and essential components of

the primary cilium (Chih et al., 2012; Ghossoub et al., 2013; Hu et al., 2010). In the inner

medullary collecting duct cells IMCD3, SEPT2 localizes to the base of the primary cilium

and is required for the formation of functional cilia (Hu et al., 2010). SEPT2 depletion

results in complete loss of primary cilia and the formation of shorter cilia, in which ciliary

membrane proteins diffuse freely into the surrounding plasma membrane (Hu et al.,

2010). Thus, SEPT2 is thought to be part of a membrane diffusion barrier, which

maintains the localization of ciliary membrane proteins. Recent studies, however,

35 indicate that SEPT2/7/9 complexes may also interact with axonemal microtubules

controlling the length of cilia (Ghossoub et al., 2013).

Podocytes are specialized epithelial cells that wrap around the capillaries of the

kidney’s glomeruli and respond to insulin. Podocytes prevent serum albumin and other

macromolecules from leaving the blood, while allowing water, salts and glucose to enter

the kidney lumen. Loss of podocyte function is prevalent in diabetic nephropathy, the

most common cause of renal failure (Pagtalunan et al., 1997). A new study shows that

SEPT7 colocalizes with the glucose transporter isoform 4 (GLUT4) storage vesicles and

interacts with nephrin, the CD2-associated protein CD2AP and the vesicle SNARE

protein VAMP2 (Wasik et al., 2012). SEPT7 knock-down increased the interaction of

VAMP2 with nephrin and syntaxin, and increased glucose uptake (Wasik et al., 2012).

Thus, SEPT7 could be involved in the regulation of glucose transport in the insulin-

sensitive podocytes.

VII. Respiratory System

Respiratory epithelia perform a number of key functions including gas exchange

and protection from particulate matter. In the lower respiratory tract, the multiciliated

bronchial epithelia protect the underlying tissue from noxious agents and push the

mucus toward the pharynx (Fahy and Dickey, 2010). Maintenance of the epithelial cell-

cell junctions is critical for limiting paracellular permeability, subepithelial inflammation

and injury (Frank, 2012). Physiological stresses in bronchial epithelia (e.g. particulate

matter) stimulate dynamic reorganization of the actin cytoskeleton at the cell cortex,

strengthening cell-cell junctions and restricting paracellular permeability (Youakim and

Ahdieh, 1999). In shear-stressed bronchial epithelia, SEPT2 is recruited to the apical

plasma membrane and interacts with the F-actin network more strongly (Sidhaye et al.,

36 2011). In SEPT2-depleted MDCK and bronchial cells challenged with shear stress or

particulate matter, cortical F-actin is reduced and low-molecular-weight dextran enters

into the basolateral space more readily (Sidhaye et al., 2011). Moreover, apical

application of EGF in SEPT2-depleted cells results in elevated activation of the

basolaterally-confined EGF receptor (Sidhaye et al., 2011). Taken together, these data

indicate that SEPT2 is critical for the maintenance of epithelial integrity during respiratory

stress.

VIII. Endocrine system

Tissues and cells of the endocrine system secrete hormones that regulate the

physiology and function of peripheral tissues and organs. In the pancreas, SEPT5 is

highly expressed in the islet of Langerhans, while its expression is markedly reduced

and absent from the acinar and ductal epithelia, respectively (Capurso et al., 2005). The

islet of Langerhans consists of α- and β-cells that secrete glucagon and insulin,

respectively, stimulating glucose release and uptake from the blood. The role of SEPT5

in exocytosis was tested in the HIT-T15 insulinoma cell line by assaying for the release

of exogenous human growth hormone (hGH) in the presence of wild-type or dominant-

negative SEPT5. Secretion of hGH was significantly reduced in cells expressing wild-

type SEPT5 indicating that SEPT5 inhibits exocytosis (Beites, 1999). In pancreatic β-

cells, SEPT2 interacts with EXO70, a component of the octameric exocyst complex that

mediates vesicle tethering to the plasma membrane (Rittmeyer et al., 2008). The

SEPT2-EXO70 complex localizes to the β-cell membrane and interacts directly with the

Rho GTPase effector, IQGAP1 (Rittmeyer et al., 2008). Interestingly, overexpression of

the SEPT2-EXO70 binding domain of IQGAP1 displaces SEPT2 from the plasma

membrane, and the constitutively-active form of CDC42 inhibits the interaction of

37 IQGAP1 with the SEPT2-EXO70 complex and blocks insulin secretion (Rittmeyer et al.,

2008). Although the precise role of septins in vesicle docking and fusion is unknown,

regulation of septin localization and function by IQGAP and Rho signaling might be

important for the secretion of insulin from pancreatic β-cells.

IX. Integumentary system

The integumentary system consists of the skin, hair follicles and sebaceous

glands. Collectively, these structures make up the largest organ system of the body

protecting all other organs from physical damage, infectious agents and water loss.

Regeneration of the epidermis after injury relies on skin and hair follicle stem cells,

whose growth and differentiation are tightly regulated so that tissue structure is

preserved without overgrowth. A recent study showed that the SEPT4 splice variant

ARTS (SEPT4_v2), is specifically expressed in hair follicle stem cells (HFSCs) (Fuchs et

al., 2013). Hair follicle bulges in mice lacking Sept4/ARTS are morphologically normal,

but contain twice as many CD34+ progenitor cells (Fuchs et al., 2013). Sept4/ARTS-

deficient CD34+ cells grow faster in culture without feeder cells and are highly-resistant

to pro-apoptotic stimuli, suggesting that SEPT4/ARTS restricts HFSC proliferation

(Fuchs et al., 2013). Remarkably, wound recovery in Sept4/ARTS-deficient mice is

extraordinarily fast. Sept4/ARTS has previously been shown to promote apoptosis by

targeting XIAP, an inhibitor of apoptosis, for degradation (Larisch et al., 2000). HFSCs

from mice deficient in both SEPT4/ARTS and XIAP showed reduced wound repair

(Fuchs et al., 2013) supporting the hypothesis that SEPT4/ARTS-mediated apoptosis

can limit stem cell growth.

38 Table 2.1 Septin expression, binding partners and links to human diseases.

39

Figure 2.1 Septin functions in organ systems and their connection to human disease Schematic diagram depicts main organ systems of the human body and outlines organ-

and cell type-specific structures and processes that require septins. Color codes indicate

the intracellular involvement of septins, which includes septin functions in the

organization of actin (red), microtubules (green) and cell membranes (grey), vesicle

fusion (blue) and apoptosis (yellow). Human diseases are listed based on genetic,

histological, genomic and proteomic evidence that implicates septins in the pathology of

various organ disorders.

40

CHAPTER 3. Septins in cancer

Septin expression is frequently altered in cancer, including tumors of epithelial

origin and hematological malignancies. On the other hand, increased assembly of

septins with actin stress fibers in cancer-associated fibroblasts, resident cells of the

tumor microenvironment, was shown recently to promote ECM remodeling,

angiogenesis and tumor cell invasion in a mouse model of breast cancer (Calvo et al,

2015). Septins are also emerging biomarkers for urological and colon cancers, but their

role in tumorigenesis and metastasis is poorly understood.

Septins in hematological malignancies

Aberrant septin expression has been implicated in blood disorders including

acute myeloid leukemia (AML) and acute lymphoblastic leukemia (ALL) (Cerveira et al.,

2011). MLL is a histone methyltransferase that regulates homeobox (HOX) gene

expression during hematopoiesis (Muntean and Hess, 2012). Chromosomal

translocations of the mixed lineage leukemia (MLL) gene account for ~10% of the de

novo and therapy-induced leukemias (Muntean and Hess, 2012). Septins 2, 5, 6, 9 and

11 have all been identified as MLL fusion partners, accounting for the largest known

protein family associated with MLL rearrangements (Cerveira et al., 2011). The MLL-

septin chimera results from double-stranded intronic breaks in the MLL and septin

genes, producing in-frame fusions of the MLL N-terminus and almost the entire open-

reading frame of the septin (Cerveira et al., 2011). Ectopic expression of the MLL-

SEPT6 chimera has been shown to immortalize hematopoietic progenitor cells,

upregulating the expression of the Hox-7a and Hox-9a genes (Ono et al., 2005b).

Although MLL-SEPT6 expression in mice was not sufficient to induce leukemogenesis

(Ono et al., 2005a), mice transduced with an active FMS-like receptor tyrosine kinase 3,

41 which is frequently overexpressed in AML, required MLL-SEPT6 to stimulate

leukemogenesis (Ono et al., 2005b). MLL-SEPT6 expression also led to a lethal

myeloproliferative disease with long latency (Ono et al., 2005b).

Recent studies have shown that hematopoietic stem cell maturation and

malignancy are affected by the SEPT4 splice variant ARTS (apoptosis-related protein in

the TGF-beta signaling; SEPT4_v2) (Garcia-Fernandez et al., 2010). SEPT4/ARTS

binds to inhibitors of apoptosis proteins, initiating caspase activation and cell death ,

2004(Gottfried et al., 2004; Larisch et al., 2000). Sept4-deficient mice contain elevated

levels of hematopoietic stem cells, including progenitor and immature B-cells that are

resistant to pro-apoptotic stimuli (Garcia-Fernandez et al., 2010). Genetic crosses

between Sept4-/- and the leukemogenic Eµ-Myc mice showed that SEPT4 ablation

enhances lymphomagenesis and reduces lifetime by 50% (Garcia-Fernandez et al.,

2010). Significantly, ARTS expression is frequently altered in bone marrow lymphocytes

derived from patients with childhood ALL, the most common malignancy found in

children (Elhasid et al., 2004). Lymphocytes from ALL patients exhibit a significant

reduction in SEPT4/ARTS mRNA levels, and SEPT4/ARTS-deficient lymphocytes are

highly-resistant to apoptosis when treated with the chemotherapeutic agent, ara-C

(Elhasid et al., 2004). DNA methylation inhibitors can rescue SEPT4/ARTS expression

suggesting that aberrant methylation downregulates its expression during tumorigenesis

(Elhasid et al., 2004). Clinically, SEPT4/ARTS expression was rescued in patient

lymphocytes following therapy, indicating that SEPT4/ARTS expression suppresses

growth and possibly malignant transformation of hematopoietic cells.

Altered septin expression in solid tumors

Septins are overexpressed in renal cell carcinoma (RCC), which is an aggressive

and highly metastatic cancer of the kidney with poor response rates to therapeutics

42 (Craven et al., 2006b). SEPT2 and SEPT11 are upregulated in RCCs defective in the

von-Hippel Lindau (VHL) tumor suppressor, a ubiquitin E3 ligase that targets the

hypoxia-inducible factor 1 (HIF-1) transcription factor for degradation (Craven et al.,

2006a). It is unknown if SEPT2 and SEPT11 are transcriptionally regulated by HIF1 or

directly degraded by VHL. Moreover, it is unclear if and how septin upregulation

contributes to RCC growth and metastasis.

SEPT9 is over-expressed in hormonally regulated breast, prostate and ovarian

cancers and endometrial cancer. SEPT9 binds directly to HIF-1 and increases its

transcriptional activity in prostate cancer cells by inhibiting HIF-1 degradation. Septin 9

expression, however, is significantly altered in ovarian cancers. Initial studies in sporadic

ovarian tumor samples showed that the chromosomal locus 17q25.3, which includes the

SEPT9 gene, was partially deleted leading to allelic imbalances and loss of

heterozygosity (Kalikin et al., 2000; Russell et al., 2000). Subsequent studies revealed

that SEPT9 has a complex genomic organization encoding for up to 18 different mRNA

transcripts and 15 protein isoforms (McIlhatton et al., 2001). Analysis of SEPT9 mRNA

transcripts in benign, malignant and borderline, low-grade ovarian carcinomas showed

that the SEPT9_v1 and SEPT9_v4* transcripts were overexpressed in serous and

mucinous borderline tumors (Scott et al., 2006). The enhanced translational efficiency of

SEPT9_ v4* relative to SEPT9_v4 suggested that increased levels of SEPT9_i4 could

promote the malignant and metastatic potential of ovarian carcinomas (McDade et al.,

2007). Consistent with this possibility, over-expression of SEPT_i4 enhances cell motility

and resistance to anti-cancer drugs that target microtubules (Chacko et al., 2012).

Upregulation of SEPT9_i1 could also enhance tumor growth and angiogenesis by

slowing down the degradation of the c-Jun-N-terminal kinase (JNK) and the hypoxia-

inducible factor HIF1a (Amir et al., 2009; Gonzalez et al., 2009). While the cause of the

43 amplification of certain SEPT9 mRNA transcripts is not understood, epigenetic changes

such as the hypermethylation of alternative intragenic promoters could alter the relative

expression of mRNA transcripts (Connolly et al., 2011b).

An increase in the mRNA and protein levels of the Bradeion isoform of SEPT4

has also been detected in urine samples from patients with RCCs and transitional

bladder cancers (Bongiovanni et al., 2012; Tanaka et al., 2003). Quantification of SEPT4

Bradeion expression by immunochromatography and RT-PCR of urine samples has

been introduced for the diagnosis of urothelial cancers in Japan and Europe

(Bongiovanni et al., 2012; Tanaka et al., 2003). Abnormal expression of SEPT4 has also

been observed in colorectal cancer (CRC). While absent from patient-matched healthy

tissues, two SEPT4 isoforms (Bradeion α and β) were identified in colorectal tissue

isolated from human patients with mucinous carcinoma and rectal malignant melanoma

(Tanaka et al., 2001). Ribozyme-mediated cleavage of either or both isoforms arrests

CRC growth in the G2 phase of the cell cycle in vitro and prevents tumor growth in mice

injected with CRC cells (Tanaka et al., 2002). The SEPT4 isoforms exhibit cytoplasmic

and nuclear localization (Tanaka et al., 2001), but further studies are necessary to

elucidate their cellular functions and the molecular mechanisms that contribute to their

abnormal expression in CRC.

In squamous cell carcinoma (SCC) and malignant melanoma tissue sections,

SEPT1 is significantly upregulated (Mizutani et al., 2013). In SCC cells, SEPT1 and

SEPT5 coimmunoprecipitate and colocalize with SEPT4 in the lamellipodia, a dendritic

actin network at the leading edge of migrating cells (Mizutani et al., 2013). SEPT1-

depleted DJM-1 cells exhibit defective cell spreading, a process that depends on actin

polymerization and cell adhesion with the ECM. While absent from healthy epidermis,

44 SEPT9 is highly expressed in cultured SCC cells and coprecipitates with

SEPT7/10/11/14 (Mizutani et al., 2013).

Conversely, SEPT9 is highly expressed in normal colonic glandular and surface

epithelia, but is markedly reduced in adenomas and almost completely absent from

tumor tissues, indicating that SEPT9 levels decrease progressively during tumorigenesis

(Toth et al., 2011). Treatment of HT29 CRC cells with a demethylating agent results in

increased levels of SEPT9, suggesting that SEPT9 expression is downregulated by

gene methylation during CRC progression (Toth et al., 2011). Epigenetic modifications in

stem cell DNA within the colonic crypt are known to change with age and have been

identified in a subset of CRCs (Humphries and Wright, 2008; Taylor et al., 2003). More

specifically, hypermethylation of CpG islands (GC-rich regulatory sequences found in the

promoters of almost half of all human genes) are common in many cancers (Herman

and Baylin, 2003). Plasma DNA from CRC patients contains abnormally high levels of

methylated SEPT9 (Grutzmann et al., 2008). Significantly, SEPT9 is hypermethylated in

87% of early stage I and II colorectal tumors (Warren et al., 2011). These findings have

led to the development of the septin 9 test, which is a commercial non-invasive, blood-

based test that screens for the methylation of the SEPT9 gene as a diagnostic of early-

stage colon cancer.

Septins in the tumor microenvironment

Surrounding the tumor mass lies a complex environment referred to as the tumor

microenvironment (TME), whose organization and constituent cells become altered

during tumor growth and metastasis (Balkwill et al., 2012). Cells of the immune,

vasculature and lymphatic systems, along with resident cells of the ECM (e.g.

45 fibroblasts) can function in part to support tumor growth by promoting angiogenesis,

growth and motility.

In particular, cancer-associated fibroblasts (CAFs) are a subpopulation of

“activated” cells that reside within the tumor microenvironment that promote tumor cell

transformation, angiogenesis and invasion (Kalluri and Zeisberg, 2006). During tumor

progression, CAFs become highly contractile through the increased the assembly of

actomyosin stress fibers, which promotes ECM remodeling by protein deposition and

upregulating cell-ECM adhesions (Butcher et al., 2009). Furthermore, tumor-associated

macrophages (TAMs) also promote cell invasion by localizing to the invasive edge of

tumors where they sustain a paracrine signaling loop that stimulates cell motility

(Patsialou et al., 2009).

In normal mouse NIH3T3 fibroblasts, septins colocalize with subnuclear actin

stress fibers (Kinoshita et al., 2002). Septin knockdown results in stress fiber

disassembly, while pharmacological targeting of actin stress fibers induces septin

filament assembly (Kinoshita et al., 2002). Conversely, septins and actin are largely

cytoplasmic in fibroblasts derived from mouse mammary glands, but co-assemble into

actomyosin stress fibers in CAFs (Calvo et al., 2015). Septin depletion inhibits

actomyosin stress fiber assembly in CAFs and reduces their ability to promote tumor

growth in vivo (Calvo et al., 2015). Thus, septins and actin stress fibers show an

interdependent relationship, but whether septins bind directly to actin filaments and how

septins regulates the organization of the actin stress fiber network during cell migration

has remained unknown.

46

CHAPTER 4. Septin overexpression in tumor cell migration,

trafficking and metabolism

As discussed in the previous chapter, septin overexpression is frequently

reported in solid tumors of epithelial origin including renal, breast, ovarian and skin; yet,

how septins contribute to tumorigenesis and tumor metastasis at the molecular level is

unknown. I hypothesize that septin overexpression promotes tumor growth and invasion

through at least two distinct mechanisms: i) promoting the mesenchymal-like motility of

renal epithelia, and ii) regulating the intracellular trafficking of macropinocytic cargo.

Because septins are multi-functional proteins that interact directly with the

cytoskeleton and cell membranes, it is not surprising that these findings implicate septins

in different cellular processes. Furthermore, tumorigenesis is a complex process that

stems largely from changes in the genome, which can alter many different cellular

pathways (Hanahan and Weinberg, 2000). Tumor cells proliferate unchecked by evading

cell death signals and adapt to growing metabolic needs by changing the mechanisms

by which energy is produced in order to sustain proliferation (Hanahan and Weinberg,

2011). Through changes in cell polarity, cytoskeletal organization and the trafficking of

intracellular cargo, tumor cells can invade the surrounding ECM and migrate towards

tissues of the blood and lymph to initiate metastasis (Bravo-Cordero et al., 2012).

Septins functions in tumor cell migration

Epithelial tumor cell invasion is activated by the aberrant induction of epithelial-

to-mesenchymal transition (EMT), a developmental transcriptional program that

regulates the patterning of epithelial tissues (Thiery et al., 2009). During this process,

epithelia breakdown cell-cell contacts, upregulate the attachment and turnover of

adhesions with the ECM, and adopt a front-rear, mesenchymal-like polarity (Thiery et al.,

47 2009). Tumor cell invasion into the ECM is initiated by membrane protrusions, such as

invadopodia and pseudopodia, which are driven by the assembly of actin filaments

against the ventral cell membrane (Murphy and Courtneidge, 2011). Within these

protrusions, actin-binding proteins, cell-ECM receptors and ECM-degrading enzymes

are enriched (Linder, 2007).

Tumor cell invasion into the ECM is a dynamic process. Tumor cells adapt to

ECM organization and topography, which becomes stiffer and more crosslinked during

tumorigenesis, and can interconvert between different modes of motility (Figure 4.1): a)

mesenchymal-like; b) amoeboid; c) lobopodial (Friedl and Wolf, 2010; Petrie et al.,

2012). The mesenchymal-like motility of epithelia has been characterized extensively

(Figure 4.1). At the leading cell edge, actin polymerization by the Arp2/3 complex drives

the formation of protrusions termed lamellipodia, which make new contacts with the

ECM through integrin receptors (Parsons et al., 2010; Svitkina and Borisy, 1999). These

focal complexes mature into focal adhesions, which is induced under the tension and

contractile forces of a network of actin stress fibers. Radial actin stress fibers emanate

from growing focal adhesions and link up with the contractile transverse arcs in the

leading lamella, which transmits forces to focal adhesions to promote forward

translocation (Gardel et al., 2010). Assembly of this dynamic network involves proteins

that nucleate, bundle and cross-link actin filaments (Choi et al., 2008). Force

transduction is regulated by the myosin II regulatory light chain, which modulates the

ATPase activity and step rate of myosin II motor in response to ECM stiffness (Davis et

al., 2002; Schwartz, 2010). Thus, the force-generating mechanisms that drive cell

motility are in a direct crosstalk with the extracellular microenvironment. In contrast, bleb

and lobopodial-based cell migration rely upon the assembly of contractile actomyosin

fibers at the cell cortex, which induces the formation of membrane protrusions that help

48 tumor cells navigate through gaps in the ECM (Lammermann and Sixt, 2009; Petrie and

Yamada, 2012) (Figure 4.1). The conversion to amoeboid and lobopodial-based cell

motility correlates with the expression of actin-binding proteins and ECM-degrading

enzymes and ECM topography (Petrie and Yamada, 2012), but the mechanism by

which tumor cells interconvert between these motility modes is not fully known.

Importantly, invasive tumor cells frequently show elevated expression of actin-

binding proteins (Wang et al., 2004), which control the assembly and organization of the

actin cytoskeleton during cell migration (Parsons et al., 2010). Septins have been

reported previously to promote cell migration, but the mechanism has remained elusive.

In metastatic cancer cell lines, SEPT9 expression is elevated and essential for the

formation of actin-rich pseudopods and cell migration (Shankar et al., 2010). In

agreement with this study, overexpression of five different SEPT9 isoforms in the non-

invasive MCF-7 breast cancer cells promotes cell migration (Connolly et al., 2011b).

Thus, septins promote tumor cell migration, but how septins influence the organization of

the actin cytoskeleton at the molecular level during cell motility has been unclear.

Septins have been linked to the assembly of branched actin networks in epithelia

and branched actin patches and neurons. SEPT1 knockdown in squamous cell

carcinoma cells inhibits cell spreading, which is dependent upon the formation of the

lamellipodia by the Arp2/3 complex (Mizutani et al., 2013). Interestingly, SEPT6

overexpression promotes the recruitment of cortactin, an activator of the Arp2/3

complex, to actin patches in neurons and renal epithelial lamellipodia (Hu et al., 2012).

Experiments in vitro with purified actin filaments and the Arp2/3 complex show that

recombinant SEPT6 localizes to actin branch points (Hu et al., 2012), which correlates

with Arp2/3 localization. Despite these findings, how septins interact with and regulate

actin filament assembly remained unknown.

49

In the first part of this thesis, I present a novel role for septins in promoting the

mesenchymal-like motility of renal epithelia undergoing EMT by directly crosslinking

actin filaments and regulating focal adhesion stability. Septins localizes to transverse

actin arcs and their orthogonal junctions with radial stress fibers, which are anchored to

focal adhesions. By disrupting the actin stress fiber network, septin depletion reduces

focal adhesion stability and cell migration. I identified further that SEPT9 expression

specifically increases during EMT, suggesting that SEPT9 may be part of the EMT

program that promotes the mesenchymal-like motility of epithelia. Indeed, SEPT9

overexpression promotes the migration of renal carcinoma cells and the collective

migration of renal epithelia undergoing EMT in 3D collagen matrix. Conversely, SEPT9

depletion in single cells migrating in a 3D matrix inhibits motility and induces a transition

to a more rounded and protrusive shape, indicating that SEPT9 expression promotes

specifically the mesenchymal-like motility mode.

Because stress fiber assembly during mesenchymal motility is regulated by

plethora of actin-binding proteins, whose spatial and temporal recruitment is tightly

regulated, I investigated further the interaction between SEPT9 and actin filaments at the

molecular level. SEPT9 binds to three specific sites on the actin filament, two of which

are shared by other actin-binding proteins. Strikingly, SEPT9 interferes with the binding

of the myosin motor domain in its low-affinity state and the actin-severing protein cofilin,

which could stabilize and protect filaments from depolymerizing forces. Thus, I

hypothesize that SEPT9 protects actin filaments by regulating spatially the recruitment of

actin-binding proteins during filament assembly. In agreement with this hypothesis, I

investigated the assembly and stability of nascent actin stress fibers in live renal

epithelia. Indeed, septin depletion results in unstable radial actin stress fibers that

disassemble at a faster rate.

50

At the molecular level, septins operate collectively as hetero-polymers and

filaments. In renal epithelia, SEPT9 colocalizes and immunopreciptates with SEPT2,

SEPT6 and SEPT7, indicating that they co-assemble into complexes and filaments.

While SEPT6 and SEPT9 bind directly to actin, SEPT2 does not interact with actin

filaments, but it binds directly to myosin II and is posited to control the phosphorylation of

its regulatory light chain and myosin contraction (Joo et al., 2007). These differential

interactions may regulate spatially the binding and activation of myosin along the actin

filament and contraction of actomyosin stress fibers during cell motility. In the absence of

septins, myosin motor proteins may bind and contract uncrosslinked actin filaments,

which can induce filament buckling and disassembly. Conversely, actin filaments lacking

septins may contain more available binding sites for the actin-severing protein cofilin,

which is enriched in the leading cell edge. Thus, overexpression of septins in cancer

may reinforce the mesenchymal-like motility mode by controlling the interaction of actin-

binding proteins with actin filaments during stress fiber assembly and contraction.

Septins in membrane traffic and metabolism

Septins also associate with the microtubule cytoskeleton and cell membranes,

regulating the intracellular trafficking of membrane cargo (Spiliotis and Gladfelter, 2012).

Importantly, membrane traffic is an emerging regulator of tumor cell migration, growth

and metabolism (Jones et al., 2006). Intracellular vesicles are derived from organelles

and the plasma membrane via endocytosis. Through the recruitment of adaptor and

motor proteins, vesicles move along actin filaments and microtubules to distinct

subcellular compartments (Fu and Holzbaur, 2014). Intracellular vesicle dynamics are

controlled by the large family of Rab GTPases, which can recruit motor proteins (e.g.

dynein) and promote vesicle docking, tethering and fusion (Stenmark, 2009). Moreover,

51 intracellular vesicles can serve as signaling hubs that activate pathways to promote cell

motility and growth. Rab-dependent recycling of integrins to focal adhesions (Caswell et

al., 2008) and active members of the Rho family of GTPases (Palamidessi et al., 2008),

which regulate actin polymerization at the leading cell edge, promotes tumor cell motility

and endocytosis. Mutations present in human cancers that render Rab GTPases (e.g.

Rab35) constitutively active were identified recently to stimulate tumor cell proliferation

via aberrant trafficking of growth factor receptors to lysosomes and activation of the

PI3K/AKT/mTOR signaling pathway (Wheeler et al., 2015).

Septins regulate the intracellular delivery of golgi-derived to the plasma

membrane domains in renal epithelia, which is important for epithelial polarity and

morphogenesis (Spiliotis, 2008). Septin knockdown or inhibition with a function-blocking

antibody impedes the transport of plasma membrane proteins through the exocytic

pathways (Spiliotis, 2008). Septins colocalize with polyglutamylated microtubules and

regulate the organization and dynamics of the microtubule network (Bowen et al., 2011;

Spiliotis, 2008).

In contrast, little is known about the function of septins in endocytic pathways.

Recent evidence shows septins regulate epidermal growth factor receptor (EGFR)

surface levels and degradation (Diesenberg et al., 2015), which occurs through

endocytic recycling, and promote the formation of multivesciular bodies (Traikov et al.,

2014). Importantly, altered endocytosis is commonly observed in tumor cells.

Upregulation of the endocytic mechanisms that internalize adhesion proteins promote

the breakdown of cell-cell junctions (Yang et al., 2006), and increased macropinocytosis,

a form of clathrin-independent endocytosis, promotes tumor cell growth and possibly

migration (Commisso et al., 2013).

52

Macropinosomes are derived from plasma membrane ruffles, which fold back on

themselves forming large vesicles that can capture membrane proteins, extracellular

fluids and solutes (Lim and Gleeson, 2011). Following ruffle closure, macropinosomes

undergo a maturation process that involves changes in lipid composition, the recruitment

of Rab GTPases and fusion with late endosomes, which can recycle proteins back to the

plasma membrane, and lysosomes, which degrade proteins for catabolic pathways

(Bright et al., 2005; Swanson, 2008) (Figure 4.2). In motile fibroblasts, growth factor

stimulation induces the formation of macropinosomes that recycle integrins back to

nascent focal adhesions in the leading cell edge; and inhibition of macropinocytosis or

Rab-dependent integrin trafficking attenuates fibroblast migration (Gu et al., 2011). Thus,

macropinocytic trafficking of cell adhesion molecules may be necessary for tumor cell

invasion.

In addition, recent evidence also supports a role for macropinocytosis in tumor

cell growth via carbon metabolism (Commisso et al., 2013). Uncontrolled tumor cell

proliferation requires increased energy metabolism in order to sustain growth. Tumor

cells frequently bias energy production towards glycolysis and glutamine metabolism, or

glutaminolysis, which have emerged as hallmarks of tumor cell metabolism

(Deberardinis et al., 2008). Notably, the glycolytic and glutamine metabolic pathways are

frequently linked to cellular transformation by oncogenes, such as Ras GTPases (White,

2013). In addition, pancreatic tumor cells expressing a constitutively-active mutation in

the KRas oncogene show increased uptake of amino acids through macropinocytosis

and the delivery of glutamine to the central metabolic pathway (Commisso et al., 2013).

Strikingly, inhibition of macropinocytosis and subsequent glutamine metabolism reduces

tumor cell proliferation in vitro and tumor growth in vivo (Commisso et al., 2013).

53

In the last part of this thesis, I discovered a population of membrane-associated

septins that localize to macropinosomes and their contact sites with endosomes by

interacting directly with PI(3,5)P2, a low-abundant lipid that concentrates on late

endosome and lysosomes. Septin knockdown results in clusters of macropinosomes and

abrogates macropinosome fusion with lysosomes. Conversely, septin overexpression

enhances the trafficking of macropinocytic cargo to the lysosomes. Using an in vitro

vesicle fusion assay, I found that septins promote membrane fusion independently of the

actin and microtubule cytoskeleton.

Membrane-associated septins control the organization of plasma membrane

lipids and proteins (Spiliotis and Gladfelter, 2012). In addition, septins have been

reported to interact directly with exocytic SNAREs, whose clustering into membrane

domains is required for fusion. (Aoyagi et al., 2005). Based on this knowledge, septins

may promote vesicle fusion by controlling SNARE organization in membrane domains or

the dynamic assembly and disassembly of SNARE complexes during macropinosome

fusion with lysosomes. Through these potential mechanisms, septin overexpression may

promote tumor cell proliferation by promoting the trafficking of macropinocytic cargo to

the lysosome, which can supply molecular intermediates for the central metabolic

pathway.

In a similar manner, autophagy is a process by which stressed or starved cells

capture cellular material (e.g. organelles, proteins) into autophagosomes, which are

targeted to the lysosome for degradation (Kaur and Debnath, 2015; Levine and Klionsky,

2004). Autophagy has been linked to prolonged tumor cell survival, but also defects in

the autophagic machinery are associated with tumorigenesis (Mathew et al., 2007).

During microbial infection, septins target intracellular bacteria (e.g. Shigella) to

autophagosomes, which restricts their replication and reduces infection in vitro and in

54 vivo (Mostowy et al., 2010; Mostowy et al., 2013). Septins are posited to interact directly

with the autophagy machinery, which also targets autophagosomes to lysosomes by

interacting with lysosomal SNAREs (Liu et al., 2015; Mostowy et al., 2010). How septins

function in autophagy is not fully understood, and whether septin overexpression may

support autophagy-mediated cell survival in tumor cells is unknown. However, I

speculate that septins may have a wider role in targeting intracellular vesicles to

lysosomes.

55

Fig 4.1 Modes of single tumor cell migration. (A) Mesenchymal motility is characterized by the formation of focal adhesions and an

actin stress-fiber network containing radial stress fibers (SFs) and transverse arc

SFs that link up to promote focal adhesion growth.

(B) Amoeboid motility is characterized by the assembly of actomyosin fibers at the cell

cortex and the formation of protrusive blebs.

(C) Lobopodial motility is characterized by the assembly of an actomyosin network at the

cell cortex that connects with a network of intermediate filaments near the nucleus.

Through this cytoskeletal connectivity, the nucleus pressurizes the cell front in a

piston-like mechanism that promotes the extension of blunt membrane protrusions

termed lobopodia.

56

Figure 4.2 Macropinosome formation, maturation and traffic to endolysosomes. Plasma membrane ruffles fold back upon themselves to engulf soluble cargo (red) into

macropinosomes. Macropinosomes undergo a maturation process that is controlled by

changes in phosphoinositide lipid composition and the recruitment of Rab GTPases,

which collectively regulate macropinosome retrograde movement and fusion with late

endosomes and lysosomes.

57

CHAPTER 5. Septins promote stress fiber-mediated maturation of focal

adhesions and renal epithelial motility.

Abstract

Organogenesis and tumor metastasis involve the transformation of epithelia to

highly motile mesenchymal-like cells. Septins are filamentous G proteins, which are

over-expressed in metastatic carcinomas, but their functions in epithelial motility are

unknown. Here, we show that a novel network of septin filaments underlies the

organization of the transverse arc and radial (dorsal) stress fibers at the leading lamella

of migrating renal epithelia. Surprisingly, septin depletion resulted in smaller and more

transient and peripheral focal adhesions. This phenotype was accompanied by a highly

disorganized lamellar actin network and rescued by the actin bundling protein α-actinin-

1. We show that pre-assembled actin filaments are crosslinked directly by SEPT9,

whose expression is increased after induction of renal epithelial motility with the

hepatocyte growth factor. Significantly, SEPT9 over-expression enhanced renal cell

migration in 2D and 3D matrices, while SEPT9 knock-down decreased migration. These

results suggest that septins promote epithelial motility by reinforcing the cross-linking of

lamellar stress fibers and the stability of nascent focal adhesions.

Introduction

Embryonic patterning and organogenesis depend on the morphogenetic

movements of epithelial cells, which transition to migratory mesenchymal-like cells

(Thiery et al., 2009). In the developing kidney, extensive epithelial movements

accompany the formation, branching and tubulogenesis of the ureteric bud (Dressler,

2002; Zegers et al., 2003), and epithelial-to-mesenchymal transition (EMT) underlies

58 renal fibrosis and the metastasis of renal cell carcinomas (RCCs) (De Craene and Berx,

2013; Liu, 2010). EMT is characterized by disruption of cell-cell adhesions, cell

scattering and the enhancement of invasive cell motility with a front-rear polarity

(Guarino et al., 2007; Lim and Thiery, 2012).

Mesenchymal motility is driven by the actomyosin cytoskeleton and involves

adhesion to the ECM and its degradation (Bravo-Cordero et al., 2012; Friedl and Wolf,

2010). Nascent adhesions are stabilized and grow to focal adhesions (FAs) under the

tension exerted by a network of actin stress fibers (Geiger and Yamada, 2011; Schwartz,

2010). At the leading lamellae of migrating cells, forces are transduced to adhesions by

the radial (dorsal) stress fibers (RSFs), which extend dorsally from FAs and connect with

the transverse arc (TA), a contractile network of curved actin filaments (Burridge and

Wittchen, 2013; Gardel et al., 2010; Parsons et al., 2010). Actin nucleating and cross-

linking proteins and myosin II, whose activity is regulated by the phosphorylation of its

regulatory light chain (RLC), mediate the assembly, organization and contraction of actin

stress fibers (Burridge and Wittchen, 2013; Parsons et al., 2010). Actomyosin

organization and contractility are regulated by the small GTPases of the Ras

superfamily, which affect cell motility (Parri and Chiarugi, 2010; Raftopoulou and Hall,

2004).

Septins are evolutionarily and structurally related to the Ras GTPases (Leipe et

al., 2002), but unlike the monomeric small GTPases, septins form filamentous polymers

that consist of non-polar hetero-octameric complexes (Mostowy and Cossart, 2012;

Weirich et al., 2008) . Septins interact with the actomyosin cytoskeleton (Joo et al., 2007;

Kinoshita, 2002; Mavrakis et al., 2014) and are abnormally over-expressed in renal cell

carcinomas, but their precise function in stress fiber organization and mesenchymal-like

59 motility is unknown. Here, we show that in the leading edge of motile renal epithelia, a

novel network of septin filaments promotes cell motility by reinforcing the organization of

the lamellar stress fiber network and the stability of nascent FAs.

Materials and methods

Tissue culture and transfections.

MDCK II/G cells and the stable MDCK-SEPT2-YFP (Spiliotis et al., 2005), MDCK-

paxillin-GFP (Yamada and Nelson, 2007), MDCK-SEPT9-mCherry, MDCK-ABD-

ABP140-mCherry cell lines were cultured in low glucose DME with 1g/liter NaHCO3,

10% FBS (Cell Generation), and 1% PSK (penicillin, streptomycin, kanamycin) at 5%

CO2. The stable MDCK-SEPT9-mCherry and MDCK-ABP140-mCherry cells were

generated by standard selection with G418 sulfate (0.8 mg/mL; Corning) and stable

transfectants were cloned in 96-well plates by serial dilutions and levels of expression

were analyzed by western blotting. The renal cell carcinoma 786-O cell line was

purchased from ATCC and maintained in RPMI with 1 g/liter NaHCO3, 10% FBS, and

1% PSK. Coverslips were acid-washed with 1 M HCl for 4h at 50ºC and coated with 10-

20 µg/mL fibronectin (Invitrogen) for 2h at 37ºC or rat tail collagen. Cells were

transfected with 1.5 µg of plasmid or co-transfected with 1.5 µg plasmid and 1µg

shRNAs with Lipofectamine 2000 (Invitrogen). Prior to fixation or live-imaging, MDCK

cells were stimulated with 20 ng/mL recombinant HGF (Sigma) in DME supplemented

with 0.5% FBS for at least six hours. Live imaging studies were performed in phenol red-

free media containing 20 mM HEPES.

To generate polarized cysts, MDCK cells and cells that express stably SEPT9-

mCherry at low and high levels were cultured in collagen gels composed of 2mg/mL type

I collagen (Advanced Biomatrix) on transwell filters (0.4 µm pore; BD Biosciences) for

ten days. Polarized cysts were stimulated with 30 ng/mL HGF for three days (Fig. 4 C-E)

60 and with or without 50 µm FCF (Fig. 4E). To image and quantify single cell migration in

3D (Fig. 4 H and I), MDCK cells were transfected with control or SEPT9-targeting

shRNAs for 24h on tissue culture plates. Cells were dissociated by trypsinization and

cultured in 2 mg/mL type I collagen gels on 35 mm glass-bottom dishes (live imaging of

MDCK-ABP140-ABD-mCherry; Fig. 4 H) or transwell filters (morphometric analysis of

fixed MDCKs; Fig. 4I) and treated with 40 ng/mL HGF, 16 h prior to imaging.

Plasmids and shRNAs.

Canine SEPT2 was targeted with control (5’-TGA GTTCACACTAATGGTGGTCG-3’),

SEPT2 shRNA #1 (5’-CCTTAGACGTCGCAT TCATGAAA-3’) and SEPT2 shRNA #2 (5’-

GCAACTACAAGCCCGAAGAATAA-3’) (Bowen et al., 2011). To target human SEPT2,

the SEPT2 shRNA 5’-AAGGTGAATATTGTGCCTGTC-3’ was subcloned into the GFP-

expressing pG-SUPER vector. The SEPT9 shRNA construct was made by inserting the

SEPT9 shRNA 5’- GACCGGCTGGTGAACGAG AAGTT-3’ into mCherry-expressing

pSuper vector. The shRNA constructs against canine and human SEPT9 were made by

subcloning the shRNAs 5’-GACCGGCTGGTGAACGAGAAGTT-3’ and 5’-

GACCATCGAGATCAAGTCC-3’ into the mCherry-expressing pSuper and GFP-

expressing pGFP-V-RS vectors, respectively. ABP140-ABD-mCherry was generated by

annealing the primers 5’-ATCCTCCTCCTTG

CTGATGCTTCGAACTTCTTGATCAGGTC GGCCACGC CCATGGTGGCG-3’ (forward)

and 5’-AATTCGCCACCATGGGCGTGGCCGACCTGATCAAGAAGTCGAGAG

CATCAGCAAGGAGG-3’ (reverse), which correspond to the first 17 amino acids

(Lifeact) of ABP140, and inserting the sequence into the N1-mCherry vector (Invitrogen)

using the NcoI restriction site. SEPT9_mCherry was constructed by PCR amplification of

SEPT9_i1 from pET28a-SEPT9_i1 (Bai et al, 2013) using the primers 5’-

61 CGTAAGCTTGCATGAAGAAGT-3’ and 5’-GTAGTCGACCTACATCTCTGGGGC-3’ and

the product was ligated into the HindIII and SalI sites of the pmCherry-C1 vector. Mouse

SEPT2 was inserted into N1-YFP (Spiliotis et al, 2005). Avian paxillin was inserted into

C1-EGFP (West et al, 2001). Mouse myosin IIA-GFP, human α-actinin-1-GFP and

human myosin II RLC-DD-GFP (pEGFP-MRLC1 T18D, S19D) were purchased from

Addgene. The actin-binding domain of α-actinin-1 (amino acids 30-253) was deleted

from α-actinin-1-GFP (Roca-Cusachs et al., 2012). The myosin IIA-N93K-GFP mutant

was generated by site-directed mutagenesis using the primers 5’-

CTCACGTGCCTCAAGGAAGCTTCGGTG-3’ (forward) and 5’-CACCGAAGCTTCCTTG

AGGCACGTGAG-3’ (reverse).

Protein expression and purification.

SEPT2/6/7 was expressed by co-transforming E. coli BL21(DE3) cells with a bicistronic,

kanamycin-resistant pET-28a(+) plasmid that expresses SEPT6 and His-SEPT7 in

tandem, and an ampicillin resistant pET-15b(+) plasmid that expresses untagged SEPT2

(Bai et al., 2013). His-SEPT9_1 was expressed by transforming E. coli BL21(DE3) cells

with pET28a-SEPT9_i1 (Bai et al, 2013). Bacterial cultures of at least OD600 of 0.6 were

induced with 0.5 mM IPTG for 16h at 18C, centrifuged at 5,000 rpm for 5 m at 4ºC, and

resuspended in buffer containing 1% Triton X-100, 50 mM Tris, pH 8.0, 150 mM NaCl,

10% glycerol and 10 mM imidazole. Bacteria were lysed with a French pressure cell at

1,280 psi. Cell lysates were cleared using centrifugation (14,000 x g) for 30min at 4ºC

and the supernatants were passed through Ni-NTA beads by gravity flow. The columns

were washed with 10mL washing buffer (50mM Tris, pH 8.0, 300mM NaCl, 10%

glycerol, 25 mM imidazole) and proteins were eluted with elution buffer (50 mM Tris,

pH8.0, 300 mM NaCl, 10% glycerol, and 250 mM imidazole). Purification of SEP2/6/7

62 was verified by size exclusion chromatography using an AKTA FPLC system (GE

Healthcare). Purified proteins were dialyzed overnight in buffer containing 50mM Tris,

pH 8.0, 150 mM NaCl, and 10 mM imidazole.

Immunoprecipitations and immunoblots.

For immunoprecipitations, equal number of MDCK and SEPT2-YFP MDCK cells were

lysed with buffer containing 50 mM HEPES pH 7.4, 100 mM NaCl, 0.5% Triton X-100, 1

mM PMSF, and protease inhibitors (Millipore) and incubated with anti-GFP antibody

(3E6; Invitrogen) overnight at 4ºC. Protein A beads (Thermo Scientific) were incubated

for 4 h at 4ºC and were stripped with SDS buffer at 95ºC for 10min. Equal volumes were

resolved on a 10% SDS-PAGE gel and transferred to a nitrocellulose membrane using

100 V for 1 h at 4ºC. Membranes were blocked with buffer containing 5% milk and 1%

BSA for 1h at 25ºC. Primary antibodies targeting rabbit anti-SEPT2 (1:5000, Sigma),

rabbit anti-SEPT6 (1:5000, Santa Cruz Biotechnology, Inc.), rabbit anti-SEPT7 (1:8000,

ImmunoBiological Laboratories, Inc.) and rabbit anti-SEPT9 (1:2500, ITG Labs), and

anti- rabbit or mouse secondary antibodies (LiCor) conjugated to infrared dyes were

diluted in buffer containing 2% BSA, 0.1% Tween-20, and 0.025% sodium azide and

sequentially incubated on each membrane prior to scanning with an Odyssey Li-Cor

imaging system.

To test for changes in SEPT9 expression following HGF treatment, MDCK cells

were cultured on collagen-coated plates and stimulated with HGF for the indicated times.

Cell lysates of equal protein concentration were run on a 10% SDS-PAGE gel,

transferred to a nitrocellulose membrane and probed using primary antibodies targeting

rabbit anti-SEPT9 (PTG Labs), mouse anti-actin (1:8000, Sigma) and mouse anti-

GAPDH (1:8000, Abcam). The western blot shown in Fig. 4 A is representative of three

63 different experiments, which were reproduced with no limitations. To quantify SEPT9

overexpression in MDCK cells, untransfected cells and cells that express stably SEPT9-

mCherry at high levels were lysed, processed and blotted as described above (Fig. S2

D).

Immunofluorescence microscopy and quantifications.

Cells were simultaneously fixed and extracted with 4% PFA in warm PBS buffer

containing 0.1% Triton X-100 for 20min, or fixed with 4% PFA in cytoskeleton buffer (10

mM MES pH 6.1, 0.32 M sucrose, 138 mM KCl, 3 mM MgCl2, 2 mM EGTA) for 20 min

and extracted with 0.5% Triton X-100 for 10min (Fig. 3 D). Excess PFA was quenched

with 0.25% ammonium chloride. Prior to antibody incubation, cells were blocked with

PBS containing 2% BSA for 20 min. Primary antibodies to rabbit anti-SEPT2 (N5N; gift

from M. Kinoshita, Nagoya University, Nagoya, Japan), rabbit anti-SEPT6 (Tada et al.,

2007), rabbit anti-SEPT7 (IBL, Inc.), SEPT9 (ITG Labs), mouse anti-paxillin (BD, 349),

rabbit anti-pY118-paxillin (Millipore), rabbit anti-myosin-IIA (Sigma), mouse anti-pSer19-

MLC (Cell Signaling Technologies), and secondary donkey DyLight 488-, 594-, 647-

conjugated F(ab’)2 to mouse or rabbit IgG (Jackson ImmunoResearch Laboratories,

Inc.) were diluted in PBS with 2% BSA and 0.1% Triton X-100. F-actin was visualized

using phalloidin conjugated to Alexa-488 or -647 (Life Technologies). Coverslips were

mounted on slides using Vectashield (Vector Laboratories) or FluorSave (EMD) and

imaged using a confocal laser-scanning microscope (FluoView 1000; Olympus) with a

Plan Apocrhomat 60x/1.42NA objective. Optical sections (0.2 µm thick) were exported to

the Slidebook 5.0 (Intelligent Imaging Innovations), ImageJ or Volocity (Perkin Elmer)

softwares for processing and quantification. Kymographs were generated in Slidebook

by drawing a six pixel wide line down the center of a growing FA marked by paxillin or

64 ABP140-ABD. Super-resolution structured illumination microscopy was performed using

the DeltaVision OMX V4 (GE Healthcare) microscope with a 60x/1.42NA objective and

immersion oil with a refractive index of 1.514. A z-step size of 0.125 µm was used and

images were acquired with sCMOS pco.edge cameras (PCO) and reconstructed in the

softWoRx software.

FA sizes were quantified using masks of paxillin fluorescence intensity

segmentation; the area of individual masks were exported to excel and quantified. FA

distance from the cell edge was measured by converting the phalloidin-stained cell into a

binary image, which was subsequently inverted and transformed into a Euclidean

distance map (Schober et al., 2007). FAs were overlaid on the map and the distance

from the FA center to the cell edge was measured. To assess relative levels of pTyr118

paxillin to total paxillin, confocal images were obtained using identical laser settings

between control and septin-depleted samples. Individual FAs were masked using

intensity segmentation and the ratios were derived from the sum intensities.

Colocalization of septins with actin in the leading cell edge was quantified in

background subtracted images using the Mander’s Coefficient analysis in Slidebook.

Fluorescence intensity of actin as a function of the distance from the cell edge was

quantified from confocal images, which were acquired in during one sitting at identical

laser intensities. Protrusions at the leading cell edge were sampled 2-3 times using a 10

µm x 1 µm line scan drawn perpendicular from the cortex and were averaged and

plotted using custom software written in MATLAB (MathWorks). The number of RSFs

per micron was calculated by drawing a line parallel to the cell edge and counting the

number of radial fibers that intersected with the line. Fluorescence intensity ratios of

pSer19-myosin II light chain to myosin IIA heavy chain (Fig. S3) was quantified by

65 masking 5 µm segments of the TAs parallel to the leading cell edge. In each cell, the

sum intensities of at least six different segments were analyzed.

To visualize 3D cyst structure (Fig. 4 C), gels were briefly treated with

collagenase (Sigma) in PBS at 37ºC and fixed with 4% PFA in PBS with 0.5% Triton X-

100. Gels were incubated with phalloidin for 1 h under gentle agitation and mounted in

Vectashield. Optical sections (1.25 µm thick) were stacked as mass projections. The

number of extensions per cyst and the number of cells per extension were quantified

and plotted. Extensions were defined as protrusions from the main cyst body consisting

of three cells or more. To quantify single cell morphology in 3D (Fig. 4 I), gels were fixed,

stained and mounted using the same method but without the initial collagenase

treatment. The long and short axes were measured in Slidebook to derive the axial ratio

of each cell. To measure the velocity of cell movement in a 3D matrix (Fig. 4 H), time-

lapse optical sections were imported into the Volocity software, which identified, tracked

and quantified the movement of the cell centroid.

Time-lapse imaging.

All live imaging experiments were performed at 37ºC in DME lacking Phenol red

supplemented with 0.5% FBS, 1% PSK, and 20mM HEPES. Widefield microscopy was

performed with an inverted microscope (IX-81; Olympus) equipped with a motorized

stage (ProScanII; Prior), a camera (OrcaR2; Hamamatsu Photonics), a custom built

stage-top chamber and temperature controller (Air-Therm ATX; World Precision

Instruments) and the Slidebook 5.0 software. Fluorescent images were captured every

20s with a Plan Apocrhomat 60x/1.40 NA objective (Olympus). Spinning disk confocal

microscopy was performed at 37ºC using an inverted microscope (IX-71; Olympus)

equipped with a camera (ImagEM; Hamamatsu Photonics), an airstream incubator

66 (LiveCell; Pathology Devices), scan head (CSU10; Yokogawa), and the MetaMorph

software (Molecular Devices). Fluorescent images were captured every 20s with a 60x

1.49 NA objective. TIRF microscopy was performed at 37ºC with the DeltaVision OMX

V4 inverted microscope (GE Healthcare) equipped with 60x/1.49NA TIRF objective

(Olympus), motorized stage, sCMOS pco.edge cameras (PCO), stage-top incubator with

temperature controller and the softWoRx software. 3D cell migration experiments were

performed using a laser scanning confocal microscope (LSM700; Carl Zeiss) equipped

with a stage-top incubator and temperature controller (XL modules; Carl Zeiss), and the

Zen software (Carl Zeiss). One micron optical sections were acquired every two minutes

with a 40x C-Apochromat/1.2 NA water-objective (Carl Zeiss). Supplemental videos

were rendered using the Photoshop software (Adobe).

Quantification of FA dynamics.

FA dynamics were obtained using the stable MDCK-paxillin-GFP cell line and TIRF

microscopy; background subtraction and photobleach correction algorithms in the

Slidebook software were applied to normalize fluorescence intensities. Individual FA

lifetimes were quantified from the length of each adhesion signal in three cells using the

open-source Focal Adhesion Analysis Server (Berginski and Gomez, 2013). FA

assembly and disassembly rates were quantified as previously described (Webb et al.,

2004). In brief, individual FAs were masked in Slidebook and a semilogrithmic plot of

paxillin intensity as a function of time was generated. The rate constants were derived

from the slopes of 45 adhesions from the leading edge (three cells per condition).

Low speed co-sedimentation assay.

Human platelet actin (Cytoskeleton, Inc.) in G-buffer (5mM Tris-HCl, 0.2mM CaCl2) was

polymerized using F-buffer (20 mM HEPES, 100 mM KCl, 1 mM MgCl2) supplemented

67 with 0.5 mM ATP and 4 mM DTT for 1h at 25ºC to generate F-actin. 2 µM recombinant

SEPT2/6/7 and 4 µM recombinant his-tagged SEPT9 were incubated with 2 µM F-actin

for 1h at 25ºC and centrifuged atop a glycerol cushion (20 mM HEPES, 100 mM NaCl, 1

mM MgCl2, 10% glycerol) at 8,000 x g for 20min at 25ºC. Equal volumes of the

supernatant and pellet fractions were resolved using a 7.5% SDS-PAGE gel and stained

with Coomassie brilliant blue.

Electron Microscopy.

Skeletal striated muscle G-actin (2.5 μM) was polymerized in 10 mM HEPES buffer (pH

7.4), 100 mM KCl, 1 mM MgCl2, 0.5 mM DTT, and 0.5 mM ATP for 2 h. Actin filaments

were incubated with 14 μM His-SEPT6 or His-SEPT9 for 15-30 min at 4°C. To visualize

complexes of F-actin with septins, aliquots (10 µl) were applied to glow-discharged

carbon-coated grids (Electron Microscopy Sciences) and stained with 2% uranyl acetate.

The grids were examined in a JEOL 1200EXII transmission electron microscope (TEM)

(JEOL Inc, Peabody, MA) under regular-dose conditions at an accelerating voltage of 70

keV. The images shown in Fig. 3 F and G are representative of similar results obtained

from three independent experiments.

Transmigration Assays.

786-O cells were transfected with plasmids encoding for the indicated constructs for 48h,

trypsinized and plated on collagen-coated transwell filters (8 µm pore) for 12h. Filters

were fixed with 4% PFA in PBS and stained for nuclei with DAPI. Apical and basolateral

sections were imaged at 20x magnification using confocal microscopy. The number of

transfected cells in each section were quantified and plotted in excel.

Statistical Analyses

Data sets were analyzed using R and the Excel software. Normal distribution and equal

variance were assessed using the Shapiro and Levene tests, respectively. Data sets

68 with normal distribution and variance were analyzed using an unpaired student’s t-test.

Data sets that did not exhibit normal distributions or variance were further analyzed

using the Kruskal-Wallis non-parametric ANOVA. All values represent the mean +/-

SEM.

Results and Discussion

A network of septin filaments interfaces with RSF and TA stress fibers in the

leading lamella of renal epithelia.

To examine septin functions in the motility of renal epithelia, we used the MDCK

model of partial EMT, which allows the study of cell migration in 2D and 3D matrices

after stimulation with the hepatocyte growth factor (HGF) (Zegers et al., 2003).

Treatment of MDCKs with HGF disrupted cell-cell adhesions and induced scattering and

development of front-rear cell polarity as previously shown (de Rooij et al., 2005).

Resembling the stress fiber organization of migratory cells, the lamellae of fan-shaped

MDCK cells contained radial stress fibers that extended dorsally from FAs and

connected with transverse arc (TA) filaments (Fig. 1 A; arrows). Staining for endogenous

SEPT2, SEPT6, SEPT7 and SEPT9 revealed an extensive network of septin filaments

that localized at the interface of TA filaments and RSFs (Fig. 1 A, Fig. S1).

Colocalization and co-immunoprecipitation of SEPT2, SEPT6, SEPT7 and SEPT9 (Fig.

S2, A and B) indicated that these septin filaments consist of SEPT2/6/7/9 complexes,

which are the basic structural units of mammalian septin filaments (Kim et al., 2011;

Sellin et al., 2011b). Thus, we used SEPT2 as a marker for the lamellar network of

septin fibers, and SEPT2 and SEPT9 shRNAs were interchangeably applied to perturb

the localization and function of septins; knock-down of SEPT2 or SEPT9 affected the

69 expression and localization of both of these septins as well as SEPT6 and SEPT7 (Fig.

S2, C-D) as reported previously (Kim et al., 2011).

To determine the precise localization of septins with respect to the lamellar

network of stress fibers and FAs, we used confocal microscopy in combination with 3D

image reconstructions and super-resolution structured illumination microscopy (SIM).

Septins localized mainly on and between the curved actin bundles of the TA network and

on a subset of RSFs (Fig. 1 B-E; Video 1). Septins were present on the dorsal ends of

80% ± 2% RSFs (n = 72; 12 cells) that connected with the TA (Fig. 1, B and D). On the

quartile end of these RSFs, 1.9 ± 0.16 µm away from FAs (n = 58; 12 cells), septins

interdigitated with actin filaments as they flared from their FA-anchored bundle toward

the TA (Fig. 1, E; panel 1). Time-lapse microscopy showed that septins are recruited to

the distal ends of RSFs after the assembly of nascent FAs (Fig. 1 F and G). In live cells,

septins persisted on RSFs during their interactions with the TA, spanning segments of

both RSF and TA stress fibers (Fig. 1 H). Taken together, these data indicate that

septins localize at the interface of the contractile TA and RSFs, and thus, may be part of

the actomyosin machinery that drives the growth and turnover of nascent FAs in motile

renal epithelia.

Septins are required for the stabilization of nascent FAs and the organization of

the lamellar stress fiber network.

To test whether septins affect the formation and dynamics of FAs, HGF-treated

MDCK epithelia were depleted of SEPT2 or SEPT9. Septin knock-down with two

different SEPT2 shRNAs and a SEPT9 shRNA increased the number of FAs, which

were smaller and localized closer to the cell edge compared to control cells (Fig. 2 A-D).

Time-lapse imaging of paxillin-GFP showed that septin knock-down reduces the life time

70 of nascent FAs (Fig. 2E), which fail to grow significantly in size and do not move closer

to the cell body (Video 2). Analysis of the kinetics of paxillin-GFP fluorescence revealed

no differences in the rates of FA assembly and disassembly (Fig. 2 F), but septin

depletion abolished the stabilization phase of the FA life cycle (Fig. 2 G). Consistent with

lack of FA stabilization, which is characterized by dephosphorylation of paxillin’s Tyr118

(Zaidel-Bar et al., 2007), septin depletion increased the levels of pTyr118-paxillin relative

to total paxillin in MDCK and 786-0 RCC cells (Fig. 2 H-I).

Maturation of nascent FAs is mediated by stress fibers, which exert mechanical

forces and provide a template for the growth of cell-ECM adhesions (Burridge and

Wittchen, 2013; Gardel et al., 2010; Vicente-Manzanares et al., 2009). While myosin-

mediated tension is thought as the main trigger of FA maturation, recent work shows that

myosin II-dependent tension is necessary but not sufficient for FA maturation, which

requires actin-binding proteins that mediate the formation and maintenance of the

lamellar actin network (Choi et al., 2008; Oakes et al., 2012; Roca-Cusachs et al., 2013;

Stricker et al., 2013).

Next, we examined the organization and dynamics of the lamellar actin network

in septin-depleted cells. Strikingly, the lamellar actin meshwork was thin and devoid of

an organized TA (Fig. 2 J). F-actin was significantly reduced in the lamellar regions of

the cell (5-10 µm from the cell edge; Fig. 2 K), and the length and density of RSFs

decreased by ~50% (Fig. 2 L and M). In live SEPT2-depleted cells, RSFs were short-

lived, dissipating quickly after their formation, while in control cells the actin fluorescence

of RSFs persisted and increased over time (Fig. 2 N; Video 3). Taken together with the

localization of septins to the TA and distal ends of RSFs, these data suggest that septins

71 are required for the stabilization and maturation of nascent FAs by maintaining the

organization and possibly contractility of the lamellar stress fiber network.

Septins function as actin cross-linking proteins in FA maturation.

Because SEPT2 binds directly myosin II and this interaction is posited to facilitate the

activation of the myosin RLC (Joo et al., 2007), we sought to determine whether septins

affect FA maturation through their role in the organization of the lamellar actin network or

by a possible involvement in myosin-mediated tension. To distinguish between these

possibilities, we tested first whether septins affect the levels of activated myosin RLC on

the contractile TA, and second whether the actin bundling protein α-actinin-1 or the

constitutively active di-phosphomimetic myosin RLC-DD can rescue septin loss of

function. Analysis of the fluorescence intensity ratio of the phosphorylated pSer19

myosin RLC to total myosin IIA showed no difference between control and SEPT2-

depleted cells (Fig. S3 A and B). In agreement, RLC-DD did not rescue FA size and

number in SEPT2-depleted cells (Fig. 3 A-C). In contrast, α-actinin-1 restored the

number and size of FAs to near control levels (Fig. 3 A-C). This effect of α-actinin-1 was

also accompanied by a rescue of the lamellar actin network in septin-depleted cells,

which now contained a network of TA filaments and RSFs (Fig. 3 D). Notably, α-actinin-

1-∆ABD, which lacks its actin-binding domain, did not rescue FAs (Fig. 3 B and C), while

the actin-binding motor-dead myosin II-N93K did (Fig. S3 C-E). Thus, loss of septin

function in FA maturation is specifically rescued by the actin-binding and cross-linking

properties of α-actinin-1 and myosin IIA heavy chain.

72 Previous studies have reported that septin complexes comprising of SEPT2,

SEPT6 and SEPT7 do not bind actin filaments directly (Kinoshita et al., 2002), but new

work shows that copolymerization of actin with SEPT2/6/7 results in circular actin

filaments (Mavrakis et al., 2014). Because in the lamellar regions of motile MDCK

epithelia, septins interact with pre-assembled RSFs and TA filaments, we examined

whether septins have the ability to cross-link pre-polymerized actin filaments using low

speed actin sedimentation assays and negative stain EM. We found that SEPT9 directly

cross-links pre-polymerized actin filaments into bundles, while SEPT2/6/7 does not (Fig.

3 E-G). These results suggest that septins promote the maturation of nascent FAs by

cross-linking the lamellar stress fibers of motile epithelia.

SEPT9 is up-regulated in renal epithelia undergoing EMT and enhances renal cell

migration.

Given that septins are over-expressed in renal cell carcinomas and other cancers

(Connolly et al., 2011a; Craven et al., 2006a; Dolat et al., 2014a), we asked whether

SEPT9 is upregulated during renal EMT. Treatment of MDCK cells with HGF increased

the levels of SEPT9 relative to actin, while actin levels did not change (Fig. 4 A and B).

To test if increased SEPT9 expression promotes cell motility, we established stable

MDCK cell lines expressing SEPT9-mCherry at low endogenous-like and high levels of

expression, and cultured these cells in a 3D ECM, which enables the morphogenesis of

renal cysts. Treatment of the 3D MDCK cysts with HGF causes tubulogenesis, a process

that mimics the collective and invasive migration of metastatic tumors (Debnath and

Brugge, 2005). Over-expression of SEPT9 doubled the number of multicellular

extensions that protruded from the main cyst and the extensions were longer and

contained more cells (Fig. 4 C and D). In contrast, treatment of cysts with

73 forchlorfenuron (FCF), a septin-targeting compound (Hu et al., 2008), reduced

significantly the number of HGF-induced extensions (Fig. 4 E). Moreover, dissociated

single MDCK cells migrated slower in 3D and were less elongated and fibroblast-like

after SEPT9 depletion (Fig. 4 H and I; Video 4). To corroborate the effects of SEPT9

over-expression on renal cell motility, we performed a transmigration assay using the

786-0 renal carcinoma cells. SEPT9 over-expression increased the migration of 786-0

cells along a serum gradient, while SEPT9 knock-down decreased migration (Fig. 4 F

and G). These results, therefore, indicate that SEPT9 promotes the mesenchymal-like

migration of renal epithelia.

Previous studies have implicated septins in the amoeboid motility of T-cells and

neuronal migration (Shinoda et al., 2010; Tooley et al., 2009), which are mechanistically

different from the stress fiber-driven mode of mesenchymal migration. Septins have

been reported to colocalize with stress fibers in fibroblasts (Kinoshita et al., 1997;

Kremer et al., 2007; Schmidt and Nichols, 2004b), but this interaction has been

considered to be indirect (Joo et al., 2007; Kinoshita et al., 2002), and the precise

function(s) of septins in epithelial and mesenchymal motility are unknown (Chacko et al.,

2005; Connolly et al., 2011b). Here, we have found that septins are required for the

stabilization of nascent FAs. Our results indicate that septins cross-link stress fibers,

maintaining the organization of the lamellar actin network, and thereby, promoting the

transmission of forces from the contractile TA to FAs. This septin function relies on the

ability of SEPT9 to crosslink preassembled actin filaments. Thus, septin upregulation

might be part of the EMT program that retools the actin cytoskeleton for a migratory

mesenchymal-like phenotype.

74

75 Figure 5.1. Septin filaments interface with RSF and TA stress fibers in the leading

lamella of renal epithelia. (A) Confocal images of SEPT2 (green), actin stress fibers

(phalloidin; red) and FAs (paxillin; white) in HGF-treated (16 h) MDCK epithelia. White

arrows point to RSFs that connect with the TA. (B) 3D rendering of the lamellar region

outlined in panel A. Arrows point to SEPT2 fibers that localize on FA-anchored RSFs.

Scale bars, 1 µm. (C) Quantification of SEPT2 colocalization with F-actin (green) and

vice versa (red). (D) SIM images show ventral and dorsal optical sections from the

lamellar region of an HGF-treated MDCK cell. Arrowhead points to SEPT9 localization

on the dorsal segment of an RSF that connects with the TA. (E) SIM images show an

HGF-treated MDCK cell stained for F-actin (phalloidin; red), SEPT2 (green) and paxillin

(white). An FA-anchored RSF (1) and TA filaments (2) are shown in higher

magnification. Arrowheads point to SEPT2 localization on and between actin filaments.

(F) Still frames show FA formation (paxillin-mCherry; red) and subsequent recruitment of

SEPT2-YFP (green). (G) Total internal reflection fluorescence microscopy kymograph

shows recruitment of SEPT2-YFP (green) to a nascent RSF (ABP140-ABD-mCherry,

red). (H) Inverted monochrome images show frames from spinning disk confocal time-

lapse microscopy of HGF-treated MDCK cells expressing ABP140-ABD-mCherry and

SEPT2-YFP. A subset of TA filaments and an RSF were pseudo-colored red and super-

imposed onto the inverted monochrome image of the SEPT2-YFP channel. SEPT2-YFP

elements recruited to the junction of the RSF with the TA were pseudo-colored green.

Overlay image shows the pseudocolored stress fibers and SEPT2 outlined in the

inverted monochrome frames. Arrow points to the RSF end and its junction point with the

TA.

76

77 Figure 5.2. Septins regulate the organization of the lamellar actin network and are

required for the stabilization of nascent FAs. (A) Confocal images of MDCK cells,

which were transfected with mCherry-expressing plasmids that encode for control or

SEPT2 shRNAs, and stained for paxillin after treatment with HGF. Red dashed line

highlights the cell edge. (B and C) Graphs show FA size and number from MDCK cells

(n = 20) treated with control, SEPT2 and SEPT9 shRNAs. (D) Graph shows FA distance

from the cell edge of MDCK cells (n = 10) treated with control (n = 903 FAs), SEPT2 # 1

(n = 749 FAs), SEPT2 # 2 (n = 512 FAs) and SEPT9 shRNAs (n = 840 FAs). (E) Graph

shows FA lifetime in three different MDCK cells treated with control (n = 6,066 FAs) and

SEPT2 shRNAs (n = 10,528 FAs). (F) Graph shows the rates of FA (n = 45) assembly

and disassembly in cells treated with control and SEPT2 shRNAs. (G) Representative

profiles of the kinetics of paxillin-GFP fluorescence in MDCK cells treated with control

and SEPT2 shRNAs. (H) Confocal images show 786-O cells stained for total paxillin

(red) and phosphorylated pY118-paxillin (green). Insets show GFP expression from

plasmids encoding for control and SEPT2 shRNAs. (I) Bar graphs show the ratio of the

fluorescence intensity of pY118-paxillin to total paxillin in MDCK cells (n = 10) treated

with control (n = 740 FAs) and SEPT2 shRNAs (n = 962 FAs), and 786-O cells (n = 10)

treated with control (n = 757 FAs) and SEPT2 shRNAs (n = 602 FAs). (J) Confocal

images of control and SEPT2-depleted MDCK cells stained for F-actin (phalloidin; red)

and paxillin (white). Insets show GFP expression from plasmids encoding for control and

SEPT2 shRNAs. (K) Plot shows line scans of phalloidin fluorescence across the leading

edges of MDCK cells (n = 13) treated with control (n = 54 line scans) and SEPT2

shRNAs (n = 56 line scans). (L and M) Graphs show the mean number of RSFs per

lamellar length (L) and mean length of RSFs (M) in MDCK cells treated with control (n =

43) and SEPT2 shRNAs (n = 72). (N) Inverted monochrome frames show ABP140-ABD-

78 mCherry from total internal reflection fluorescence (TIRF) microscopy of MDCK cells

treated with control or SEPT2 shRNAs and HGF. Arrows point to RSFs that persist and

grow overtime in control cells, and stress fibers that dissipate in SEPT2-depleted cells.

79

80 Figure 5.3. SEPT9 functions as an actin cross-linking protein in FA maturation. (A)

Confocal images of paxillin-stained MDCK cells transfected with mCherry-expressing

plasmids that encode for control or SEPT2 shRNAs, and RLC-DD-GFP, α-actinin-GFP

or GFP vectors. Scale bars, 10 µm. (B and C) Graphs show FA size (B) and number per

µm2 (C) in control cells expressing GFP (n = 1,650 FAs; 25 cells), and SEPT2 knock-

down cells expressing GFP (n = 1,567 FAs; 24 cells), α-actinin-1-GFP (n = 1,556 FAs;

20 cells), RLC-DD-GFP (n = 1,962 FAs; 18 cells) and α-actinin-1-∆ABD-GFP (n = 1,962

FAs; 20 cells). (D) Confocal images of MDCK cells transfected with mCherry-expressing

plasmids encoding for control or SEPT2 shRNAs, and α-actinin-1-GFP or GFP vectors.

Cells were stained for F-actin (phalloidin). Scale bars, 10 µm. (E) Coomassie-stained gel

shows equal volumes of supernatant (S) and pellet (P) fractions from low speed

sedimentation of pre-polymerized actin filaments in the presence of recombinant

SEPT2/6/7 and SEPT9. (F-G) Negative stain EM images of actin filaments in the

absence and presence of recombinant SEPT6 or SEPT9. Outlined regions (F) are

shown in higher magnification in G. Scale bars, 0.5 µm.

81

82 Figure 5.4. SEPT9 promotes the motility of renal epithelia during EMT. (A) Western

blot of HGF-treated MDCK cell lysates probed for SEPT9, actin and GAPDH. (B)

Graphs show the median, lowest and highest (error bars) ratio values of the SEPT9,

actin and GAPDH band intensities from three independent experiments. Values were

normalized to the protein band intensity ratios at 0 h of HGF treatment. (C) Maximum

intensity projections from 3D confocal microscopy images of HGF-treated renal cysts of

stable MDCK cells expressing SEPT9-mCherry (green) at low and high levels of

expression. 3D renal cysts were grown in collagen and stained for nuclei (DAPI; blue)

and F-actin (phalloidin; red). (D) Graphs show number of extensions per MDCK cyst (n =

23) and number of cells per extension from untransfected (n = 18), low- (n = 35) and

high-expressing (n = 51) MDCK cysts (n = 15). (E) Mean number of extensions per cyst

after treatment with DMSO (n = 23) and FCF (n = 19). (F and G) Graphs show the mean

percentage of transmigrated 786-0 cells expressing GFP and SEPT9-GFP (F), and

control and SEPT9 shRNAs from 20 different areas of a transwell filter. (H and I) Graphs

show the mean velocity (H; n = 16) and axial ratio of the long to short axes (I; n =31) of

MDCK cells migrating in a 3D matrix in the presence of HGF after transfection with

control or SEPT9 shRNAs.

83

84 Figure 5.5. Lamellar septin fibers are composed of SEPT2/6/7/9. (A-C) Confocal

images of MDCK cells stained for paxillin (white), F-actin (phalloidin; red) and

endogenous SEPT7, SEPT9 and SEPT6 (green). Outlined lamellar regions (dashed

rectangles) are shown in higher magnification. (D-F) Fluorescence images show MDCK

cells that expresses SEPT9-mCherry at endogenous-like levels after staining with

antibodies against SEPT2, SEPT6 and SEPT7. Scale bars, 10 µm. (G) Cell lysates from

a stable cell line that expresses SEPT2-YFP at subendogenous levels (see relative

intensity of SEPT2-YFP and SEPT2 bands in the “input” lanes) and untransfected

MDCKs were incubated with mouse anti-GFP antibody and immunoprecipitates were

blotted for rabbit antibodies against SEPT2, SEPT6, SEPT7 and SEPT9.

85

Figure 5.6. Quantification of septin knockdown and overexpression in MDCK cells

(A) Confocal microscopy images show MDCK cells transfected with mCherry-expressing

plasmids encoding for scrambled control, SEPT2- and SEPT9-targetting shRNAs. Cells

were stained with antibodies against endogenous SEPT2, SEPT9, SEPT7 and SEPT6.

Scale bars, 10 µm. (B) Bar graphs show quantification of SEPT2 and SEPT9

fluorescence intensity (arbitrary units; AU) per µm2 in cells transfected for 48 h with

control or SEPT2 shRNA # 1 (n = 20), SEPT2 shRNA #2 (n = 15) and SEPT9 shRNA (n

= 15). (C) Cell lysates from untransfected MDCKs and MDCKs that overexpress SEPT9-

mCherry stably were blotted for rabbit antibodies against SEPT2, SEPT6, SEPT7 and

SEPT9, and a mouse antibody against GAPDH as a loading control. (D) Bar graph

shows the ratio of fluorescence intensity (arbitrary units; AU) for each septin relative to

GAPDH.

86

87 Figure 5.7. b. (A) Confocal microscopy images show MDCK cells treated SEPT2 and

control shRNAs and stained for myosin-IIA heavy chain (red) and phosphorylated Ser19-

myosin II regulatory light chain (green). Scale bars, 10 µm. (B) Bar graph shows

quantification of the ratio of pSer-myosin II light chain to myosin IIA heavy chain

fluorescence from 5 µm-long myosin filament segments, which were blindly selected

from the lamellar regions of cells treated with control (n = 93; 12 cells) and SEPT2 (n =

74; 11 cells) shRNAs. (C) Confocal microscopy images of paxillin-stained MDCK cells

transfected with mCherry-expressing plasmids that encode for control or SEPT2

shRNAs, and GFP or the GFP-tagged non-muscle myosin IIA heavy chain mutant N93K.

Scale bars, 10 µm. (D and E) Bar graph shows FA size (D) and number per µm2 (E) in

control cells (n = 16) expressing GFP (n = 1,722 FAs), and SEPT2 knock-down cells (n =

16) expressing GFP (n = 1,515 FAs) and myosin IIA N93K-GFP (n = 1,139 FAs).

88 CHAPTER 6. Septin 9 exhibits polymorphic binding to F-actin and inhibits

myosin and cofilin activity

Abstract

Septins are a highly conserved family of proteins in eukaryotes that is recognized

as a novel component of the cytoskeleton. Septin 9 (SEPT9) interacts directly with actin

filaments and functions as an actin stress fiber cross-linking protein that promotes the

maturation of nascent focal adhesions and cell migration. However, the molecular details

of how SEPT9 interacts with F-actin remain unknown. Here, we use electron microscopy

and image analysis to show that SEPT9 binds to F-actin in a highly polymorphic fashion.

We demonstrate that the basic domain (B-domain) of the N-terminal tail of SEPT9 is

responsible for actin cross-linking, while the GTP-binding domain (G-domain) does not

bundle F-actin. We show that the B-domain of SEPT9 binds to three sites on F-actin,

and the two of these sites overlap with the binding regions of myosin and cofilin. SEPT9

inhibits actin-dependent ATPase activity of myosin and competes with the weakly-bound

state of myosin for binding to F-actin. At the same time, SEPT9 significantly reduces the

extent of F-actin depolymerization by cofilin. Taken together, these data indicate that

SEPT9 protects actin filaments from depolymerization by cofilin and myosin, and

suggest a mechanism by which SEPT9 could maintain the integrity of growing and

contracting actin filaments.

Introduction

Septins were first discovered in the budding yeast as genes that are crucial for

cell division (Hartwell, 1971). Since the discovery of septins in yeast, proteins with

homologous sequences have been found in almost all eukaryotic cells. The number of

septin genes per organism is variable and ranges from two isoforms in Caenorhabditis

89 elegans to 13 isoforms in humans (Russell and Hall, 2011). Based on sequence

similarity, human septins are classified into four homology groups named after their

founding members: SEPT2, SEPT3, SEPT6 and SEPT7. In this manuscript we focus on

SEPT9 which belongs to the SEPT3 subgroup and has a unique N-terminal domain

(NTD) consisting of a basic domain (B-domain) and an acidic domain (A-domain) (Fig.

1a) (Bai et al., 2013). All septins have a GTP-binding domain (G-domain), which is

evolutionarily and structurally related to the Ras GTPases (Leipe et al., 2002). Unlike the

monomeric small GTPases, septins form polymers that consist of non-polar oligomeric

complexes(Bertin et al., 2008; Sirajuddin et al., 2007)

Septins have been recognized as the fourth component of the cytoskeleton

because of their filamentous appearance and their interdependence with actin filaments

and microtubules. The N-terminal domain of SEPT9 has been shown to bind and bundle

microtubules by interacting with the acidic C-terminal tails of β-tubulin (Bai et al., 2013).

Septins have been shown to bind actin filaments indirectly via anillin or the motor protein

myosin II(Joo et al., 2007; Kinoshita, 2002). Recent data, however, demonstrated that

Drosophila septin complexes (Sep1-Sep2-pnut) can directly interact with actin and

cross-link actin filaments into curved bundles, which are critical for the organization and

function of contractile actin rings during Drosophila embryonic cleavage (Mavrakis et al.,

2014). Moreover, we recently found that human SEPT9 directly binds and bundles F-

actin, and that cross-linking of actin filaments by SEPT9 promotes focal adhesion

maturation and epithelial cell motility (Dolat et al., 2014b).

Despite the new evidence that septins can directly interact with the actin

cytoskeleton, the molecular details of these interactions and how septins interact with

the surface of the actin filament remain unknown. Here we use electron microscopy,

90 image analysis and co-sedimentation assays to identify how SEPT9 interacts with F-

actin. Our data indicate that the B-domain of SEPT9 is responsible for its F-actin

bundling activity, while the G-domain of SEPT9 does not bundle actin filaments. We

demonstrate that SEPT9 binds to three sites on the surface of F-actin that are commonly

bound by other actin binding proteins. Importantly, two sites overlap with the regions of

the actin molecule involved in the binding of myosin and cofilin. As predicted by the

structural data, SEPT9 inhibits F-actin depolymerization by cofilin, which severs actin

filaments, and reduces the actin-dependent ATPase activity of myosin. Our results

suggest that SEPT9 could protect growing actin filaments from the combined

depolymerizing activity of cofilin and myosin during actin filament assembly and

contraction.

Materials and Methods

Protein expression and purification. G-Mg2+-actin was prepared as described (Orlova

and Egelman, 1995). SEPT9 constructs were prepared as follows. E. Coli BL21 (DE3)

cells were transformed with SEPT9_i1, SEPT9-NTD, SEPT9-G, SEPT9-A, or SEPT9-B

in kanamycin-resistant pET-28a (Bai et al., 2013). Plasmids expressing His-tagged

SEPT9-N (aa 1–283 of SEPT9_i1), SEPT9-B (aa 1–142 of SEPT9_i1), SEPT9-A (aa

143–283 of SEPT9_i1), SEPT9-G (aa 284–586 of SEPT9_i1) were made with the

QuikChange II Site-Directed Mutagenesis kit (Agilent Technologies) using the pET-

SEPT9_v1 and pET-SEPT9_v3 plasmids. Bacterial cultures of OD600 were induced with

0.5 mM IPTG for 16h at 18°C, collected by centrifugation and resuspended in lysis buffer

containing 50 mM Tris, pH 8.0, 1% Triton X-100, 150 mM NaCl, 10% glycerol, and 10

mM imidazole. Bacteria were lysed using a French pressure cell at 1,280 psi, and

lysates were cleared using centrifugation (14,000 g) for 30 min at 4°C. Columns

91 containing Ni-NTA beads were equilibrated with lysis buffer prior to passing

supernatants through by gravity flow. The beads were washed with buffer containing

50mM Tris, pH 8.0, 300 mM NaCl, 10% glycerol, and 10 mM imidazole, and proteins

were eluted with buffer containing 50mM Tris, pH 8.0, 300 mM NaCl, 10% glycerol, and

250 mM imidazole.

MyosinV-S1 heavy chain, 907 amino acid residues including 6IQ domains was

expressed in a baculovirus/Sf9 cell system. The proteins were purified using FLAG-

affinity chromatography and then concentrated and fractionated on a MonoQ ion-

exchange column with a linear gradient of 0.1−0.5 M NaCl in 10 mM MOPS, 0.1 mM

EGTA, 3 mM NaN3, and 1 mM DTT, pH 7.2, using HPLC. The concentration of myosinV-

S1 was determined from the A280 after correction for light scattering (A280−1.5A320).

The extinction coefficient was calculated to be 1.1 × 105 M-1 cm-1 from the number of

tyrosine and tryptophan residues and their molar extinction coefficients:  ε = (n = 10)trp

× 5690 + (n = 42)tyr × 1280. All preparations were analyzed by SDS protein gel

electrophoresis and by active site titration with deac-aminoATP. Cofilin-1 was a

generous gift from Dr. Emil Reisler (UCLA).

Electron microscopy of negatively stained SEPT9 domains. G-actin was

polymerized for 1–2 hours in F-buffer (10mM HEPES, pH 7.4, 100mM KCl, 1mM MgCl2,

0.5mM ATP). For EM samples shown in Fig. 1c-g, 2 µM F-actin was incubated for 2 min

in tube with 34 µM NTD, B-domain, A-domain, or G-domain. Samples were blotted and

negatively stained with 2% (wt/vol) uranyl acetate. To obtain single actin filaments

decorated with SEPT9 domains was 2 µM F-actin was incubated for 1 min on carbon

coated glow discharged grids in a humidified chamber, gently blotted and incubated

again with 34 µM NTD, B-domain, A-domain, or G-domain. Samples were negatively

92 stained with 2% (wt/vol) uranyl acetate. A JEM-2100F (JEOL) Field Emission

Transmission Electron Microscope, equipped with 11-megapixel Gatan SC1000 ORIUS

CCD camera, was used at an accelerating voltage of 200 kV, and a nominal

magnification of 60,000x to record micrographs at a raster of 1.58 Å/pixel.

Electron microscopy of SEPT9 in presence of cofilin-1 and SEPT9. F-actin (2 µM)

was mixed with 10 µM of cofilin-1, or preincubated with 10 µM of SEPT9 before addition

of 10 µM of cofilin-1. Samples were left at room temperature and analyzed by TEM at 2

and 30 min time points.

Image analysis. The SPIDER software package (Galkin et al., 2001) was used for most

image processing, while the EMAN package (Galkin et al., 2003) was used to extract

filament images from micrographs. Images were corrected for the contrast transfer

function (CTF) using theoretical CTF obtained with the CTFIND software (Frank, 1984)

and decimated to 3.16 Å/pixel scale. From these CTF-corrected images segments (each

120 pixels long) of F-actin decorated with the NTD (n=8,193), B-domain (n=2,708), A-

domain (n=5,435) constructs were extracted. Segments of filament decorated with the

NTD were reconstructed using IHRSR approach (Ludtke et al., 1999). The resultant

reconstruction showed three additional densities bound to F-actin - one at the front

interface between SD1 and SD2 of two adjacent actin protomers, the other one at the

side of SD1, and one on the top of SD4 (Fig. S1). Three model volumes were created

by using pseudo-atomic model of F-actin (Mindell and Grigorieff, 2003) having model

density attached to the first site, second site, or third site. These volumes, along with a

naked F-actin volume, were projected onto 120 × 120-pixel images with an azimuthal

rotational increment of 4°, and the resultant 360 reference projections (4 × 90) were

cross-correlated with the 8,193 NTD-decorated actin filament segments. Each of the four

93 classes was reconstructed starting from the two independent starting models (Fig. S2).

The naked class (n=4,222) yielded a symmetry of 166.6°/27.3 Å, the mode1 class

(n=1,590) yielded a symmetry of 166.7°/27.2 Å, the mode 2 class (n=1,305) converged

to 166.1°/27.2 Å, while a set of images classified as mode 3 (n=1,079) reached a stable

solution having 167.0°/27.2 Å symmetry values. The same sorting procedure was

applied to actin filaments decorated with the B-domain. The naked segments extracted

from the filaments decorated with the B-domain (n=1,571) converged to a 166.6°/27.2 Å

solution, the mode1 class from that set (n=473) yielded 166.3°/27.1 Å, the mode 2 class

(n=332) converged to a symmetry of 166.1°/27.3 Å, while the mode 3 class (n=332)

yielded a symmetry of 166.5°/27.2 Å. Segments of filaments incubated with the A-

domain were mostly naked actin (n=4,935) with only 218 segments assigned to mode 1

class, 135 segments assigned to the mode 2 class, and 147 segments assigned to the

mode 3 class. Neither of these classes yielded reasonable 3D reconstruction (data not

shown). The naked class converged to the stable solution of 166.5°/27.1 Å. Fourier Shell

Correlation approach was used to determine the resolution of the 3D-reconstructions.

Each image set was split into two non-overlapping subsets, and these subsets were

reconstructed independently. Resultant volumes were used for resolution determination.

Using 0.5 criterion all maps yielded 27-28 Å resolution (Fig. S7). All the 3D-

reconstructions shown in the paper were filtered to the measured resolution. The

Chimera software (Egelman, 2000) was used to dock a protomer from the atomic model

of F-actin into the reconstruction manually. Figures 2, 3 and 4 were produced using the

Chimera software.

Actin activated steady-state ATPase experiments. Actin activated steady-state

experiments were done by the NADH coupled assay at 25 °C. 20 nM Myosin V-S16IQ

was mixed with 10 µM actin and increasing concentrations of SEPT9 A- domain , B-

94 domain or full length SEPT9 in a buffer containing 10 mM MOPS (pH 7.0), 2 mM MgCl2,

25 mM KCl, 0.15 mM EGTA, 2 mM ATP, 40 units/ml lactate dehydrogenase, 200

units/ml pyruvate kinase, 1 mM phosphoenolpyruvate, and 200 μM NADH (Fujii et al.,

2010). Changes in NADH absorption at 340 nm (€340 = 6220 M-1cm-1) were followed in

a Beckman DU640 spectrophotometer. The ATPase activity of blanks containing actin

but no myosin was subtracted from the actomyosin data. The data was fit to the equation

F (x) = Vmax/1+x/Ki .

Low-speed actin pelleting assays. G-actin in G buffer (5 mM Tris-HCl and 0.2 mM

CaCl2) was polymerized in buffer containing 20 mM Hepes, pH 7.4, 100 mM KCl, and 1

mM MgCl2 supplemented with 0.2 mM ATP and 4 mM DTT for 1 h at 25°C. In Figure 1, 8

μM of recombinant SEPT9-N, SEPT9-G, SEPT9-A, or SEPT9-B were incubated with 2

μM F-actin for 1 h at 25°C. In Supplemental Figure 6, increasing concentrations of full-

length SEPT9 or SEPT9-B were incubated with 2 μM F-actin for 1 h at 25°C. Reactions

were subsequently centrifuged on top of a glycerol cushion (20 mM Hepes, 100 mM

NaCl, 1 mM MgCl2, and 10% glycerol) at 8,000 g for 20 min at 25°C. Supernatant and

pellet fractions were collected in SDS sample buffer, and equal volumes of each fraction

were resolved with a 10% or 6-12% gradient SDS-PAGE gel and stained with

Coomassie brilliant blue. Protein band densities from scanned gels were quantified using

the Odyssey infrared scanning system (LI-COR Biosciences) and analyzed in excel. All

experiments were repeated 3 times.

High-speed actin pelleting assays. Competition between SEPT9 and the myosin V-S1

heavy chain for F-actin binding were performed in buffer containing 10mM MOPS, pH

7.0, 100 mM KCl, 2 mM MgCl2, 0.5 mM EGTA, and 2 mM DTT with 1.25 mM ATP or

without ATP. For experiments without ATP, the F-actin stock was dialyzed in F-buffer

95 without ATP. F-actin (2 μM) was pre-incubated with increasing concentrations of SEPT9

for 30 minutes at 25°C. Reactions were incubated with myosin V-S1 heavy chain (0.75 –

1 μM) for an additional 30 minutes at 25°C, placed on a glycerol cushion (10mM MOPS,

pH 7.0, 100 mM KCl, 2 mM MgCl2, 0.5 mM EGTA, 20% glycerol) and centrifuged at

200,000 g for 30 minutes at 21°C. Equal volumes of supernatant and pellet fractions

were collected in SDS sample buffer, and equal volumes of each fraction were resolved

using a 10% SDS-PAGE gel and stained with Coomassie brilliant blue. Protein band

densities were quantified using the Odyssey infrared scanning system and analyzed in

excel. Each experiment was repeated 3 times.

Co-sedimentation assays (Fig. 4) that tested for cofilin-mediated

depolymerization were performed by pre-incubation of aged F-actin with recombinant

SEPT9_i1 (5 μM) or 5 μM SEPT9 fragments (NTD, B-domain or A-domain) in buffer

containing 10 mM Hepes, pH 7.6, 100 mM KCl, 1 mM MgCl2, and 2 mM DTT for 30 min

at 25°C. Reactions were incubated with 5 μM recombinant human cofilin-1 for an

additional 30 min at 25°C, placed on a glycerol cushion (20 mM HEPES, pH 7.6, 100

mM NaCl, 1 mM MgCl2, and 10% glycerol) and centrifuged at 200,000 g for 30 min at

21°C in an ultracentrifuge (Optima TL100, Beckman Coulter). Supernatant and pellet

fractions were collected in SDS sample buffer, and equal volumes of each fraction were

resolved using a 8-15% gradient SDS-PAGE gel and stained with Coomassie brilliant

blue. Protein band densities from scanned gels were quantified using the Odyssey

infrared scanning system (LI-COR Biosciences) and analyzed in excel. Each experiment

was repeated 3 times.

High-speed pelleting assays between SEPT9 and F-actin (Fig. S5) were

performed by incubating full-length SEPT9 (1 μM) with increasing concentrations of F-

96 actin for 30 minutes at 25°C in buffer containing 20 mM Hepes, pH 7.6, 100 mM KCl,

and 1 mM MgCl2 supplemented with 0.2 mM ATP and 4 mM DTT. Reactions were

placed on a glycerol cushion (20 mM HEPES, pH 7.6, 100 mM NaCl, 1 mM MgCl2, and

20% glycerol) and centrifuged at 200,000 g for 30 min at 21°C. Supernatant and pellet

fractions were collected in SDS sample buffer, and equal volumes were resolved using a

10% SDS-PAGE gel. Gels were stained with Coomassie brilliant blue, and protein band

densities from scanned gels were quantified using the Odyssey infrared scanning

system. The fraction of SEPT9 protein in each pellet fraction was plotted against the

concentration of actin in GraphPad and fitted with a one-site specific binding curve to

determine the dissociation constant.

RESULTS

The B-domain of SEPT9 promotes F-actin bundling.

Using negative stain EM and low-speed pelleting assays, we sought to

understand how SEPT9 interacts with F-actin by expressing and purifying each of the

three SEPT9 domains; the domain composition of SEPT9 is shown in Fig. 1a. Full-length

SEPT9 (isoform 1) as well as the N-terminal domain (NTD) of SEPT9 effectively

assemble F-actin into bundles (Fig. 1c and d, respectively). Strikingly, the B-domain

alone promotes formation of ordered thick bundles (Fig. 1e), while the A-domain does

not possess any bundling activity (Fig. 1f). Similarly to the A-domain, the G-domain of

SEPT9 does not possess any bundling activity (Fig. 1g). Our results indicate that the B-

domain of SEPT9 is responsible for the SEPT9 F-actin bundling activity. To

independently corroborate these EM observations, we examined the ability of individual

SEPT9 domains to cross-link actin filaments using a low speed actin sedimentation

97 assay (Fig. 1h and i). In agreement with the EM data, the NTD (Fig. 1h) and B-domain

(Fig. 1i) of SEPT9 pelleted F-actin at low speed, while the G-domain (Fig. 1h) and the A-

domain (Fig. 1h) did not. Taken together, these data indicate that the B-domain of the

NTD of SEPT9 is responsible for the actin-bundling activity of SEPT9.

SEPT9 binds to three sites on the surface of F-actin.

To identify the binding site of SEPT9 on the surface of F-actin we performed a

3D-reconstruction of actin filaments decorated with the two SEPT9 actin bundling

fragments – the NTD and the B-domain (Fig. 2). In order to obtain single actin filaments

suitable for image analysis, the NTD or B-domain were incubated with actin filaments

that were pre-bound to a carbon film to reduce bundle formation. The overall 3D-

reconstruction of F-actin segments (each segment contained ~14 actin subunits)

decorated with the NTD of SEPT9 revealed three sites of interaction of the NTD with the

actin filament (Fig. S1). We used a modeling approach and cross-correlation sorting (see

Methods) to compute individual 3D-reconstructions for each of the three modes of

binding of the NTD to F-actin revealed in the overall reconstruction (Fig. 2a). Since

model-based sorting may be biased, we used two independent approaches to confirm

that our 3D-reconstructions reflected the intrinsic properties of the images. First, we

computed two independent 3D-reconstructions for each of the three structural modes

starting the IHRSR algorithm from a featureless solid cylinder, or from a 3D-

reconstruction of a different structural mode (Fig. S2). This approach proved that the

resultant EM map was independent from the starting model and hence all its features

reflected the intrinsic properties of the raw images (Fig. S2). Second, we calculated

reference free 2D-averages for each class of NTD-decorated F-actin segments to

confirm that all three 2D averages were different from each other and hence, the

98 differences in the 3D reconstructions reflected the differences in the raw images (Fig.

S3).

The NTD of SEPT9 binds to three distinct sites on the surface of F-actin. In mode

1 (Fig 2a, magenta arrows), the NTD binds to the side of the SD1/SD2 interface of the

actin molecule which includes actin residues 80-101, 125-130, and 357-360 (Fig. 2d,

shown in magenta). In mode 2 (Fig. 2a, red arrows), the NTD is bound to the front

portion of the SD1/2 interface, which involves residues 1-7, 21-30, 56-60, and 92-103

(Fig. 2d, shown in red). In mode 3 (Fig. 2a, cyan arrows), actin residues 223-238 and

248-251, which belong to the SD4 of F-actin (Fig. 2d, shown in cyan), are located at the

interface with the NTD.

We used the same set of references to determine how the B-domain of SEPT9

binds to F-actin (Fig. 2b). We found the B-domain is bound to the same three sites on

the surface of the actin molecule. We applied the same sorting procedure to F-actin

incubated with the A-domain of SEPT9 and compared the frequency of decorated and

naked actin segments between the three SEPT9 domains (Fig. 2c). In contrast to the

NTD and the B-domain, more than 90% of actin segments incubated with the A-domain

yielded a best match with naked F-actin. The remaining 10% generated an

uninterpretable 3D-reconstruction (data not shown). To exclude any possibility that the

A-domain binds to F-actin, we used the same set of references to run cross-correlation

sorting of pure F-actin segments (Fig. S4). Not surprisingly, the sorting showed a very

similar distribution of the pure F-actin segments between the four classes which

reflected the margin of error in the sorting procedure.

We did not find a predominant mode of NTD or B-domain binding to F-actin as

the frequencies of the three structural modes were quite similar (Fig. 2c). Because we

99 did not find any segments with all three sites occupied simultaneously, individual binding

modes of SEPT9 appear to be highly cooperative and mutually exclusive; note that full

length SEPT9 exhibited the same three modes of actin binding as the NTD and B-

domains (data not shown). It is therefore likely that if a SEPT9 molecule is bound to an

actin protomer in either mode 1, 2 or 3, adjacent SEPT9 molecules would also bind actin

protomers in the same mode. Such a cooperativity has been reported for many actin

binding protein and relies on cooperative structural transitions that can propagate along

the actin filament (Galkin et al., 2005; Galkin et al., 2010; Orlova and Egelman, 1997). In

addition to the structural cooperative transitions within the actin filament, the spatial

arrangement of the SEPT9 domains on the surface of F-actin could result in mutually

exclusive modes of SEPT9 binding to F-actin. Binding modes 1 and 2 have an

overlapping interface on the surface of F-actin (Fig. 2d, 80-101 helix in mode 1 and 92-

103 in mode 2), and modes 2 and 3 would clash spatially if present simultaneously

within the same actin filament. Taken together, our results indicate that SEPT9 interacts

with F-actin via the B-domain of its NTD in a polymorphic fashion. Importantly, this

interaction involves actin-binding sites, which are commonly bound by actin-binding

proteins including myosin and cofilin (Behrmann et al., 2012; Galkin et al., 2011).

SEPT9 inhibits actomyosin activity

The localization of myosin S1-fragment on the surface of F-actin in the absence

of ATP (rigor state) has been previously resolved (Behrmann et al., 2012). While the

position of myosin on the surface of the actin filament in the presence of ATP (weak

binding mode) has not been visualized, modeling studies based on the X-ray diffraction

from permeabilized muscles suggest that the N-terminus of F-actin is at the interface

with the weakly bound myosin head (Gu et al., 2002). Our data suggest that the position

100 of myosin in a rigor state would clash with the position of the NTD of SEPT9 in two of its

three binding modes (Fig. 3a-b, red arrows); the N-terminus of F-actin (denoted with red

asterisk in Fig. 3a-c) is also at the interface of NTD with F-actin. Thus, binding of SEPT9

to F-actin in modes 1 or 2 could interfere with the actin binding of myosin in both rigor

and weak states (Fig. 3a-b), while binding of SEPT9 to F-actin in mode 3 may not

interfere with the acto-myosin interaction (Fig. 3c).

To test these structural predictions, we first used an actin-activated steady-state

ATPase assay. In this assay, myosin ATP hydrolysis is coupled with its interaction with

F-actin. Thus, any protein that interferes with the actomyosin interaction will reduce the

ATPase activity of the myosin head. To evaluate the effect of SEPT9 on the actomyosin

cross-bridge cycle, we carried out the actin activated steady-state ATPase assay using

the NADH coupled method (Fig 3d). The S1 fragment of myosin V and F-actin were

mixed with increasing concentrations of full length SEPT9, or its individual B- and A-

domains domains (see Methods for details). The full length SEPT9 exhibited the largest

inhibition with an apparent Ki of 0.46 µM followed by the B-domain with a Ki of 2.44 µM.

The A-domain had almost no effect on myosin ATPase activity with an apparent Ki of

185 µM.

Since bundling would reduce the number of F-actin sites that are available for

myosin binding, we tested whether the reduction in myosin ATPase activity correlated

with an increase in F-actin bundling. Low-speed actin sedimentation assays showed that

both the full length SEPT9 and SEPT9 B-domain did not significantly affect F-actin

bundling over concentration ranges that caused a drastic reduction in myosin activity

(Fig. S5). Hence, the bundling activity of SEPT9 by itself could not account for full

inhibition of the acto-myosin ATPase activity. Taken together with our structural data that

101 predict a mutually exclusive binding of myosin and SEPT9 in modes 1 and 2 (Fig. 3a,b),

our results indicate that SEPT9 inhibits actin-dependent myosin ATPase activity by

competing with myosin for binding to the same sites of F-actin.

ATP hydrolysis and subsequent dissociation of the nucleotide from the myosin

head changes acto-myosin interaction from the initial weakly-bound state to a rigor-

bound state. This transition in the acto-myosin interaction is an essential part of myosin’s

ATPase cross-bridge cycle. To test how SEPT9 inhibits acto-myosin activity, we

performed competitive high-speed actin pelleting assays of myosin V S1-fragment with

the full length SEPT9 in the presence or absence of ATP, so we can distinguish between

the weakly-bound (ATP) and rigor-bound (no ATP) states of myosin. In the presence of

ATP, full length SEPT9 efficiently competed with the myosin S1-fragment for binding to

F-actin (Fig. 3e). Interestingly, in the absence of ATP (only ADP was present in solution)

SEPT9 co-sedimented with myosin S1-fragment (Fig. 3f). The affinity of the ADP-bound

myosin V to F-actin (~7.6 nM) is ~1000 fold higher than its affinity to F-actin in the

presence of ATP (~13 µM) (De La Cruz et al., 1999; Yengo et al., 2002). Taking into

account that the affinity of septins to F-actin is in the micromolar range(Mavrakis et al.,

2014) (Fig. S6), SEPT9 molecules that are bound to F-actin in modes 1 and 2 are likely

to be displaced by ADP-bound myosin, which has a high affinity for F-actin.

Cosedimentation of SEPT9 with rigor-bound myosin V suggests that SEPT9 is bound to

F-actin in mode 3 (Fig. 3c), which does not clash with the position of rigor-bound myosin.

In the presence of ATP, however, SEPT9 is likely to compete with myosin for the binding

modes 1 and 2, which account for ~70% of the binding of SEPT9 to F-actin (Fig. 2c), and

thereby, SEPT9 can inhibit the interaction of myosin with F-actin. To summarize, our

data indicate that SEPT9 inhibits the weak acto-myosin interaction through its B-domain,

while it can bind to F-actin in the presence of rigor-bound myosin.

102 SEPT9 reduces cofilin-driven F-actin depolymerization.

Superimposition of the 3D reconstructions of SEPT9-decorated F-actin in mode 1

(Fig. 4a, top) and mode 2 (Fig. 4a, bottom) revealed significant clashes between the

SEPT9 density (Fig. 4a, black arrows) and cofilin (Fig. 4a, shown as solid red surface).

Similar to our results with myosin, we hypothesized that SEPT9 may inhibit cofilin-actin

binding. Using electron microscopy and high-speed pelleting assays, we sought to test

whether SEPT9 affects cofilin-driven F-actin depolymerization.

EM imaging of actin filaments showed that within 2 min of incubation, cofilin

disrupts the meshwork of naked actin filaments, which are disintegrated into short

twisted filaments (Fig. 4b, 2 min). After 30 min, only a few cofilin-decorated actin

filaments remain (Fig. 4b, 30 min). In the presence of full length SEPT9, F-actin bundles

(Fig. 4c, 0 min) remain intact after 2 min of incubation with equimolar concentration of

cofilin-1 (Fig. 4c, 2 min). Even after 30 min of incubation with cofilin-1, F-actin bundles

are still present (Fig. 4c, 30 min). To further test whether SEPT9 protects against actin

depolymerization by cofilin, we performed high-speed sedimentation assays in the

presence of full length SEPT9 or its domains and cofilin-1 (Fig. 4,d-f). In our

experiments we used equimolar concentrations of SEPT9 and cofilin, because the

affinity of human ADF/cofilin to ADP-F-actin is similar to septins (Mavrakis et al., 2014) (Fig. S6).

After 30 min incubation of F-actin with cofilin-1, ~90% of actin is recovered in the

supernatant (Fig. 4d). Pre-incubation of F-actin with SEPT9 reduced the amount of actin

in the supernatant to ~30% (Fig. 4d,). The NTD (Fig. 4e) and the B-domain (Fig. 4f) of

SEPT9 also reduced the amount of depolymerized actin in the supernatant to ~50-60%

of total actin, while the A-domain had no effect (Fig. 4f). Thus, similar to the inhibition of

103 myosin binding to F-actin, SEPT9 appears to inhibit cofilin binding, and thereby, protect

actin filaments from cofilin-mediated depolymerization.

Discussion

Drosophila septins have been recently reported to directly bind and bundle F-

actin (Mavrakis et al., 2014). Mammalian septins (SEPT2/6/7 oligomers) have been

proposed to interact with F-actin indirectly via anillin and myosin II, but it was recently

shown that the mammalian SEPT9 can bind and bundle actin filaments (Dolat et al.,

2014b; Joo et al., 2007; Kinoshita et al., 2002). Our data show that the actin bundling

activity of SEPT9 resides in the NTD – a unique domain that is not present in other

septin paralogs. SEPT9 interacts with F-actin in a highly polymorphic fashion (Fig. 2a-b).

Such multiplicity in binding modes in not unique to SEPT9 and has been reported for

other actin binding proteins (ABPs) such as Arg/Abl kinase, drebrin, myosin binding

protein C, utrophin, and a giant muscle protein nebulin (Galkin et al., 2005; Galkin et al.,

2002; Grintsevich et al., 2010; Lukoyanova et al., 2002; Orlova et al., 2011) Since, the

N-terminal domain of SEPT9 does not possess any detectable amino acid identity with

these ABPs, it suggests that multiplicity of actin-binding modes is a common feature of

ABPs independently of their actin-binding sequences and domains.

Recently we showed that F-actin can exist in multiple structural states and that

the structural transitions in the mobile regions of the actin protomers are coupled (Galkin

et al., 2010). SEPT9 binds to the two mobile regions of actin molecule including

residues 1-7 of the N-terminus and residues 56-60 of the SD2 (Fig. 2d). It is plausible

that in the cell the structural mode of SEPT9 binding to F-actin is determined by the

structural state of the actin filament and the presence of other actin-bound ABPs. Our

results suggest that SEPT9 may switch from mode 1 and 2, where it interferes with the

104 actin-myosin interaction (Fig. 3a-b), to mode 3 where both rigor-bound myosin and

SEPT9 can simultaneously interact with F-actin. By switching between these structural

modes of binding, SEPT9 may alter acto-myosin interactions in migrating cells, where

tensile forces are tightly controlled and are linked to focal adhesion turnover (Parsons et

al., 2010; Peacock et al., 2007) and cell-ECM adhesion (Vicente-Manzanares et al.,

2007). Thus, SEPT9 may influence actomyosin activity at stress fibers and focal

adhesions.

SEPT9-dependent actin bundling activity is required for the maintenance of a

functional stress fiber network in the leading edge of migratory epithelial cells (Dolat et

al., 2014b). Interestingly, our results indicate that SEPT9 protects actin filaments from

depolymerization by cofilin. Actin cross-linking proteins have been shown to protect actin

filaments from depolymerization in vitro (Cano et al., 1992; Lebart et al., 2004; Zigmond

et al., 1992). In living cells, varying levels of expression of actin bundling proteins have

been shown to alter the rates of actin filaments turnover (Mukhina et al., 2007; Tilney et

al., 2003) and myosin has been shown to regulate the turnover of actin bundles in

neuronal growth cones (Ishikawa et al., 2003; Medeiros et al., 2006).

In addition, recent studies show that actin depolymerization by ADF/cofilin is

differentially regulated by actin motors and actin-cross-linking proteins (Schmoller et al.,

2011). While myosin II enhances the disintegration of F-actin bundles, cross-linking

proteins inhibit the disassembly of actin bundles by cofilin (Schmoller et al., 2011).

Notably, cross-linking proteins such as fascin may affect the severing activity of cofilin by

reducing the flexibility of bundled F-actin, and thereby, limit the torsional twist that cofilin

triggers on F-actin (Breitsprecher et al., 2011). Because the efficiency of cofilin severing

is the highest between decorated and non-decorated segments of F-actin, the severing

105 activity of cofilin will be the highest on more rigid bundled actin filaments, which do not

allow for a uniform, cooperative binding that is enabled by a torsional twist (Breitsprecher

et al., 2011). Thus, depending on the ABP composition of F-actin bundles and their

flexibility, cross-linking proteins such as fascin may either increase or decrease the

activity of cofilin (Breitsprecher et al., 2011; Elkhatib et al., 2014)

Our results indicate that SEPT9 inhibits both myosin and cofilin interaction with

F-actin, and hence may work as a protective factor for growing actin bundles. Given that

both myosin (Juanes-Garcia et al., 2016)and cofilin (Bernstein and Bamburg, 2010) are

involved in the regulation of actin organization and contractility in a diversity of cellular

processes and structures, our results suggest that the ubiquitously expressed SEPT9

(Dolat et al., 2014a) may be a key player in the regulation of F-actin organization and

function.

106

107 Figure 6.1. SEPT9 bundles F-actin through the B-domain of its NTD. (a) Schematic

representation of SEPT9 domains. In addition to the GTP-binding domain (G-domain)

found in all septin paralogs, SEPT9 has a unique N-terminal domain (NTD) consisting of

a basic domain (B-domain) and an acidic domain (A-domain). (b) Pure F-actin, scale

bar: 1 nm. (c) Full-length SEPT9 efficiently bundles actin filaments within 2 min of

incubation. (d) NTD consisting of the B- and A-domains packs F-actin into tight bundles.

(e) The B-domain bundles F-actin into tight bundles morphologically similar to those

shown in (d). (f) The A-domain does not bind or bundle F-actin. (g) The G-domain does

not bundle F-actin. (h and i) Low-speed sedimentation of F-actin in the presence of

SEPT9 domains. Coomassie-stained gels with the corresponding bar graphs show that

F-actin is recovered in the pellet in the presence of the NTD [(h), SEPT9-NTD] and the

B-domain [(i), SEPT9-B]. Error bars correspond to the maximum and minimum values

from three independent experiments. In the presence of the G-domain [(h), SEPT9-G]

and the A-domain [(i), SEPT9-A], most of the actin is recovered in the supernatant.

108

Figure 6.2. The NTD and B-domain of SEPT9 bind to three sites on the surface of F-

actin. (a and b) 3D reconstructions show the three modes of binding of the NTD (a) and

B-domain (b) to F-actin. Three sites on the surface of the actin filament are shown: side

of actins SD1/SD2 (magenta arrows), front portion of the same subdomains (red arrows)

and the top of SD4 (cyan arrows). Actin molecules are shown as blue ribbons while non-

filled empty envelopes represent additional mass on the surface of the actin filament that

corresponds to the sites of binding of SEPT9 constructs to F-actin. (c) Cross-correlation

sorting of segments of actin filaments decorated with SEPT9 domains shows that the

NTD and B-domain bind to the three sites on F-actin with similar frequency, while the A-

domain does not bind to F-actin. (d) Actin residues that are likely to be involved in the

interaction with SEPT9 are marked on the actin monomer (yellow ribbons) for mode 1 in

magenta, mode 2 in red and mode 3 in cyan.

109

110 Figure 6.3. SEPT9 inhibits the activity and interaction of myosin with actin. (a–c) An

atomic model of the actomyosin complex in rigor state [14] was docked onto the 3D

reconstructions of the three modes of binding of the SEPT9 NTD to F-actin (Fig. 2a)

using the best fit of the actin molecules. Actin molecules are shown as blue ribbons and

myosin head is depicted as solid surface. The clashes between the myosin head and the

NTD density (red arrows) in modes 1 and 2 are marked with red arrows (a and b). There

are no such clashes in mode 3 (c). The position of the N-terminus of F-actin that resides

at the interface between the NTD and F-actin is denoted with red asterisks. (d) Plot

shows the ATPase activity of the S1 fragment of myosin V in the absence and presence

of full-length SEPT9 and the A- and B-domains of SEPT9. As predicted by the structural

data, full-length SEPT9 and the B-domain inhibit the ATPase activity of myosin, while the

A-domain does not have a significant effect. (e) Coomassie-stained gels show the pellet

(P; top) and supernatant (S; bottom) fractions from a competitive high-speed actin

pelleting assay in the presence of ATP (1.25 μM). F-Actin (2 μM) was incubated with

increasing concentrations of recombinant full-length SEPT9 and the myosin S1 fragment

was added at a constant concentration (0.75 μM). (f) Bar graphs show the fraction of

total S1 fragment in the pellet and supernatant. Error bars correspond to the minimum

and maximum values obtained from three independent experiments. (g) Coomassie-

stained gels show the pellet (P; top) and supernatant (S; bottom) fractions from a

competitive high-speed actin pelleting assay in the absence of ATP. F-Actin (2 μM) was

incubated with increasing concentrations of recombinant full-length SEPT9 and the

myosin S1 fragment was added at a constant concentration (1 μM). (f) Bar graphs show

the fraction of total S1 fragment in the pellet and supernatant. Error bars correspond to

the minimum and maximum values obtained from three independent experiments.

111

112 Figure 6.4. SEPT9 reduces the extent of cofilin-driven F-actin depolymerization. (a)

Atomic model of actin–cofilin complex [15] docked into the 3D reconstructions of modes

1 and 2 (transparent gray surfaces) shows a clash between the cofilin molecule (red

surface) and SEPT9 density (marked with black arrows). (b) Long F-actin filaments

formed after 2 h of polymerization (b; 0 min) are disrupted into short segments within

2 min of incubation with cofilin-1 (b; 2 min); after a 30-min incubation with cofilin (b;

30 min), only few filaments that are heavily decorated with cofilin remain. (c) F-actin

forms tight bundles in the presence of SEPT9 (c; 0 min). After 2 min of incubation with

cofilin-1, SEPT9-induced F-actin bundles still remain intact (c; 2 min), and even after

30 min of incubation, F-actin bundles still persist (c; 30 min). (d–f) High-speed

sedimentation assay of F-actin in the presence of SEPT9 domains and cofilin-1. Actual

gels with the corresponding bar graphs (d–f) show that, in the presence of cofilin-1, most

of the actin is found in the supernatant (d), while in the presence of full-size SEPT9 (d),

NTD (e) or the B-domain (f), more than half of the actin is recovered in the pellet. In the

presence of the A-domain, most of the actin remains in the supernatant (f).

113

Figure 6.5. Overall 3D reconstruction of F-actin decorated with the NTD suggest

the sites of SEPT9 binding to the actin filament. Segments of actin filaments

decorated with the NTD of SEPT9 were reconstructed using IHRSR approach starting

from a solid cylinder. The resultant reconstruction possessed three additional densities

on the surface of F-actin. The first one is at the front interface between SD1 and SD2 of

the two adjacent actin protomers (red arrow). The second density is at the side of SD1

(magenta arrow), while the third one is on the top of SD4 (cyan arrow)

114

Figure 6.6. Convergence of structural classes validates model-based

crosscorrelation sorting. Every structural class was independently reconstructed using

IHRSR algorithm starting from either a solid featureless cylinder (purple) or 3D

reconstruction obtained from another structural class. 3D reconstructions of mode 1,

mode 2 and mode 3 are shown in yellow, cyan and green, respectively. Each class

converged to the same structural solution regardless of the starting model.

115

Figure 6.7. Reference free 2D averages of occupied classes. SPIDER1 AP SR

routine was used to generate reference free 2D averages of the three structural modes

recovered in the NTDdecorated actin filaments. Red arrows indicate visible differences

in the SEPT9-NTD positions.

116

Figure 6.8. Comparison of the frequencies of structural classes upon cross-

correlation sorting of F-actin segments in the presence or absence of the A-

domain of SEPT9. We performed cross-correlation sorting of pure F-actin segments

and F-actin segments in the presence of the A-domain using the same set of references

that was employed in the sorting of the NTD-decorated F-actin segments. Comparison of

the classes’ frequencies does not reveal any difference between the two data sets and

suggests that the small number of F-actin segments assigned to the occupied classes in

the presence of A-domain is not due to the decoration by the A-domain, but rather

reflects the margin of error in the sorting procedure.

117

Figure 6.9. Full length SEPT9 and SEPT9 B-domain do not significantly affect F-

actin bundling over concentration ranges that inhibit myosin activity. (a - b)

Coomassie-stained SDS-PAGE gel shows the supernatant (S) and pellet (P) fractions

from a low-speed actin sedimentation assays in the absence and presence of

recombinant full length SEPT9 (a) or Bdomain (b). (c - d) Bar graph shows the

percentage of F-actin in the supernatant and pellet fractions in the absence and

presence of SEPT9 (c) or B-domain (d). Error bars indicate the lowest and highest

values from three independent experiments. Note that the percentage of pelleted actin is

similar between 1 and 3 µM of full length SEPT9, while this concentration increase

results in a 2-3-fold decrease in myosin ATPase activity (Fig. 3d). The percentage of

pelleted actin is also similar in the range of 3 – 12 µM of B-domain, which causes >5-fold

decrease in myosin ATPase activity (Fig. 3d).

118

Figure 6.10. Cosedimentation assay of full length SEPT9 with F-actin. (a)

Coomassie-stained gels show the pellet and supernatant fractions (equal volumes) from

a representative high-speed sedimentation assay. SEPT9 (1 µM) was incubated with

increasing concentrations of F-actin for 30 min at room temperature and was centrifuged

at 200,000 g. (b) Binding curve shows the fraction of F-actin-bound SEPT9 as a function

of increasing concentrations of F-actin. Data were fitted in GraphPad with one site

specific binding curve and an estimated Kd of 2.23 ± 0.60 µM was derived based on

three independent experiments.

119

Figure 6.11. Resolution determination. Each structural class was split in two non-

overlapping sets and the IHRSR procedure was used on each set. The correlation

between the corresponding Fourier shells of the two resultant volumes was calculated

for each data set.

120 CHAPTER 7. Septins promote macropinosome maturation and traffic to the

lysosome by facilitating membrane fusion

Abstract

Macropinocytosis, the internalization of extracellular fluid and material by plasma

membrane ruffles, is critical for antigen presentation, cell metabolism and signaling.

Macropinosomes mature through homotypic and heterotypic fusion with endosomes,

and ultimately merge with lysosomes. The molecular underpinnings of this clathrin-

independent endocytic pathway are largely unknown. Here, we show that the

filamentous septin GTPases associate preferentially with maturing macropinosomes in a

phosphatidylinositol 3,5-bisphosphate-dependent manner, and localize to their

contact/fusion sites with macropinosomes/endosomes. Septin knock-down results in

large clusters of hemifused macropinosomes, which persist longer and exhibit less

fusion events. Septin depletion and over-expression down-regulate and enhance,

respectively, the delivery of fluid-phase cargo to lysosomes, without affecting Rab5 and

Rab7 recruitment to macropinosomes/endosomes. In vitro reconstitution assays show

that fusion of macropinosomes/endosomes is abrogated by septin immunodepletion and

function-blocking antibodies, and is induced by recombinant septins in the absence of

cytosol and polymerized actin. Thus, septins regulate fluid phase cargo traffic to

lysosomes by promoting macropinosome maturation and fusion with

endosomes/lysosomes.

Introduction

Macropinocytosis is a clathrin-independent pathway of endocytosis that

originates from the closure of plasma membrane ruffles into large vesicles, which

121 internalize extracellular fluid and particles (Kerr and Teasdale, 2009; Swanson, 2008).

Macropinocytosis is upregulated by growth factors and oncogenes, which stimulate the

uptake of nutrients for cell proliferation and growth (White, 2013). Many bacterial and

viral pathogens invade cells by hijacking macropinocytosis, which is critical for the

antigen presentation (de Carvalho et al., 2015; Lim and Gleeson, 2011; Mercer and

Helenius, 2012). By modulating the internalization of plasma membrane receptors and

adhesion molecules, macropinocytosis is also implicated in cell signaling, adhesion and

motility (Bryant et al., 2007; Gu et al., 2011; Schmees et al., 2012).

Macropinosomes traffic to lysosomes while undergoing a process of maturation,

which is characterized by size reduction concomitant with membrane tubulation and

fusion with other macropinosomes and early/late endosomes (Bright et al., 2005;

Racoosin and Swanson, 1993). Macropinosome maturation is accompanied by changes

in phoshoinositide and Rab GTPase content (Levin et al., 2015). Nascent

macropinosomes become enriched with phopshatidylinositol 3-phosphate (PI3P) and

Rab5 (Feliciano et al., 2011; Yoshida et al., 2009), while maturing macropinosomes

contain phopshatidylinositol 3,5-biphosphate (PI(3,5)P2) and Rab7 (Kerr et al., 2006;

Kerr et al., 2010). Despite our knowledge of these transitions, the molecular basis and

regulation of the membrane fusion events that underlie macropinosome maturation and

merging with the lysosome remain unknown.

Septins are filamentous heteromeric GTPases that associate with cell

membranes and the cytoskeleton (Fung et al., 2014; Mostowy and Cossart, 2012).

Plasma membrane septins are essential for the maintenance of diffusion barriers and

modulate exocytosis (Bridges and Gladfelter, 2015; Caudron and Barral, 2009). Much

less is known about how septins function in the endomembrane system of organelles

and their communication. Septin mRNAs and proteins associate with endosomes

122 (Baumann et al., 2014; Baust et al., 2008) and are implicated in multivesicular body

formation (Traikov et al., 2014). However, how septins function in endocytic membrane

traffic is not known.

Materials and methods

Plasmids

Mouse SEPT2 was inserted into N1-YFP (Spiliotis et al., 2005). Mouse SEPT2

was inserted into N1-GFP (Bowen et al., 2011). PM-mCherry was constructed by

inserting the membrane-binding domain of lyn kinase in pcDNA 3.1-mCherry (Benjamin

et al., 2010). Rab5A was inserted into C1-GFP using the HindIII and SalI restriction

sites, and Rab7A was inserted into C1-GFP using the Xho1 and KpnI restriction sites.

Rab5B-mCherry and Rab7A-mCherry were purchased from addgene. Rat SEPT7v2 and

human SEPT6v3 were inserted into the pET-28a(+) vector, which expresses in tandem

His-SEPT7 and an untagged SEPT6 (Bai et al., 2013). Mouse SEPT2 was inserted into

pET-15b and expresses an untagged SEPT2 (Bai et al., 2013).

For experiments using shRNAs, the non-targeting control (5′-

TGAGTTCACACTAATGGTGGTCG-3′) was inserted in pG-SUPER-GFP (Bowen et al.,

2011) and pSUPER-mCherry (Dolat et al., 2014b). To target canine SEPT2, the shRNA

#1 (5′-CCTTAGACGTCGCATTCATGAAA-3′) was inserted in pSUPER-mCherry (Dolat

et al., 2014b) and SEPT2 shRNA #2 (5′-GCAACTACAAGCCCGAAGAATAA-3′) was

inserted in pG-SUPER-GFP. Human SEPT2 was targeted using the SEPT2 shRNA 5′-

AAGGTGAATATTGTGCCTGTC-3′, which was inserted in pG-SUPER-GFP (Bowen et

al., 2011). For siRNA experiments (Fig. 5), the SEPT2 siRNA oligonucleotides

(5’AAGGUGAAUAUUGUGCCUGUC-3) and mock GFP siRNA oligonucleotides (5’-

123 GGCUACGUCCAGGAGCGCACC-3’) were purchased from Dharmacon Research

(LaFayette, CO).

Tissue culture and transfections

MDCK II/G cells and the stable cell lines MDCK-SEPT2-YFP (CMV promoter;

N1-YFP) (Spiliotis et al., 2005) and MDCK-PM-mCherry (CMV promoter; pcDNA3.1)

were cultured in low glucose DME with 1 g/L NaHCO3, 10% FBS (Cell Generation), and

1% PSK (penicillin, streptomycin and kanamycin) at 37 ˚C with 5% CO2. HT1080 cells

(CCL-121) were purchased from ATCC and cultured in high glucose DME with 1 g/L

NaHCO3, 10% FBS (Cell Generation), and 1% PSK (penicillin, streptomycin and

kanamycin) at 37 ˚C with 5% CO2. Cells were plated on No. 1.5 glass coverslips (VWR),

which were pre-coated with 10 µg/mL type I collagen (PurCol; BioMatrix) in 0.1% acetic

acid for five minutes, and subsequently were dried and rinsed twice with media before

cell plating. Cells were transfected with 1.5 µg plasmid or co-transfected with 1.0 µg of

one plasmid and 1.5 µg of the second plasmid using lipofectamine 2000 (Invitrogen).

Dextran uptake and pulse-chase assays

Cells were cultured in low-serum media (0.5% FBS) overnight and switched to

media with 10% FBS to stimulate macropinocytosis for the duration of incubations and

pulse-chase treatments with fluorescent dextrans. Pulse-chase experiments were

performed by incubating cells with media containing 10% FBS and 0.2 mg/mL FITC- or

TR-dextran (10,000 MW, ThermoFisher Scientific) for 5 min and then washing three

times and incubating cells in dextran-free media for the indicated times. In Fig. 2C, cells

were incubated with 800 nM YM201636 (SelleckChem, Inc.) or vehicle control (DMSO)

124 for 2 h in low-serum media and then stimulated with media containing 10% FBS and 0.2

mg/mL TR-dextran with YM201636 or DMSO for 10 min.

Immunofluorescence microscopy

Cells were fixed with warm (37 ˚C) 2% paraformaldehyde (PFA) in PBS

(phosphate-buffered saline) and simultaneously permeabilized and blocked with 2%

BSA (bovine serum albumin; Sigma) and 0.1% saponin (Calbiochem) for 30 min. In Fig.

1A, cells were extracted prior to fixation by incubating for 30 s with warm (37 ˚C)

cytoskeleton buffer (CB) containing 10 mM MES (pH 6.1), 138 mM KCl, 3 mM MgCl2, 2

mM EGTA, 320 mM sucrose and 0.1% Triton X-100. Cells were subsequently rinsed

twice with CB and fixed with 2% PFA in CB.

Rabbit antibodies against SEPT2 (N5N; a gift from M. Kinoshita, Nagoya

University, Nagoya, Japan) and Rab5 (Stressgen), and mouse antibodies against Rab7A

(Abcam), LAMP1 (H4A3, Developmental Studies Hybridoma Bank; University of Iowa),

Sec6 (Exoc3; 10C2), Sec8 (Exoc4; 9H5), Exo70 (Exoc7; 13F3), gifts from C. Yeaman

(University of Iowa), were diluted in PBS containing 2% BSA and 0.1% saponin.

Secondary donkey Alexa Fluor 488-, AlexaFluor 594- or Alexa Fluor 647-conjugated

F(ab’) to mouse or rabbit IgG (Jackson ImmunoResearch Laboratories, Inc.) were

diluted in PBS containing 2% BSA and 0.1% saponin. Cells were stained for F-actin

using phalloidin conjugated to Alex Fluor 555 (ActiStain; Cytoskeleton, Inc.). Coverslips

were mounted on coverslides using Vectashield (Vector Laboratories).

Slides were imaged using a confocal laser-scanning microscope (FluoView 1000;

Olympus) equipped with three Hamamatsu R7862 photomultiplier tubes (PTMs) and

multi-line Argon ion (458 nm, 488 nm, 515 nm), Helium-Neon (543 nm) and diode 401

nm and 635 nm lasers. Images were acquired using 60x/1.42 NA oil objective and the

125 FV10-ASW 02.01 software. 3D rendering of confocal stacks was performed in the

Volocity software (Perkin Elmer).

Time-lapse imaging

All time-lapse microscopy experiments were performed at 37˚C using DME

without Phenol red supplemented with 10% FBS, 1% PSK and 20 mM Hepes. In Fig. 1

and 3A,, PM-mCherry dynamics were imaged with 3D deconvolution wide-field

microscopy using an inverted microscope (DeltaVision OMX v4; GE Healthcare)

equipped with a motorized stage, stage-tope incubator with temperature controller, and

three pco.edge sCMOS cameras (custom version). 3D stacks (1 – 2 µm; 0.125 nm

slices) were acquired every 10-20s using a 60x/1.49 NA objective lens (Olympus) and

were deconvolved in the softWoRx 6.1.1 software. In Fig. 3B-D and 3H, macropinosome

dynamics were imaged using an inverted microscope (IX81; Olympus) equipped with a

motorized stage (ProScanII; Prior), an OrcaR2 CCD camera (Hamamatsu Photonics), a

custom-built stage-top chamber and temperature controller (Air-Therm ATX; World

Precision Instruments) and the Slidebook 6.0 software. Fluorescent images were

acquired every 5 s for 8 min with a Plan-Apochromat 60x/1.40 NA objective. Time-lapse

videos were rendered in Photoshop (Adobe).

Protein expression and purification

Recombinant SEPT2/6/7 was expressed by co-transforming Escherichia coli

BL21(DE3) cells with a kanamycin-resistant, bicistronic pET-28a(+) that expresses His-

SEPT7 and SEPT6, and an ampicillin-resistant pET-15b(+) plasmid that expresses an

untagged SEPT2. Bacterial cultures were grown to 0.6 OD and induced with 0.5 mM

IPTG for 16 hours at 18 ˚C. Cultures were centrifuged (4,000 g) for 10 min at 4 ˚C, lysed

126 in buffer containing 50 mM Tris pH 7.5, 150 mM NaCl, 1 mM PMSF, 1 mM DTT, 10 mM

imidazole, and 10% glycerol using a French pressure cell at 1,280 psi. Cell lysates were

centrifuged (14,000 g) for 30 min at 4 ˚C and passed through a 0.45 µm filter before

applying to a gravity flow column with Ni-NTA beads. The columns were washed with

buffer containing 50 mM Tris pH 8.0, 300 mM NaCl, 1 mM DTT, 20 mM imidazole, and

10% glycerol, and SEPT2/6/7 complexes were eluted using buffer containing 50 mM Tris

pH 8.0, 150 mM NaCl, 10% glycerol, and 250 mM imidazole. Protein preps were

dialyzed overnight in dialysis buffer (50 mM Tris, pH 8.0, 150 mM NaCl, 1 mM DTT, and

10% glycerol).

Liposome flotation assays

All lipids were purchased from Avanti Polar Lipids, except PI(4,5)P2 diC8

(Echelon Biosciences; p-4508). L-α-phosphatidylcholine (840051) and L-α-

phosphatidylinositol (840042) were diluted in chloroform. PI(3,4,5)P3 (850156),

PI(3,5)P2 (850154), PI(3)P (850150), PI(4)P (850151), and PI(5)P (850152) were diluted

in a solution made of chloroform, methanol and water (20:9:1). PI(4,5)P2 was diluted in

water. To generate small unilamellar vesicles, lipids were mixed to a final concentration

of 75 %mol L-α-phosphatidylcholine, 20 %mol L-α-phosphatidylinositol, and 5 %mol of

each phosphoinositide. Lipid mixtures were dried completely under a nitrogen stream,

rehydrated in liposome buffer (50 mM Hepes pH 7.2, 50 mM KCl) for 30 min at 25 ˚C,

freeze/thawed five times, and passed through a mini-extruder using 0.1 µm pore filter at

least 12 times to isolate small unilamellar vesicles (SUVs). SUVs were stored at 4˚C and

used within 1-2 days.

To test for septin binding, 0.5 mM SUVs were mixed with 0.5 µM recombinant

SEPT2/6/7 in 50 µl liposome buffer containing 1 mM MgCl2 and incubated with rotation

127 at 25 ˚C for 1 h. Reactions were mixed with 200 µl liposome buffer with sucrose (75%)

and placed at the bottom of a polycarbonate tube (Beckman Coulter). Liposome buffer

(150 µl) with 25% sucrose was gently overlaid and on top of this layer, 50 µL of liposome

buffer was gently placed. Samples were centrifuged (220,000 g) in a SW60 Ti rotor

(Beckman Coulter) for 1 h at 20 ˚C. Equal volumes of the top and bottom fractions were

collected with a thin pipette tip, run on a 10% SDS-PAGE gel and stained with

Coomassie Brilliant Blue. Gels were scanned using the Odyssey imaging system (LI-

COR Biosciences).

In vitro fusion assay

Reconstitution of endosome fusion was performed in a cell-free fluorescence-

based assay as previously described (Barysch et al., 2010). MDCK cells were cultured

on 15 cm plates to 80% confluence, trypsinized and centrifuged (300 g) for 5 min. Cells

were resuspended in cold PBS containing 5 mg/mL BSA (PBS-BSA) and centrifuged

(250 g) for 5 min at 4˚C. The cell pellet was resuspended in cold internalization buffer

(0.2 mg/mL glucose in OPTI-MEM) and centrifuged (250 g) for 5 min at 4˚C. The

supernatant was discarded and the pellets were incubated at 37˚C for 5 min before

resuspending in warm internalization buffer with either FITC- or TR-dextran (0.5 mg/ml)

and incubating for an additional 5 min at 37 ˚C. Cells were transferred to ice and washed

three times with cold PBS-BSA and once with homogenization buffer (250 mM sucrose,

3 mM imidazole pH 7.4) while centrifuging (250 g) for 5 min at 4 ˚C in between washes.

The cell pellet was resuspended in 1 ml homogenization buffer containing protease

inhibitors (Calbiochem), passed through a ball-bearing homogenizer 20 times, and

centrifuged (1,200 g) for 15 min at 4 ˚C. The post-nuclear supernatant (PNS) was

recovered and protein concentration was determined using a Bradford assay (Thermo

128 Scientific). Small aliquots of the post-nuclear supernatants were flash frozen in liquid

nitrogen and stored in -80 ˚C. Canine kidney cytosol (gift from Dr. James Nelson) was

isolated as previously described (Davis et al., 1974). Canine kidney cortex was washed

in cold PBS, cut into 1 cm cubes and homogenized using a Waring blender in a PBS

containing 250 mM sucrose. The homogenate was filtered, centrifuged (100,000 g) to

remove cell membranes, and aliquots of the supernatant were drop flash frozen in liquid

nitrogen and stored at -80˚C.

Fusion reactions were assembled by mixing 5 µg canine kidney cytosol with 0.1

µg of post-nuclear extracts with FITC- and TR-dextran-labeled

macropinosomes/endosomes in homogenization buffer (50 µl) with protease inhibitors

and with or without 3.3 mM ATP. All reactions were incubated at 37 ˚C for 45 min. To

inhibit SEPT2 function, the cytosol was pre-incubated in homogenization buffer

containing 0.5 mg/ml rabbit anti-SEPT2 (N5N) or rabbit IgG (Sigma) for 1 h at 4 ˚C.

SEPT2 was immunodepleted from the cytosol as previous described (Pagano et al.,

2004). Cytosol was diluted to 1 mg/ml in buffer containing 20 mM Hepes pH 7.2, 90 mM

potassium acetate and 2 mM magnesium acetate with protease inhibitors and

subsequently incubated with 5 µg rabbit anti-SEPT2 or rabbit IgG overnight with rotation

at 4 ˚C. Protein A beads (Thermo Scientific) were blocked with 2% BSA overnight,

washed five times with homogenization buffer, and added to the cytosol for 3 hours at

4˚C. Samples were centrifuged (1,000 rpm) for 5 minutes at 4˚C and the supernatant

was collected. SEPT2 protein levels in the cytosol samples were assessed by western

blots.

To assess fusion microscopically, reactions (30 µl) were floated into chambers

created by immobilizing a glass coverslip on a slide with two strips of a double-side tape;

prior to mounting, coverslips were cleaned with 70% ethanol and flame-dried. Chambers

129 were washed with homogenization buffer prior to adding each reaction mix and sealing

with vacuum grease. Slides were centrifuged (300 g) upside-down for 10 min at 25 ˚C

using a swing-bucket rotor with microtiter adapters. Samples were imaged using an

inverted Zeiss AxioObserver Z1 wide-field microscope equipped with a Hamamatsu

Orca-R2 digital CCD camera, X-cite 120 metal halide lamp and a motorized z-drive.

Images were acquired using a 63X/1.4 NA objective and the Slidebook 6.0 software

(Intelligent Imaging Innovations).

Western blots

Cells were lysed in buffer containing 50 mM Hepes, pH 7.4, 100 mM NaCl, 0.5%

Triton X-100, 1 mM MgCl2, 1 mM PMSF and protease inhibitors. Samples were

centrifuged at 8,000 rpm for 10 min at 4˚C and the supernatants were collected. Equal

volumes were resolved with a 10% SDS-PAGE gel, transferred to a nitrocellulose

membrane at 100V for 1 h at 4˚C. Membranes were blocked with 5% dried milk (w/v) in

PBS containing 0.1% Tween-20. A rabbit antibody against SEPT2 (Sigma; 1:5000) and

a secondary anti-rabbit antibody (LiCor) conjugated to infrared dye was diluted in PBS

containing 2% BSA, 0.1% Tween-20, and 0.025% sodium azide, and sequentially

incubated on the membrane before scanning with the Odyssey imaging system (LI-

COR).

In Fig. 5, equal volumes of canine kidney cytosol samples were resolved on a

10% SDS-PAGE gel and transferred to a nitrocellulose membrane. Membranes were

blocked in 5% dried milk (w/v) in PBS containing 0.1% Tween-20. Rabbit antibody

against SEPT2 (Sigma; 1:5000), mouse anti-actin (Sigma, 1:5000), and anti-rabbit or –

mouse secondary antibodies (LiCor) conjugated to infrared dyes were diluted in PBS

containing 2% BSA, 0.1% Tween-20, and 0.025% sodium azide, and sequentially

130 incubated on each membrane before scanning with the Odyssey imaging system (LI-

COR).

Image analysis and quantifications

Image analysis was performed with the Slidebook 6.0 software (Intelligent Imaging

Innovations) unless otherwise noted. In Fig. 1B, the peripheral lamellar regions of cells

were masked by segmentation of F-actin fluorescence and within these regions, the sum

of SEPT2 fluorescence intensity per µm2 was quantified in each cell. In Fig. 2, the

percent of dextran particles with SEPT2 was calculated by creating dextran and SEPT2

masks using fluorescence intensity segmentations. SEPT2 and dextran masks were

subtracted to create a new artificial channel with masks that corresponds to all dextran

fluorescence that overlaps with SEPT2. The particles of this overlay channel were

automatically counted and their number was divided by the total number of dextran

particles to derive the percentage of dextran that colocalizes with SEPT2. The same

approach was used to derive the percentage of Rab5- and Rab7-mCherry compartments

with SEPT2 in peripheral lamellae (Fig. S1) and the percentage of dextran particles with

Rab5, Rab7 and LAMP1 (Fig. 3 and 4); LAMP1 background fluorescence was

subtracted by Gaussian smoothing. In Fig. 2B-D, time-lapse movies of PM-mCherry of

identical capture intervals (5 s) and duration (8 min) were analyzed after applying the 2-

D Laplacian of Gaussian function, which improves the clarity of membrane vacuoles and

tubules. The formation of nascent macropinosomes from membrane vesicles was

tracked manually and clusters of three or more macropinosomes per cell were scored

(Fig. 3B). Macropinosome lifetime was quantified by measuring the time that it took for a

nascent macropinosome to disappear or merge with another membrane vacuole or

tubule (Fig. 3C). The number of nascent macropinosomes that fused with another

membrane compartment was calculated as percentage of total macropinosomes

131 tracked. Number of fusion events per min was the inverse of the time that it took for a

nascent macropinosome to undergo a fusion event (Fig. 3D). In Fig. 3E, 2 µm-long line

masks were drawn along the hemifused membrane contact of two macropinocytic

vacuoles and along their non-contacting perimeters. The intensity profiles of each line

scan was exported and plotted in Excel. In Fig. 3F-G and S2, the number and size of

dextran particles were quantified in Image J (NIH) by generating masks of dextran

fluorescence intensity, which were gated by size (0.2 – 20 µm2) as previously done by

others (Wang et al., 2014). The rates of lysosomal delivery of dextran corresponded to

the slope of the lines derived from a least squares linear regression fit to the 5, 10 and

30 min points of the plot in Fig. 4C. In vitro fusion (Fig. 5) was quantified by generating

fluorescence segmentation masks that correspond to FITC- and TR-dextran-containing

macropinosomes/endosomes. The masks were subsequently subtracted from each

other to identify macropinosomes/endosomes with overlapping fluorescence. The

number of these new masks was automatically calculated and divided by the total

number of macropinosomes/endosomes, which was calculated by subtracting the

number of masks with FITC/TR overlap from the total number of FITC- and TR-labeled

masks.

Statistical analyses

All datasets were analyzed using the Shapiro and Levene tests, respectively, for

normality and equal variance in the R software, and an unpaired student’s t test used to

derive p values (95% confidence intervals) in the Excel software. All values represent

the mean ± SEM.

132 Results and Discussion

Septins associate preferentially with maturing macropinosomes in a PI(3,5)P2-

dependent manner

We previously identified a network of septins fibers in the leading lamellae of

migrating MDCK epithelia (Dolat et al., 2014b). This network is interwoven with actin

stress fibers, but it is reduced upon mild detergent extraction, indicating that a subset of

septins is membrane-bound (Fig. 1, A and B). Given that membrane ruffling activity is

high in cellular lamellae, we imaged the localization of septins in MDCK cells that stably

express PM-mCherry (Corbett-Nelson et al., 2006), a plasmalemmal marker that marks

membrane ruffles, macropinocytic vacuoles and vesicular-tubular endosomes (Fig. S1, A

and B). We stained for endogenous SEPT2 as a proxy for the lamellar network of septin

fibers, which consist of SEPT2/6/7/9 (Dolat et al., 2014b). SEPT2 was largely absent

from membrane ruffles, but localized at the contact sites and membranes of PM-

mCherry-labeled vacuoles (Fig. 1, C and D). Consistent with septin association with

macropinosomes, SEPT2 localized to the periphery and in between compartments that

contained PM-mCherry and dextran, a fluid phase cargo marker that enters cells by

macropinocytosis (Fig. 1E). Time-lapse microscopy of PM-mCherry and SEPT2-GFP

showed that SEPT2 accumulates at the fusion sites of macropinosomes (Fig. 1F, Video

1); SEPT2 was present on 87% ± 12% of macropinosome contact sites (n = 11 cells).

To examine whether SEPT2 associates with early or late stage macropinosomes,

we performed a pulse-chase experiment with fluorescent dextran. SEPT2 co-localization

with dextran peaked after 5-10 min of internalization and diminished after 20-30 min (Fig.

2A), when dextran accumulates in the lysosome (see Fig. 4C). Given that early

macropinosomes contain Rab5 while maturing macropinosomes become enriched with

Rab7 before fusing with lysosomes (Feliciano et al., 2011; Kerr et al., 2006; Kerr et al.,

133 2010), we analyzed SEPT2 localization in cells that expressed Rab5- and Rab7-

mCherry (Fig. S1, C-E). In peripheral lamellae, SEPT2 was present on 64% ± 4% of

Rab7-positive compartments compared to 21% ± 2% of Rab5 (Fig. S1E). SEPT2 was

also present on 72 ± 4% (n = 15 cells) of peripheral vacuoles labeled with the PI(3,5)P2-

binding ML1Nx2-GFP probe (Li et al., 2013), indicating that septins associate with

mature PI(3,5)P2-positive macropinosomes (Fig. 2B). Treatment of MDCK cells with

YM201636, an inhibitor of the FYVE finger containing phosphoinositide kinase (PIKfyve)

that converts PI(3)P to PI(3,5)P2 (Jefferies et al., 2008), reduced the intensity of the

lamellar septin network and SEPT2 localization on dextran-containing macropinosomes

(Fig. 2, C-E). Liposome floatation assays showed that SEPT2/6/7 associates

preferentially with PI(3,5)P2-containing membranes (Fig. 2, F and G). In contrast to

previous results with giant unilamellar vesicles (GUVs) (Tanaka-Takiguchi et al., 2009),

SEPT2/6/7 did not exhibit significant binding to PI(4,5)P2. Given that septin-lipid binding

is sensitive to membrane curvature (Bridges et al., 2016), this discrepancy may arise

from differences in the diameter of liposomes, which is 50-to-200-fold smaller than

GUVs. Our results nevertheless indicate that septins associate preferentially with mature

Rab7- and PI(3,5)P2-positive macropinosomes.

Septin depletion impedes the maturation and turnover of macropinosomes, and

reduces macropinosome fusion events

We sought to determine how septins affect macropinosome formation and

dynamics by imaging PM-mCherry in live MDCK cells, which were treated with control or

SEPT2 shRNAs that deplete SEPT2, but also SEPT6, 7 and 9 (Dolat et al., 2014b). In

control cells, membrane ruffles gave rise to multiple macropinosomes, which fused into

single vacuoles or dissipated within seconds of formation (Fig. 3A, Video 3). In SEPT2-

134 depleted cells, membrane ruffling and macropinosome formation were not affected.

Strikingly, however, SEPT2 knock-down resulted in clusters of numerous

macropinosomes (Fig. 3A, Video 3). Clusters of three or more macropinosomes were

present in four-fold more cells upon SEPT2 depletion and the average lifetime of

nascent macropinosomes increased by 1.7-fold (Fig. 3, B and C). Notably, several

macropinosomes appeared docked with their membranes adjoining laterally (Fig. 3E).

In agreement with a slower macropinosome turnover, dextran uptake assays

showed a significant increase in the number and size of dextran particles in SEPT2-

depleted cells (Fig. 3, F and G). Notably, this increase in number and size persisted after

30 and 60 min of dextran internalization, indicating a defect in the macropinocytic traffic

of dextran (Fig. 3, F and G). Similar results were obtained with a different SEPT2 shRNA

(Fig. S2, A and B) and in human fibrosarcoma HT1080 cells (Fig. S2, C and D), which

have intense membrane ruffling activity due to the expression of an oncogenic mutant of

N-ras (Paterson et al., 1987). Expression of an shRNA-resistant SEPT2-YFP rescued

the phenotype, reversing the number and size of macropinosomes to control levels (Fig.

S2B).

Because membrane tubulation and fusion events underlie the size reduction of

maturing macropinosomes, we analyzed macropinosome membrane dynamics in

MDCK-PM-mCherry cells. Septin depletion did not alter the percentage (~40%) of

macropinosomes that formed membrane tubules and had no effect on their detachment

(Fig. 3H, Video 4). Membrane fusion, however, was severely affected. The percentage

of nascent macropinosomes that fused with another macropinosome was reduced from

74% to 25% (n = 9 cells). Moreover, the frequency of fusion events between nascent

macropinosomes and tubulo-vesicular endosomes decreased markedly (Fig. 3D). Thus,

135 septins are required for the membrane fusion events underlying the shrinkage and

turnover of macropinosomes.

Septins regulate fluid phase cargo traffic and delivery to the lysosome

Given that macropinosomes transport fluid phase cargo from the plasma

membrane to the lysosome, we sought to determine whether septins affect the endocytic

trafficking and lysosomal delivery of extracellular dextran. Using a pulse-chase

approach, MDCK cells were pulse-labeled with fluorescent dextran and its localization

with respect to Rab5, Rab7 and the lysosomal associated membrane protein 1 (LAMP1)

was analyzed after 5, 10 and 30 min, respectively. Septin depletion did not affect dextran

colocalization with Rab5 and Rab7 (Fig. S2, E-H). However, the percentage of dextran

particles that colocalized with LAMP1 decreased by >60% and this defect was rescued

by shRNA-resistant SEPT2-YFP (Fig. 4, A and B). These data suggest that septins are

required for the delivery of dextran to lysosomes, but do not affect dextran traffic to

Rab5- and Rab7-positive endosomes or the recruitment of Rab5/Rab7 to dextran-

containing macropinosomes.

To further test whether septins regulate the lysosomal delivery of fluid phase

cargo, we analyzed kinetically the accumulation of dextran in lysosomes. Dextran

colocalization with LAMP1 was quantified after 5, 15 and 30 min of internalization under

conditions of septin depletion and over-expression. Between 5 and 30 min of

internalization, SEPT2 depletion decreased the rate of lysosomal delivery from 2.34% to

1.02% of total dextran per min (Fig. 4C). Notably, the percentage of dextran colocalizing

with LAMP1 in SEPT2-depleted cells did not change between 30 and 60 min of

internalization (45 ± 6% vs. 50 ± 4%), which was indicative of a block in the traffic of fluid

phase cargo.

136 Next, we tested the effects of SEPT2 over-expression using stable MDCK cell

lines that express SEPT2-YFP at low and high levels (Fig. 4D). In the cells that express

SEPT2-YFP at low levels, dextran delivery to the lysosome was similar to cells treated

with control shRNAs (Fig. 4E). High levels of SEPT2-YFP expression, however,

increased the efficiency of dextran delivery to the lysosome (Fig. 4E). Thus, lysosomal

delivery of fluid phase cargo is dependent on the expression levels of SEPT2.

Septins promote fusion of macropinosomes with endosomes/lysosomes.

To test whether septins regulate lysosomal delivery of fluid phase cargo by

directly modulating membrane fusion, we performed dual color dextran mixing assays in

live cells and in vitro using a reconstitution assay of endosomal fusion (Barysch et al.,

2010).

In MDCK cells, we tested how septin knock-down affects the delivery of FITC-

dextran into a cohort of endosomes/lysosomes that were pre-loaded with TexasRed

(TR)-dextran. After a 5 min pulse/15 min chase with TR-dextran, we performed 5 min

pulse/15 min chase with FITC-dextran to assess its macropinocytic traffic into TR-

dextran/LAMP1-positive lysosomes. Septin depletion decreased FITC-dextran entry into

lysosomes with TR-dextran by 50% (Fig. 5, A and B).

Next, we performed a cell-free assay of macropinosome/endosome fusion using

post-nuclear extracts from two MDCK cell populations, which were labeled separately

with TR- and FITC-dextran; fusion was induced with canine kidney cytosol and ATP and

assayed microscopically by quantifying the fraction of endosomes with both TR- and

FITC-dextran (Fig. 5, C and D). Addition of a SEPT2 function-blocking antibody or

immunodepletion of SEPT2 from the cytosol abrogated fusion (Fig. 5, C-F). Surprisingly,

recombinant SEPT2/6/7 and ATP induced fusion in the absence of cytosol (Fig. 5G). Of

137 note, addition of recombinant SEPT2/6/7 to the cytosol did not increase fusion, possibly

due to saturation by cytosolic septins. Cytochalasin D, an actin depolymerizing

compound, had no effect on the endosome fusion induced by SEPT2/6/7 in the absence

of cytosol and did not alter fusion in the presence of cytosol (Fig. 5, D and G). Similarly,

fusion was not altered by the microtubule depolymerizing drug nocodazole. Thus,

septins can directly induce membrane fusion independently of F-actin and microtubules.

Taken together, our results show that septins promote the membrane fusion

events that underlie the maturation of macropinosomes and the delivery of fluid phase

cargo to lysosomes. This is the first evidence for a direct role of septins in clathrin-

independent endocytosis and the fusion of endomembranes. In the context of

exocytosis, septins interact with SNARE proteins and the exocyst complex (Beites et al.,

1999; Hsu et al., 1998; Ito et al., 2009; Taniguchi et al., 2007). The SNARE and vesicle

tethering complexes that mediate the fusion events of the macropinocytic pathway are

largely unknown. The exocyst Sec6/8 complex localizes to the endosomes of MDCKs

(Oztan et al., 2007), but septins did not colocalize with Sec6/8 or Exo70 at the contact

sites of macropinosomes/endosomes (Fig. S3, C-E). Septin knock-down also did not

affect the localization of Vps39 (Fig. S3, F and G), a component of the HOPS vesicle-

tethering complex (Balderhaar and Ungermann, 2013). Because septin depletion

increases dramatically the fraction of docked macropinosomes/endosomes (Fig. 2),

septins appear to play a distinct role in membrane fusion without affecting membrane

tethering and/or docking. Notably, this septin role is coupled to PI(3,5)P2, a low

abundance phosphoinositide that is specifically involved in the fusion of mature

macropinosomes/endosomes with lysosomes (Kerr et al., 2006; McCartney et al., 2014).

Thus, similar to the evolutionarily related Rab GTPases, septin GTPases function as

phosphoinositide effectors that catalyze SNARE-mediated fusion. Future studies will

138 determine whether this function involves the stabilization of specific SNARE complexes

and the induction or sensing of membrane curvature, which promotes lipid mixing

(Martens and McMahon, 2008).

Oncogenic proteins of the Ras family of GTPases are long known to up-regulate

membrane ruffling and macropinocytosis, which in turn increases nutrient uptake to

support the metabolic needs of growing cancers (Kimmelman, 2015; White, 2013).

Moreover, lysosomal delivery of fluid phase cargo (e.g., amino acids) is critical for the

activation of signaling pathways that regulate cell growth and division (Commisso et al.,

2013; Settembre et al., 2013). Septins are over-expressed in many cancers, but their

functions in cancer are poorly understood (Connolly et al., 2011a; Dolat et al., 2014a).

Because SEPT2 alone does not induce fusion in vitro (Fig. 5G) and SEPT2-YFP over-

expression does not alter the expression levels of SEPT6/7 (data not shown), we posit

that over-expression of a single septin (e.g., SEPT2) may promote the formation of

membrane-bound SEPT2/6/7 polymers, which could functionally support the elevated

macropinocytic activity of cancer cells. Future studies may explore how this novel

function of septins might be therapeutically targetable.

139

140 Figure 7.1. Septins localize to macropinocytic vacuoles and tubulo-vesicular

endosomes, and their contact/fusion sites. (A) Confocal images of MDCKs stained

for SEPT2 and F-actin with or without pre-extration with 0.1% Triton-X100 prior to

fixation. (B) Bar graph shows the sum intensity of SEPT2 per cell area (n = 15). (C-D)

3D-rendered confocal images of MDCK-PM-mCherry cells stained for endogenous

SEPT2. Arrows point to SEPT2 at the contact sites between macropinocytic vacuoles

(C) and tubulo-vesicular endosomes (D). (E) MDCK-PM-mCherry cells were incubated

with FITC-dextran and stained for SEPT2. Images show SEPT2 at the periphery and in

between (arrow) macropinosomes/endosomes. (F-G) MDCK-PM-mCherry cells were

transfected with SEPT2-GFP and imaged live with wide-field deconvolution microscopy.

Still frames show SEPT2 accumulation (arrows) at the site of fusion between two

macropinosomes (F) and the contacting membranes of a macropinosome and a tubular

endosome (G).

141

142 Figure 7.2. Septins associate preferentially with maturing macropinosomes in a

PI(3,5)P2-dependent manner. (A) MDCKs were pulsed with TR-dextran for 5 min,

chased for the indicated times, stained for SEPT2 and imaged with confocal microscopy.

Bar graph shows the percent of dextran-containing macropinosomes/endosomes with

SEPT2 (n = 12 cells). (B) Recombinant SEPT2/6/7 was mixed with liposomes of the

indicated phosphoinositides and centrifuged on a sucrose gradient. Coomassie-stained

gels show equal volumes from the top (liposome-bound SEPT2/6/7; B) and bottom

(unbound SEPT2/6/7; U) fractions. (C) Maximum intensity projections of confocal images

of MDCKs, which were incubated with DMSO (control) or YM201636 (800 nM) for 2 h,

and treated with TR-dextran for 10 min before fixing and staining for SEPT2. Insets show

outlined areas in higher magnification. (D) Bar graph shows the fraction of TR-dextran

puncta (>0.2 µm2) with SEPT2 (n = 20 cells).

143

144 Figure 7.3. Septin depletion decreases macropinocytic fusion events and hinders

macropinosome maturation and turnover. (A) Time-lapse frames from widefield

deconvolution microscopy show the dynamics of PM-mCherry-labeled macropinosomes

in MDCK cells treated with control and SEPT2 shRNAs. (B) Control (n=41) and SEPT2-

depleted MDCK-PM-mCherry cells (n=31) were imaged live every 5 s for 8 min. Bar

graph shows percent cells with a cluster of three or more macropinosomes formed from

a lamellar ruffle. (C - D) Bar graphs show the lifetime (C) and fusion events (D) of

nascent macropinosomes (n = 24-27). (E) Quantification of PM-mCherry fluorescence

along the lateral hemi-fused contact (1) and non-contacting segments (2, 3) of two

macropinosomes from a SEPT2-depleted cell. (F-G) MDCKs were incubated with FITC-

dextran for the indicated times. Bar graphs show the number of FITC-dextran-containing

macropinosomes/endosomes (F) and their average size (G) per cell (n=18). (H) Still

frames show the formation and detachment of PM-mCherry-labeled membrane tubules

(arrows) in live MDCKs.

145

146 Figure 7.4. Septins regulate the lysosomal delivery of fluid phase cargo. (A)

MDCKs were transfected with GFP-expressing control or SEPT2 shRNAs for 48 h. After

a 5 min pulse/15 min chase with TR-dextran, cells were stained for LAMP1. Images

show maximum intensity projections of confocal image stacks. (B) Bar graph shows

percent of TR-dextran particles that colocalize with LAMP1 (n = 20 cells). (C) Plot shows

percent of TR-dextran with LAMP1 in control and SEPT2-depleted MDCKs (n=15), which

were pulsed with TR-dextran for 5 min and chased for the indicated times. (D) Lysates

from MDCKs that stably express SEPT2-YFP at low and high levels were analyzed by

SDS-PAGE and blotted for SEPT2. (E) Plot shows percent TR-dextran with LAMP1 in

MDCK cells (n=14) that express SEPT2-YFP at low and high levels. Cells were pulsed

with TR-dextran for 5 min and chased for the indicated times.

147

148 Figure 7.5. Septins are required for fusion of macropinosomes/endosomes with

lysosomes, and promote fusion directly in an in vitro reconstitution assay. (A)

MDCKs were treated with control or SEPT2 siRNA, and two rounds of 5 min pulse/15

min chase were performed with TR- and FITC-dextran, successively. Cells were stained

with anti-LAMP1 and imaged with confocal microscopy. Images show maximum intensity

projections of TR-dextran, LAMP1 and their areas of overlap (pseudo-colored in red) as

well as FITC-dextran (green), and its overlay with the pseudo TR-dextran/LAMP1

channel. (B) Bar graph shows percent TR-dextran/LAMP1 that contains FITC-dextran (n

= 18 cells). (C-D) MDCKs were separately incubated with TR- or FITC-labeled dextran

for 5 min, and post-nuclear supernatants (PNS) were mixed with canine kidney cytosol

with/without ATP, and with ATP plus cytochalasin D (cyto D; 10 µM), nocodazole (10

µM), control or anti-SEPT2 IgG. Images (C) show dextran-containing

macropinosomes/endosomes after incubation under the indicated conditions. Bar graph

(D) shows the fraction of fused macropinosomes/endosomes with both TR- and FITC-

labeled dextran (n = 15 images). (E) Bar graph shows the fraction of fused endosomes

after incubation of PNS with ATP and whole or SEPT2 immunodepleted (∆SEPT2)

cytosol (n = 15). (F) Gel shows equal volumes of whole and ∆SEPT2 cytosol blotted for

SEPT2 and actin. (G) Bar graphs show the fraction of fused endosomes after incubation

of PNS with/without recombinant SEPT2/6/7 in the presence/absence of 10 µM cyto D (n

= 15).

149

150 Figure 7.6. PM-mCherry and SEPT2 localization with respect to Rab5 and Rab7. (A-

B) MDCK-PM-mCherry cells were transfected with Rab5-GFP (A) and Rab7-GFP (B),

fixed and imaged by confocal microscopy. Images show 3D reconstructions from

confocal z-stacks. Outlined regions are shown in higher magnification. Arrows point to

vesicular endosomes with Rab5 or Rab7 and PM-mCherry. Arrowheads point to tubular

endosomes with Rab7 and PM-mCherry. (C-D) Confocal microscopy images show

MDCK cells that were transfected with Rab5 or Rab7-mCherry, fixed and stained for

endogenous SEPT2. Outlined regions are shown in higher magnification and 3D

reconstructions of the numbered endosomes (arrows) are shown in higher magnification.

(E) Bar graph shows the percentage of Rab5- and Rab7-mCherry-positive

compartments with SEPT2 (n = 17-18 cells).

151

152 Figure 7.7. SEPT2 knock-down increases macropinosome size and number, but

does not disrupt colocalization of macropinocytic cargo with Rab5 and Rab7. (A)

MDCKs were transfected with plasmids that express GFP or mCherry and non-targeting

or SEPT2 shRNAs (GFP/shRNA #1, mCherry/shRNA #2) for 48 h. Cells were fixed and

stained for endogenous SEPT2. Images show maximum intensity projections of SEPT2

fluorescence from 3D confocal microscopy images; insets show GFP/mCherry

fluorescence. (B) MDCKs were treated with control or SEPT2 shRNA # 2, and incubated

with FITC-dextran for 10 min. Bar graphs show the number of FITC-dextran-containing

macropinosomes/endosomes (puncta >0.2 m2) and their average size per cell (n = 21

cells). (C) HT1080 cells were transfected with plasmids that express GFP and control or

SEPT2 shRNA, and stained for endogenous SEPT2. Images show maximum intensity

projections of SEPT2 and GFP (insets) fluorescence. Bar graph shows the quantification

of SEPT2 intensity per cell area (n = 20 cells). (D) Control and SEPT2-depleted HT1080

cells were incubated with TR-dextran for 10 min. Bar graphs show the number of TR-

dextran-containing macropinosomes/endosomes (puncta >0.2 m2) and their average

size per cell (n = 20 cells). (E – F) MDCKs were transfected with GFP-expressing control

or SEPT2 shRNAs for 48 h. Cells were pulsed with TR-dextran for 5 min and stained for

endogenous Rab5 (E) and Rab7 (F) after a 5 and 10 min chase, respectively. Images

show maximum intensity projections of confocal image stacks. Outlined regions with

merged fluorescence are shown in higher magnification. Bar graphs show percent of TR-

dextran particles that colocalize with Rab5 (E) and Rab7 (F) (n = 20 cells).

153

154 Figure 7.8. SEPT2 does not colocalize with Sec6/8 and Exo70 at the contact sites of macropinosomes/endosomes. (A – C) Images show 3D reconstructions of confocal

image stacks of MDCK-PM-mCherry cells stained for endogenous SEPT2 and Exo70

(A), Sec8 (B) or Sec6 (C). Outlined regions are shown in higher magnification (scale

bars, 1 µm). Green arrows point to SEPT2 at the sites of contact between PM-mCherry-

labeled macropinosomes/endosomes, and blue arrowheads point to the localization of

Exo70, Sec8 or Sec6.

155

CHAPTER 8. Conclusions and future directions

This dissertation focuses on the function of septin GTPases in epithelial cell

migration and macropinocytosis. While septins have been shown previously to associate

with the actin cytoskeleton and cell membranes, the molecular requirements have

remained poorly understood. Moreover, septins are frequently overexpressed in

epithelial cancers but the role of septins in tumor cell growth and metastasis is unknown.

In this dissertation, I uncovered a mechanism by which septins interact directly with the

actin cytoskeleton and promote epithelial migration and invasion. In addition, I

discovered that septins associate with macropinosomes in a PI(3,5)P2-dependent

manner and regulate the delivery of macropinocytic cargo to endolysosomes, a

molecular pathway that promotes tumor cell growth.

Septins in epithelial cell migration

Previous studies have shown that septins exhibit an interdependent relationship

with the actin cytoskeleton (Kinoshita et al., 1997), but whether septins bind directly to

actin was unknown. In the first part of this thesis, I discovered that septins localize to the

contractile transverse actin fibers and their points of connection with radial actin fibers.

Septin knock-down abolishes the actin stress fiber network by reducing the number of

radial fibers and thinning the transverse actin fibers. Significantly, I found that SEPT9

directly crosslinks actin filaments and interferes with the binding of the myosin motor

domain in its low-affinity state and cofilin, an actin-severing protein. Thus, septins are

bona fide actin cross-linking proteins that may protect nascent actin filaments from

depolymerizing forces, such as contractile myosin filaments and actin-severing proteins,

during epithelial cell migration.

Because septins are overexpressed frequently in cancers, I assayed for changes

in septin expression during epithelia-to-mesenchymal (EMT) transition, a critical step in

156 tumor cell invasion. I found a significant increase in SEPT9 expression and tested the

effects of SEPT9 overexpression on the motile properties of epithelia undergoing EMT.

Indeed, SEPT9 overexpression promotes epithelial cell migration in 2D and 3D

environments, while SEPT9 knockdown or pharmacological targeting of septins inhibits

cell migration. Taken together, my findings indicate that septins promote epithelial cell

migration by directly cross-linking actin filaments and regulating the assembly and

organization of the contractile actin stress fiber network.

Future studies for septins in actin filament assembly and organization

In the leading edge of motile epithelia, the actin stress fiber network is

characterized by the assembly radial actin fibers, which are anchored to focal adhesions,

and transverse arc stress fibers, which are crosslinked by contractile myosin II filaments.

Through the linkage of these two stress fiber networks, forces are transduced to focal

adhesions to promote forward translocation of the cell. My data indicate that septins are

essential for the assembly of a cohesive stress fiber network and may regulate force

transduction during cell migration. To test this hypothesis, the forces exerted upon focal

adhesions can be measured experimentally using atomic force microscopy and gel

contraction assays, which provide direct and indirect readouts, respectively.

Furthermore, radial actin fibers link up with transverse actin fibers in an

orthogonal manner. My preliminary studies in vitro show that recombinant SEPT9 can

crosslink and promote the orthogonal organization of purified actin filaments (Appendix

1.1). Actin filaments alone assemble into short linear filaments (Appendix 1.1B).

However, when co-polymerized with recombinant SEPT9, actin filaments appear thicker,

which suggests that they are crosslinked, and are arranged in a more branched-like

organization with orthogonal linkages between filaments (Appendix 1.1C).

157

More work is necessary to better understand how SEPT9 can promote this

geometric arrangement of actin filaments. Because SEPT9 can bind to three different

sites on the actin filament, it is possible that SEPT9 promotes orthogonal interactions

when bound to a specific site. High-resolution electron microscopy studies can reveal

the precise localization of SEPT9 on orthogonal actin filaments and how its position on

the actin filament may promote this geometry. Moreover, SEPT9 is a component of the

SEPT2-6-7-9 octamer and likely functions within the cell as part of a molecular complex,

but how SEPT9 binds to actin when in an octameric filament has not been determined.

The SEPT2-6-7 crystal structure shows a hinged region at the SEPT2 interface, which

provides flexibility to the filament (Sirajuddin et al., 2007), and could promote the

branched and orthogonal arrangement of actin filaments.

In addition, I showed that SEPT6 promotes the recruitment of cortactin to the

lamellipodium, a branched network of actin filaments (Hu et al., 2012). Co-

polymerization of actin with the recombinant SEPT2-6-7 complex results in curved and

circular actin filaments (Mavrakis et al., 2014). Interestingly, the Arp2/3 complex

preferentially initiates nascent filament assembly on curved actin filaments (Risca et al.,

2012). Thus, septins may promote branched actin assembly by favoring geometrically

the spatial localization of Arp2/3 activity. Moreover, yeast two-hybrid experiments

identified that SEPT6 interacts with the Arp2 of the Arp2/3 complex (Nakahira et al.,

2010), but we were unable to identify this interaction in cells or biochemically. Future

studies involving co-polymerization of actin with the Arp2/3 complex are necessary to

better understand the role of septins in branched actin assembly. Here, testing the

effects of recombinant septins in an actin pyrene assay, which provides a fluorescence

readout for actin polymerization, can reveal whether septins promote directly actin

polymerization.

158 Future studies for septins in integrin trafficking

The assembly of radial actin fibers is initiated by the engagement of integrin

receptors with ECM proteins (Parsons et al., 2010). Cytosolic tails of engaged, or

activated, integrin receptors recruit actin nucleation factors that polymerize actin

filaments and promote focal adhesion assembly and maturation. Thus far, two integrin

trafficking pathways have been identified: fast Rab4-dependent recycling and slow

Rab11-dependent recycling (Caswell and Norman, 2006). Significantly, the trafficking of

integrins is essential for tumor cell migration and invasion.

Based on previous studies and my preliminary data, I hypothesize that septins

may also have a role in integrin trafficking. Notably, Rab11 localizes to the golgi complex

and regulates the delivery of exocytic vesicles to the plasma membrane (Chen et al.,

1998). Septins localize to post-golgi microtubules and regulate the trafficking of

intracellular vesicles to the plasma membrane. Thus, septins may also promote the

trafficking of Rab11-dependent integrins to the membrane where they can engage with

ECM proteins. Alternatively, my preliminary studies show that septins colocalize with

juxtanuclear Rab4-positive vesicles (Appendix 1.2) in Cos7 cells, a renal fibroblast-like

cell line. Future studies investigating the role of septins in intracellular trafficking of

integrin receptors may unveil a second mechanism by which septins regulate cell

migration.

Septins in tumor cell migration and metastasis

In Chapter 5, I tested the effects of septin depletion on epithelial cell migration in

3D collagen matrices, which better recapitulates the physiological environment.

Interestingly, septin depletion transitions the cell to a more rounded and protrusive

shape and slows significantly cell motility, suggesting that septins may have a role in the

mechanism that regulates the mode of tumor cell motility. Future studies using

159 biomimetic cell-derived matrices derived from fibroblasts, which better mimic the tumor

microenvironment, will provide a better context for studying tumor cell migration. In these

matrices, tumor cells can transition between three different modes of migration:

mesenchymal-like, amoeboid and lobopodial. While the molecular pathways that

regulate the transition between these modes is poorly understood, septins may play a

role in the spatial organization of the actomyosin cytoskeleton, whose contractile activity

regulates focal adhesion dynamics during mesenchymal-like motility and the protrusive

behavior in amoeboid and lobopodial motility.

Ultimately, it will be necessary to test how septin expression affects tumor cell

motility in vivo using a mouse tumor model, which can be visualized with intravital

imaging, and whether pharmacological targeting of septins inhibits metastasis.

Collectively, these studies will report on the efficacy septin-targeting therapeutics in

cancer.

Septins in membrane fusion dynamics

From yeast to mammals, membrane-associated septins are posited to restrict the

lateral diffusion of membrane proteins and promote the compartmentalization of the

plasma membrane (Caudron and Barral, 2009). Septins also modulate exocytic events

at the plasma membrane (Tokhtaeva et al., 2015), but how septins function in endocytic

membrane traffic and dynamics is little understood. In Chapter 7, I show that septins

associate with macropinosomes in a PI(3,5)P2-dependent manner. Interestingly,

PI(3,5)P2 is a low-abundance lipid that localizes exclusively to late endosomes and

lysosomes and promotes macropinosome fusion (Kerr et al., 2010). In agreement with

this finding, septins colocalize with Rab7-positive late endosomes and control the

delivery of macropinocytic cargo to lysosomes. Moreover, reconstitution of vesicle fusion

in vitro shows that septin inhibition and immunodepletion blocks vesicle fusion, while the

160 addition of recombinant septins promotes vesicle fusion. Thus, septins are an essential

component of the molecular machinery that regulates membrane fusion.

Vesicle fusion dynamics

Membrane fusion relies on the coordinated recruitment of vesicle adaptor

proteins, which can recognize specific lipid domains, and the assembly and disassembly

of SNARE proteins between two vesicles. In addition, the microtubule and actin

cytoskeleton play intricate roles in localizing and docking intracellular vesicles,

respectively, to promote vesicle fusion.

In Chapter 7, I reconstituted vesicle fusion in vitro using a crude post-nuclear

supernatant that contained fluorescently-labeled endosomes, whose lipid composition

and constituent adaptor protein population was unknown. However, I was able to rule

out the role of the actin and microtubule cytoskeleton in this assay through

pharmacological inhibition. Thus, I hypothesize that septins function mechanistically at

the level of SNARE-mediated membrane fusion.

Septins were reported to interact directly with SNAREs at the plasma membrane

and inhibit exocytosis (Beites et al., 1999). However, the SNAREs constitute a large

family of proteins that localize to specific intracellular membrane compartments. The

SNAREs that promote macropinosome fusion with lysosomes are unknown, but

syntaxin-7 is important for lysosome fusion and syntaxin-17 regulates autophagosome

fusion with lysosomes (Itakura et al., 2012; Jahn and Scheller, 2006).

At a mechanistic level, septins may function at the level of SNARE assembly

and/or endosomal membrane organization, which in turn can control SNARE dynamics.

To address the first question, biochemical analyses of septin-SNARE interactions and

vesicle fusion assays using purified liposomes reconstituted with SNARE proteins will

provide a better readout for septin functions during membrane fusion. In this assay,

161 fluorescently-labeled liposomes containing donor and acceptor SNAREs are mixed

together and assayed microscopically. If septins promote membrane fusion through a

SNARE-dependent mechanism, the addition of recombinant septins could result in more

vesicle fusion by promoting SNARE assembly, which tethers vesicles together, or

SNARE disassembly, which drives membrane fusion (Jahn and Scheller, 2006).

Alternatively, septins may function at the level of membrane organization. By

compartmentalizing the plasma membrane, septins are required for the organization of

PI(4,5)P2-rich domains that organize the Orai1 calcium channel at ER-PM contact sites

(Sharma et al., 2013a). Moreover, septins were reported recently to bind preferentially to

highly-curved membranes, which are present on the plasma membrane and

endomembranes. Interestingly, both lipid domain organization and membrane curvature

can regulate the clustering of SNAREs into distinct membrane domains, which is

necessary for vesicle docking and fusion. High resolution microscopy, such as

superresoluation microscopy, which can provide a lateral resolution of ~ 20 – 100 nm,

and electron microscopy can report on the localization of septins with respect to

SNAREs on the endosomal membrane. In addition, testing the effects of septin depletion

on endolysosomal SNAREs will report on their assembly and clustering into discrete

domains on the endomembrane.

Taken together, these experiments could uncover the mechanism by which

septins regulate vesicle fusion, and whether septins have a more universal role in the

delivery of cargo from other endocytic pathways to the lysosome.

162

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190

Appendix 1.1. In vitro actin crosslinking and co-polymerization of actin filaments with SEPT9

(A) Pre-polymerized rhodamine-labeled actin filaments were incubated with or

without recombinant full-length (FL) SEPT9. The length of actin filaments and the intensity per length were quantified and plotted.

(B) Polymerization of rhodamine-actin alone (C) Co-polymerization of rhodamine-actin with recombinant SEPT9-GFP.

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Appendix 1.2. SEPT9 colocalization with Rab4

Cos7 cells were fixed and stained for SEPT9 (green) and Rab4 (red). SEPT9 colocalizes with Rab4-positive vesicles. Bottom panel shows higher magnification of the region of interest (ROI).

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Vita Lee Dolat was born in Ledyard, CT, U.S.A on July 2, 1984. After receiving his Bachelors

of Science in Molecular and Cell Biology from the University of Connecticut, Lee worked

as a research technician at 454 Life Sciences, a high-throughput DNA sequencing

company, and Massachusetts General Hospital/Harvard Medical School. Lee conducted

his PhD studies in the laboratory of Elias T. Spiliotis, PhD in the Department of Biology

at Drexel University from 2010-2016.

Articles:

1) Lee Dolat and Elias T. Spiliotis. Septins promote macropinosome maturation and traffic to the lysosome by facilitating membrane fusion. In revision.

2) Elias T. Spiliotis and Lee Dolat. Priming for destruction: septins at the crossroads of mitochondrial fission and bacterial autophagy. EMBO Reports 17: 1-3, 2016.

3) Clayton Smith, Lee Dolat, Eva Forgacs, Elias T. Spiliotis and Vitold E. Galkin. Septin 9 exhibits polymorphic binding to F-actin and inhibits myosin and cofilin activity. Under Review in Journal of Molecular Biology 427: 3273 – 3284, 2015.

4) Lee Dolat, John L. Hunyara, Jonathan R. Bowen, Eva P. Karasmanis, Maha Elgawly, Vitold E. Galkin and Elias T. Spiliotis. Septins promote stress fiber-mediated focal adhesion maturation and renal epithelial motility. The Journal of Cell Biology 207: 225-235, 2014.

5) Lee Dolat, Qicong Hu, Elias T. Spiliotis. Septin functions in organ system physiology and pathology. Biological Chemistry 395:123-141, 2014.

6) Jianli Hu, Xiaobo Bai, Jonathan R. Bowen, Lee Dolat, Farida Korobova, Wenqian Yu, Peter W. Baas, Tatyana Svitkina, Gianluca Gallo, Elias T. Spiliotis. Septin-driven coordination of actin and microtubule remodeling regulates the collateral branching of axons. Current Biology 22: 1109-1115, 2012

Book Chapters:

1) Elias T. Spiliotis, Eva P. Karasmanis, Lee Dolat. Live-cell imaging of septins with the actin and microtubule cytoskeleton (2016). Methods in Cell Biology. In press.

2) Elias T. Spiliotis and Lee Dolat. Septins: cytoskeletal filaments with structural and regulatory functions (2016). Encyclopedia of Cell Biology. Elsevier. 2015.

Honors: Ruth L Kirschstein NRSA F31 pre-doctoral fellowship. NCI. NIH 2014 – 2016 President. Biology Graduate Student Association. 2014-2015 Teaching Assistant of the Year. University-wide nomination. 2011-2012