CINGULIN IS A COMPONENT OF THE APICAL MEMBRANE INITIATION SITE THAT INDUCES
EPITHELIAL CELL POLARIZATION AND LUMEN FORMATION
by
ANTHONY MANGAN
B.S., St. John Fisher College, 2009
M.S., Rochester Institute of Technology, 2012
A thesis submitted to the
faculty of the Graduate School of the
University of Colorado in partial fulfillment
of the requirements of the degree of
Doctor of Philosophy
Molecular Biology Program
2017
This thesis for the Doctor of Philosophy degree by
Anthony Mangan
Has been approved for the
Molecular Biology Program
by
James McManaman, Chair
Moshe Levi
Jeffrey Moore
Chad Pearson
Rui Zhao
Rytis Prekeris, Advisor
Date: 05-19-2017
ii Mangan, Anthony (Ph.D., Molecular Biology)
Cingulin is a Component of the Apical Membrane Initiation Site that Induces Epithelial Cell
Polarization and Lumen Formation
Thesis directed by Professor Rytis Prekeris
ABSTRACT
Epithelial cells are structurally and functionally polarized to transport specific molecules
while maintaining a trans-epithelial barrier. Additionally, epithelial cells coordinate their
polarization with neighboring cells to form an apical lumen, a key step in the establishment of
epithelial tissue architecture, and thereby function. Recent work from several laboratories
including ours identified Rab11 and its binding protein FIP5 as major components that regulate
apical endosome transport during apical lumen formation. I demonstrate that Rab11/FIP5-
containing endosomes (FIP5-endosomes) mediate the formation of apical lumen by targeted
delivery of apical lumen proteins. Using MDCK cells in a 3D tissue culture system, it was also
shown that targeting of FIP5-endosomes to the apical membrane initiation site (AMIS) is a key
step for the formation and expansion of an apical lumen in epithelial cysts. Furthermore,
midbody formation during telophase is the first symmetry-breaking event that determines the
site of lumen formation between two epithelial cells.
Despite recent advances in our understanding of the mechanisms mediating apical
lumen formation, many questions regarding the initiation of lumen formation remain
unanswered. Here, I also focus on the identification of the machinery that mediates AMIS
establishment and FIP5-endosome targeting during apical lumen formation. A new FIP5-
interacting protein, Cingulin, is identified. Cingulin is a tight junction protein that localizes to the
AMIS and is an ideal tether to mediate FIP5-endosome targeting during apical lumen formation.
The machinery mediating Cingulin recruitment to the midbody during late telophase was also
3 analyzed and it demonstrates that both microtubule binding and actin networks are required for
establishing the site of the apical lumen. I have also shown that Cingulin binds to microtubule C-
terminal tails (CTTs) and that this binding is likely regulated by glutamylation of the tubulin tails.
I completed immunoprecipitation, immunofluorescence, and proteomics analysis of
synchronized epithelial cells in telophase and identified components of the WAVE/SCAR
complex as putative regulators of Cingulin recruitment to the midbody. Since Rac1 is known to
activate the WAVE/SCAR complex, it was demonstrated that Rac1 is also present at the midbody
and that Rac1 activation is required for Cingulin recruitment to the midbody during apical lumen
formation. Additionally, the formation of actin flares at the midbody during late telophase were
observed and these actin flares may initiate cell polarization and apical lumen formation during
epithelial morphogenesis. This data supports a combinatorial role of microtubules and actin in
the coordination and regulation of the apical membrane initiation site and forming lumen.
The form and content of this abstract are approved. I recommend its publication.
Approved: Rytis Prekeris
4 TABLE OF CONTENTS
CHAPTER
I. POLARIZED PROTEIN TRANSPORT AND LUMEN FORMATION DURING EPITHELIAL
TISSUE MORPHOGENESIS
Abstract .……….….…………………………………………………………………………………… ...... 1
Introduction ………….………………………….…………………………………………………… ...... 1
Polarization of Individual Epithelial Cells …..……………………………………………………..2
Polarity complexes and actin cytoskeleton .……...…………………………………3
Polarized membrane traffic .…………………………...…………………………………..5
Apical Lumen Formation .……………………………………………………………………… ...... 8
In vitro models of de novo apical lumen formation ………………………………8
Mechanisms of de novo apical lumen formation during epithelial tissue
morphogenesis …………………………………………………………………………………..13
The mechanisms of lumen extension and coalescence ……………………….16
Summary and Future Objectives ..……………………………………………………………… ...... 17
II. SPATIOTEMPORAL DYNAMICS OF FIP5 AND CINGULIN DURING APICAL LUMEN
FORMATION
Abstract .……….….…………………………………………………………………………………… ...... 24
Introduction ………….………………………….…………………………………………………… ...... 25
3D Time-Lapse Method …………………………………………………………………………………..26
Materials …………………………………………………………………………… ...... 26
Detailed instructional method ……………………………….…………………………..27
Results …………………………………………………………………………………………………………….30
5 Midbody and central spindle microtubules mediate AMIS formation and
FIP5-endosome transport during telophase ……………………………………….30 Discussion
……………………………………………………………………………………………………….31
III. CINGULIN AND ACTIN MEDIATE MIDBODY-DEPENDENT APICAL LUMEN FORMATION
DURING POLARIZATION OF EPITHELIAL CELLS
Abstract .……….….…………………………………………………………………………………… ...... 38
Introduction ………….………………………….…………………………………………………… ...... 38
Results …………………………………………………………………………………………………………….40
Cingulin is a FIP5 binding protein concentrated at the AMIS ………… ...... 40
CGN is required for single apical lumen formation ……………………………..42
The WAVE/Scar complex is present at the AMIS …………………………………43
Active Rac1 is required for AMIS formation at the midbody ……………….44
CGN binding to tubulin mediates targeting to the midbody ……………….49
Discussion ……………………………………………………………………………………………………….52
Methods ………………………………………………………………………………………………………...55
Plasmids and antibodies ………………………………………………………………… .... 55
CGN knock-out by CRISPR/Cas9 in MDCK cells …………………………………...56
Immunoprecipitation and proteomic analysis …………………………………….56
Protein expression and purification ……………………………………………………57
Glutathione bead pull-down assays …………………………………………………...58
Cell culture …………………………………………………………………………………………59
Immunofluorescent, time-lapse, and FRET microscopy ………………………59
MDCK cell lines …………………………………………………………………………………..60
Rac1 inhibition studies ……………………………………………………………………….60
6 Yeast tubulin purification …………………………………………………………………..61
Microtubule binding assays ……………………………………………………………….61
Western blots ……………………………………………………………………………………62
IV. CONCLUSIONS AND FUTURE DIRECTIONS
Conclusions ………………………………………………………………………………………………………….92
Future Directions …………………………………………………………………………………………………94 References
………………………………………………………………………………………………………………………..97
vii LIST OF FIGURES
Figure
1.1 The proteins that establish and maintain epithelial cell polarity.
1.2 Cavitation (A) and Hallowing (B) models of apical lumen formation in vitro.
1.3 The role of the midbody during apical membrane initiation site (AMIS)
formation and apical endosome recruitment.
1.4 Model depicting apical lumen coalescence during the formation of the zebrafish
intestinal lumen.
2.1 Apical membrane initiation site (AMIS) formation around the midbody at late
telophase mediates FIP5-endosome targeting during lumen formation.
2.2 FIP5-endosomes deliver apical proteins along central spindle microtubules to
the midbody-associated AMIS.
2.3 Figure 2.3. Proposed models of FIP5-endosome trafficking and apical lumen
initiation.
3.1 Cingulin (CGN) is an AMIS-associated FIP5-binding protein.
3.2 CGN is a FIP5-interacting protein.
3.3 Cingulin is required for the formation of a single apical lumen.
3.4 CGN knock-down affects apical lumen formation.
3.5 CGN depletion affects actin cytoskeleton in MDCK cells.
3.6 Components of the WAVE/Scar complex and branched actin filaments are
present at the midbody during lumen formation.
3.7 Midbody and Cingulin associated actin flares.
3.8 Rac1 is activated at the AMIS.
8 3.9 Rac1 inhibition affects midbody-associated AMIS formation.
3.10 Rac1 is required for apical lumen formation.
3.11 Rac1 is required for the gp135 targeting during apical lumen formation.
3.12 Rac1 is required for the formation of a single apical lumen.
3.13 Electrostatic interactions mediate CGN binding to microtubule C-terminal tails
(CTTs).
3.14 CGN binds to tubulin C-terminal tail domains.
3.15 Generation of CGN knock-out in MDCK cells using CRISPR/Cas9.
3.16 Mutation of basic patch disrupts subcellular CGN targeting.
3.17 CGN interaction with midbody microtubules and Rac1-induced branched actin
cytoskeleton is required for AMIS formation and apical lumen initiation.
3.18 Uncropped scans of western blots.
9 CHAPTER I
POLARIZED PROTEIN TRANSPORT AND LUMEN FORMATION DURING EPITHELIAL
1 TISSUE MORPHOGENESIS
Abstract
One of the major challenges in biology is to explain how complex tissues and organs
arise from the collective action of individual polarized cells. The best-studied model of this
process is the cross talk between individual epithelial cells during their polarization to form
multi-cellular epithelial lumens during tissue morphogenesis. Multiple mechanisms of apical
lumen formation have been proposed. One of the most widely accepted is that new epithelial
lumens form from pre-existing polarized epithelial structures by wrapping epithelial sheets of
budding new epithelial branches. However, a de novo lumen formation mechanism has recently
emerged as an important driver of epithelial tissue morphogenesis, especially during formation
of small epithelial tubule networks. In this chapter I will discuss the latest findings regarding the
mechanisms and regulation of de novo lumen formation in vitro and in vivo.
Introduction
Epithelial tissues are composed of polarized cells, which function as selectively
permeable barriers. The plasma membrane of individual epithelial cells is divided into the apical
and basolateral domains, and specialized junctional complexes between adjacent cells, such as the tight junctions (TJs), maintain the separation of apical and basolateral plasma membrane
domains within an epithelial layer (Rodriguez-Boulan et al., 2004). Additionally, epithelial cells
coordinate their polarization with neighboring cells to form an apical lumen, a key step in the
1 Portions of this chapter are published with permission from our previously published article Blasky A, Mangan A, Prekeris R. Polarized Protein Transport and Lumen Formation During Epithelial Tissue Morphogenesis. Annual Review of Cell and Developmental Biology. 2015. 31:575-91.
1 establishment of renal and gut architecture, and thereby function (Bryant and Mostov, 2008).
Indeed, malfunctions in epithelial cell polarization and apical lumen formation are responsible
for a variety of renal and intestinal disorders, such as polycystic kidney disease, renal tubular
acidosis, microvilli inclusion disease (MVID) and diabetes insipidus. Additionally, tissue
morphogenesis requires coordinated action of groups of epithelial cells within three-
dimensional (3D) space to establish spatial order and long-range luminal organization. How the
formation of apical lumen is regulated and coordinated has been one of the major questions in
the field of epithelial cell biology and development.
Recent work has shown that apical lumen formation is a complex process that involves
dynamic restructuring of cytoskeleton and endocytic membrane transport, as well as formation of specialized polarity complexes. Despite these advances, many questions remain unanswered.
How are apical endosomes targeted and regulated during apical lumen formation? How do cells
establish a single apical lumen site? Do the mechanisms of lumen formation established in vitro
also apply to epithelial morphogenesis in vivo? Improvement in imaging and the establishment
of 3D tissue culture systems to study lumen formation are now permitting these questions to be
addressed in greater detail. Indeed, during the last few years, multiple new models explaining
the mechanisms of lumen formation have been proposed. Here I will discuss the latest findings
regarding the mechanisms and regulation of de novo lumen formation in vitro and in vivo.
Polarization of Individual Epithelial Cells
Epithelial cells are structurally and functionally polarized to directionally transport
specific molecules while maintaining a transepithelial barrier. This selective transport is achieved by partitioning of the plasma membrane into distinct apical and basolateral domains, which
have distinct lipid compositions and contain specific sets of proteins. The apical plasma
membrane is enriched in cholesterol, sphingolipids, glycolipids, and contains various specific
2 highly glycosylated proteins. In contrast, the basolateral membrane is enriched in E-cadherin,
integrin molecules as well as growth factor receptors (Mostov et al., 2003; Nelson, 2003;
Rodriguez-Boulan et al., 2004). Tight junctions separate the apical and basolateral domains and
provide a diffusion barrier that prevents mixing of domain specific membrane components
(Figure 1.1). Since the fidelity of apical versus basolateral protein and lipid distribution is crucial
to a variety of epithelial functions, cells have developed mechanisms to selectively retain or
deliver proteins to basolateral and apical regions of the plasma membrane (Mostov et al., 2003;
Nelson, 2003; Rodriguez-Boulan et al., 2004).
Polarity complexes and actin cytoskeleton
Polarity complexes are the key protein complexes that regulate the polarization of
epithelial cells. Three major polarity complexes have been identified, each of them localizing to
different plasma membrane (PM) domains where they mediate the establishment and identify
boundaries of apical and basolateral domains (Bryant and Mostov, 2008; Nelson, 2003). SCRIB
and Crumbs complexes localize to and regulate the formation of lateral and apical PM domains,
respectively (Figure 1.1) (Nelson, 2003; Schluter and Margolis, 2012). In contrast, the Par3/6
complex localizes predominately to tight junctions and appears to play a key role in defining
apical and lateral PM boundaries (Schluter and Margolis, 2012). The key to Par3/6 complex
function is the recruitment and activation of atypical PKC (aPKC) to tight junctions.
Phosphorylation of Crumbs by aPKC promotes apical PM formation and expansion (Schluter and
Margolis, 2012), while phosphorylation of Lgl (a component of SCRIB complex) leads to its
degradation and suppression of lateral PM expansion (Bryant and Mostov, 2008; Schluter and
Margolis, 2012). The Par3/6 complex also interacts with Cdc42, a small monomeric GTPase that
plays multiple roles during polarization. One of the main functions of Cdc42 is to regulate the
recruitment of Par3/6 to tight junctions, thus promoting apical PM formation (Bryant and
3 Mostov, 2008; Schluter and Margolis, 2012). Additionally, Cdc42 regulates apical actin
polymerization, mitotic spindle orientation and exocytosis/endocytosis of apical endosomes, the
processes that are required for epithelial cell polarization. A detailed description of polarity
complexes during epithelial cell polarization is outside the scope of this review, and several good
reviews on this topic are available (Bryant and Mostov, 2008; Nelson, 2003; Schluter and
Margolis, 2012).
The actin cytoskeleton also plays a major role in initiation and maintenance of epithelial
cell polarity. It is well established that the actin cytoskeleton accumulates just beneath the
apical plasma membrane (Figure 1.1) and that actin polymers form a dense network that also
includes tropomyosin and cytokeratin-based intermediate filaments (Apodaca et al., 2012). This
network, referred to as the terminal web, was shown to play a major role in mediating apical
localization of membrane proteins by anchoring them to the actin cytoskeleton via various actin
binding proteins, such as EBP-50 (Bretscher et al., 2000). In addition, actin also forms a
junctional belt that is required for the formation and stability of tight junctions. At least to some
extent, formation of the junctional belt is mediated by Cdc42 via recruitment of WASP and
activation of Arp2/3 (Mellman and Nelson, 2008).
In some epithelial cells, the establishment of the apical terminal web is followed by the
formation of microvilli, the apical structure that are specialized for efficient nutrient uptake.
Disruption of microvilli formation leads to microvilli inclusion disease (MVID) and microvilli
atrophy (Ameen and Salas, 2000; Erickson et al., 2008). The mechanisms mediating microvilli
formation are only beginning to emerge, but recent work demonstrates that Rap2a delivery to
the apical PM initiates a signaling cascade that leads to the apical recruitment (from the trans-
Golgi Network) of MST4 kinase and phosphorylation of Ezrin, resulting in the formation of the
actin-rich microvilli (Figure 1.1) (Gloerich et al., 2012). The terminal web is also required for
4 microvilli stability, since microvilli terminate as actin filament “rootlets”, that anchor into actin
substructure of the terminal web by binding to Myo2 and fodrin (Bretscher et al., 2002). What
remains to be understood is how Rap2a is targeted to the apical PM. Recent work has shown
that Rap2a localizes to Rab11-endosomes (Bruurs and Bos, 2014). This raises the intriguing
possibility that Rab11-endosomes deliver Rap2a to the apical plasma membrane to initiate
microvilli formation. Consistently, it was shown that Rab11-binding protein myosin-Vb is required for microvilli formation and is mutated in MVID patients (Muller et al., 2008; Szperl et
al., 2011). Collectively, these new data uncover cross talk between polarity complexes,
endocytic transport and actin cytoskeleton that underlie establishment of epithelial cell polarity.
Polarized membrane traffic
Epithelial polarity is tightly regulated by polarized membrane transport. Polarized
epithelial cells have domain specific early endosomes: apical and basolateral endosomes, as well
as, a common recycling endosomes (Figure 1.1). In addition, most polarized epithelial cells have
specialized compartment of apical recycling endosomes (ARE) that are involved in regulated
recycling of specialized apical proteins. For example, regulated insertion of H+/K+-ATPase into
the apical plasma membrane of gastric parietal cells is responsible for selective secretion of
hydrochloric acid into the stomach lumen and is mediated by apical recycling endosomes
(Calhoun et al., 1998; Duman et al., 1999). A majority of apically or basolaterally internalized
proteins eventually are transferred to common recycling endosomes, where they are selectively
segregated and sorted for transport to their final destinations (Figure 1.1). For instance,
basolateral proteins such as E-cadherin, transferrin receptor or LDL receptor are targeted from
common recycling endosomes to basolateral plasma membrane via a pathway that involves
AP1B coat protein, as well as the Exocyst protein complex (Lock et al., 2005; Lock and Stow,
2005). In addition, newly synthesized proteins exiting the trans-Golgi Network (TGN) are also
5 sorted and delivered to either apical or basolateral plasma membrane. Interestingly, some data suggest that from TGN many proteins are directed to recycling endosomes before they are sorted to the apical or basolateral surfaces.
It is now well established that polarized epithelial rely on domain-specific endosomes to act as intermediaries for polarized transport of endocytic cargo to their appropriate PM domains
(Figure 1.1) (Apodaca et al., 2012). The traffic through these domain-specific endosomes is mediated by Rabs, small monomeric GTPases that act as master regulators of membrane transport (Figure 1.1). Rabs function by recruiting and activating various membrane transport regulators, known as effectors, to their respective membranes. These Rab/effector complexes are responsible for very specific and selective regulation of distinct membrane transport steps.
To date several Rabs have been implicated in regulating epithelial transport, namely Rab8,
Rab10, Rab11, Rab14, Rab17 and Rab25 (Figure 1.1) (Apodaca et al., 2012).
Rab11-family GTPases (Rab11a, Rab11b and Rab25 isoforms) have emerged as key regulators of polarized transport in epithelial cells. Interestingly, while Rab11a and Rab11b are ubiquitously expressed, Rab25 is present only in epithelial cells (Goldenring et al., 2001;
Goldenring et al., 1993). Multiple studies have implicated Rab11-family proteins in regulating apical plasma membrane trafficking via ARE (Casanova et al., 1999). For example, expression of
Rab11a/b dominant-negative mutants inhibits apical recycling and transcytosis of IgA (Casanova et al., 1999; Wang et al., 2000). Rab11 was also implicated in regulating H+/K+-ATPase dependent secretion of hydrochloric acid into the stomach lumen. In resting parietal cells H+/K+-
ATPase is sequestered within subapical tubulo-vesicles. Upon stimulation, the H+/K+-ATPase containing tubulo-vesicles (probably equivalent of ARE) fuse with the apical plasma membrane, which results in delivery of H+/K+-ATPase and subsequent secretion of hydrochloric acid. This stimulated delivery of H+/K+-ATPase was shown to depend on the Rab11a/b GTPase, as well as,
6 its binding proteins (Calhoun et al., 1998). In addition to regulating apical recycling, Rab11 also
has been suggested to regulate transcytosis of proteins endocytosed at the basolateral for
secretion at apical plasma membrane domains (Apodaca et al., 1994). Finally, recent studies
have implicated Rab11 in regulating biosynthetic delivery of E-cadherin from TGN to basolateral
plasma membrane (Lock et al., 2005), as well as endolyn delivery from TGN to apical plasma
membrane (Potter et al., 2006).
Since Rab GTPases often bind to multiple effector proteins, much effort has been
dedicated to the identification of Rab11a/b effector proteins. It was shown that Rab11a/b binds to myosin-Vb and Sec15 (subunit of the Exocyst complex) (Hales et al., 2002; Zhang et al., 2004).
This presumably allows specific targeting of Rab11-endosomes to the apical PM since the
Exocyst complex functions as a plasma membrane tether, and myosin-Vb likely mediates the
binding of Rab11-vesicles to actin-rich terminal web. Importantly, Rab11a also binds to and
recruits Rabin8. Rabin8 is GTP exchange factor (GEF) that functions by activating Rab8, a GTPase
that was also implicated in polarized endocytic transport (Figure 1.1) (Bryant et al., 2010).
Consequently, the Rab11-Rabin8-Rab8 cascade was shown to be involved in regulating several
polarized transport steps, including membrane transport to forming primary cilia (Westlake et
al., 2011). Rab11a/b GTPases also bind to Rab11 family-interacting proteins (known as FIPs)
(Hales et al., 2001; Prekeris et al., 2001; Prekeris et al., 2000). FIPs act as scaffolds for the
recruitment of additional endocytic transport factors (Prekeris, 2003), with FIP2 and FIP5
involved in apically-directed endosomal transport (Prekeris et al., 2000; Willenborg et al., 2011).
Significantly, FIP2 was shown to bind to myosin-Vb, thus mediating Rab11/FIP2/myosin-Vb
complex formation (Hales et al., 2002; Lapierre et al., 2001). Similarly, FIP5 was shown to bind to
Kinesin-2 and sorting nexin 18, both known regulators of apical protein transport (Li et al.,
2014a; Willenborg et al., 2011). Thus, it is now clear that a complex network of endosomal
7 transport pathways, as well as different Rab/effector complexes, are required for epithelial cell
polarization.
Apical Lumen Formation
One of the major challenges in biology is to explain how the interactions of individual
polarized cells result in the formation of complex tissues. While the mechanisms that govern
individual epithelial cell polarization have been well studied, little is known about how these
cells communicate to organize and form polarized structures, such as apical lumens within
epithelial ducts and terminal end buds. There are several ways to form an apical lumen. One of
them is to form new lumens from pre-existing polarized epithelia by either folding epithelial
sheets or budding/sprouting from epithelial tubes. This type of lumen formation has been
actively studied in the last couple decades and is well-described in recent reviews (Andrew and
Ewald, 2010). Much more enigmatic is the de novo formation of apical lumens from non-
polarized cells. One of the key features of de novo lumen formation is simultaneous polarization
of epithelial cells and establishment of new lumen. The molecular machinery mediating the
coordination between polarization of individual epithelial cells and apical lumen formation in
vitro and in vivo is only beginning to be defined and will be the focus of the rest of this chapter.
In vitro models of de novo apical lumen formation
The key to the advances in our understanding of lumenogenesis was the development
of 3D tissue culture systems. These 3D assays relay on the suspension and growth of individual
epithelial cells in extracellular matrix (ECM). Typically, Matrigel or collagen matrices are used to mimic the in vivo extracellular environment. These matrices provide specific structural and/or molecular signaling interactions required for epithelial cells to form spherical cysts with a single
central hollow luminal. These cysts, or acini, are excellent model system for studying the
mechanisms of apical lumen formation during tissue morphogenesis. As the result of these
8 studies two different, but not mutually-exclusive, models of lumen formation have been
proposed: cavitation and hollowing.
Cavitation model: The first model describing the mechanisms of de novo lumen formation was
proposed by Joan Brugge and colleagues based on their work using the breast epithelial cell line
MCF10A and is currently known as the cavitation model (Debnath and Brugge, 2005). Cavitation
(also called canalization, lumenization) is the creation of a luminal space within a pre-existing
mass of cells that involves apoptosis of cells within the developing lumen. In the in vitro 3D
model system, cells that are in contact with the ECM can differentiate and establish apical/basal
polarity (Figure 1.2A) (Debnath and Brugge, 2005). In contrast, the cells that are located inside
the spheroid (sometimes referred to as the mamosphere) are not able to contact with ECM and
consequently do not receive the necessary survival signals and therefore triggering apoptosis
(Debnath and Brugge, 2005). The apoptotic cells are then cleared, resulting in the formation of
hollow acini with a single apical lumen (Figure 1.2A).
Tissues that undergo cavitation include mammary and salivary glands, which rely on
highly proliferative states to produce an interconnecting network of ducts with terminal
endbuds (Coucouvanis and Martin, 1995; Jaskoll and Melnick, 1999; Mailleux et al., 2008).
Consistently, in 3D cultures of the non-tumorogenic human mammary cell line MCF10A,
targeted knockdown of the pro-apoptotic BCL-2 family proteins Bim and Bmf cause a decrease
in apoptosis and impair luminal formation (Mailleux et al., 2007; Reginato et al., 2005; Schmelzle
et al., 2007). However, impaired luminal formation is only temporary and the acini eventually
do form hollow apical lumens after a few more days in culture (Mailleux et al., 2007).
Interestingly, activation ErbB2 (EGF receptor/Her2), the receptor known to be over-expressed in some breast cancers, causes an increase in proliferation and filling of the apical lumen
(Muthuswamy et al., 2001). Thus, it is clear that multiple, partially redundant molecular
9 mechanisms are involved in cavitation and further studies will be needed to identify all the
components of this lumen formation pathway.
Hollowing model: While the cavitation model explains lumen formation in some tissues, it is also
clear that in many cases apoptosis is not required to form an apical lumen. It was shown that
apical lumen can form de novo between two cells as the result of coordinated of cell division
and polarization (Figure 1.2B) (Bryant et al., 2010; Li et al., 2014b). This mechanism of lumen
formation model, known as hollowing, was originally proposed by Keith Mostov and colleagues
based on their work using Madin-Darby canine kidney (MDCK) cells (Bryant et al., 2010; Datta et
al., 2011). In this model the apical lumen forms between as few as two cells and is dependent on
targeted trafficking of apical cargo to a specific location known as the apical membrane initiation site (AMIS) (Figure 1.2B). AMIS formation appears to be the key step in establishing the
location of nascent apical lumen and provides the targeting site for apical endosomes (Bryant et
al., 2010; Li et al., 2014b; Willenborg et al., 2011). The AMIS is a transient structure that contains the Par3/Par6 polarity complex, the Exocyst targeting complex and the canonical tight junction
proteins Cingulin and ZO1 (Bryant et al., 2010; Li et al., 2014b). As an apical lumen forms, AMIS
matures into the tight junctions that define the boundary between apical and basolateral PM
(Bryant et al., 2010; Ferrari et al., 2008; Li et al., 2014b) (Figure 1.2B).
Targeted transport and fusion of apical endosomes to the AMIS underlies the formation
of nascent apical lumens (Figure 1.2B). When embedded into the ECM, each cyst begins as a non-polarized single cell. In these cells, apical cargo such as Crumbs3a and glycoprotein-135 (gp-
135, also called podocalyxin) are either dispersed within the plasma membrane or located
within the sub-population of recycling endosomes (Bryant et al., 2010; Ferrari et al., 2008).
During apical lumen initiation, gp135 and Crumbs3a are concentrated into Rab11a containing
transport vesicles. Formation and targeting of these vesicles is dependent on the interactions
10 between Rab11 and its effector protein FIP5 (Figure 1.3). Consistently, FIP5 depletion in MDCK
cells induces the formation of multi-luminal cysts (Willenborg et al., 2011). Furthermore, FIP5 directly interacts with a sorting nexin 18 (SNX18), the protein that was shown to increase apical
transport vesicle formation (Figure 1.3) (Willenborg et al., 2011).
FIP5 has also been shown to directly interact with the motor protein Kinesin-2 (Li et al.,
2014a). This interaction allows Rab11/FIP5-containing vesicles to be transported along
microtubules toward the forming apical domain (Figure 1.3) (Li et al., 2014a). Interestingly,
Kinesin-2 competes with SNX18 for FIP5 binding, further supporting a step-wise flow from
vesicle budding to vesicle transport (Li et al., 2014a). Although these vesicles are known to fuse
at the AMIS, a specific targeting or tethering mechanism has yet to be determined, and it is
likely the result of a combination of factors. The Exocyst complex is involved in vesicle fusion
and could serve as a tether for Rab11/FIP5 vesicles at the AMIS (Datta et al., 2011). The
Rab11/FIP5 apical vesicles also contain Rab8 and Cdc42, which could regulate targeting to
aPKC/Par3 complex located at the AMIS (Figure 1.3) (Bryant et al., 2010). Importantly, apical
endosome-associated Rab8 is activated by the Rabin8, a GEF for Rab8, which is recruited to the
apical vesicles through direct binding to Rab11a (Bryant et al., 2010). While it is clear that Cdc42 and Rab8 are important for apical lumen formation, the molecular basis for their role during
lumenogenesis remains to be fully defined.
Recently, several other molecules have been implicated in the regulation of vesicular
transport and targeting to the AMIS. It was shown that synaptotagmin-like proteins Slp2a and
Slp4a are required for single lumen formation (Figure 1.3) (Galvez-Santisteban et al., 2012). It
was proposed that Slp2a is concentrated at the AMIS through its interactions with PIP2, thus
mediating the apical targeting of Rab27-containing transport vesicles. Similarly, Slp4a targets
Rab3 to the same Rab27-containing vesicles and may participate in targeting these vesicles to
11 AMIS (Galvez-Santisteban et al., 2012). What remains unclear is whether Rab11 and FIP5 are
also present in Rab27-containing organelles. One possibility is that Rab27/Rab3 and Rab11
dependent targeting pathways may function as a coincidence detection system ensuring the
fidelity of apical protein targeting to AMIS. Alternatively, Rab27/Rab3 and Rab11 vesicles may
be transporting different apical cargo to the AMIS during lumen formation.
One question that still remains is how cells create a single AMIS. Recent work has shown
that formation of the midbody during the first cell division is the first “symmetry-breaking”
event that initiates the polarization of the two daughter cells and induces lumen formation
(Figure 1.3) (Li et al., 2014b; Schluter et al., 2009). It was also shown that the AMIS forms around
the midbody during late telophase, thus marking the apical domain that is retained and
expanded with subsequent cell divisions (Li et al., 2014b). Importantly, during metaphase and
anaphase of the first cell division Rab11/FIP5-endosomes are localized at the centrosomes and
are not transported to plasma membrane until after midbody formation in late telophase
(Figure 1.3) (Li et al., 2014a; Li et al., 2014b). The machinery that mediates the timing of
Rab11/FIP5-endosome transport remains to be fully defined. However, it was shown that FIP5-
T276 phosphorylation by GSK-3 during metaphase and anaphase inhibits FIP5 interaction with
SNX18, thus preventing apical vesicle budding until late telophase (Li et al., 2014b).
Both the cavitation and hollowing methods provide viable mechanisms of de novo
lumen formation in vitro. Why two different means of lumen formation are needed in vivo
remains to be understood. It is possible that distinct lumenogenesis mechanisms function in
different tissues to accommodate the tissue-specific dynamics and regulation of morphogenesis.
Consistently, it was proposed that cavitation is a predominant mechanism of lumen formation
during mammary duct morphogenesis and remodeling. It is also possible that, at least in some
tissues, the formation and maintenance of a single lumen is a result of both lumenogenesis
12 methods. For example, apical lumen may be initially formed by hollowing, and apoptosis may
only be used during lumen expansion and maintenance to remove any cells that begin to invade
the luminal space.
Mechanisms of de novo apical lumen formation during epithelial tissue morphogenesis
Our understanding of apical lumen formation has been derived primarily through in
vitro 3D lumen formation assays (as described above). Although in vitro cell culture is a powerful
experimental approach for identifying candidate molecular mechanisms of lumen formation, it
lacks the inherent complexity of in vivo morphogenesis and organogenesis, which draw on
dynamic polarity cues and positional information. Despite the wealth of information on lumen
formation derived from in vitro studies, our understanding of how lumens form in vivo remains
limited. Consequently, experimentally manipulatable animal models are needed to understand
how lumen forms in the context of in vivo development. The next section highlights recent
advancements in understanding epithelial de novo lumen formation in vivo using fly, worm and
fish models.
Drosophila melanogaster tracheal cells: The Drosophila tracheal system is a powerful model
system for identifying and investigating molecular mechanisms of lumen formation in vivo. The
fly tracheal system is comprised of a network of epithelial tubes that transport oxygen to
tissues. During embryonic development the tracheal system forms by invagination of epidermal
placodes. Cells migrate from sites of placode invagination to form primary branches. These
primary branches connect with cognate branches from adjacent primordia, building an interconnected network with a continuous lumen (Samakovlis et al., 1996).
De novo lumen formation occurs throughout the Drosophila developing tracheal system.
Specialized fusion cells mediate lumen formation and elongation within primary branches. The
site at which fusion cells contact each other acquires apical characteristics that depend on
13 localized increase in nucleation of actin and microtubule cytoskeleton. Actin and microtubules
aid in targeted transport of apical cargo and establishment of cell structure (Lee et al., 2003; Lee
and Kolodziej, 2002). Vesicles and apical proteins, including polarity proteins aPkc, Bazooka, and
Crumbs are then targeted to the contact region to aid in lumen formation (Gervais et al., 2012).
The small GTPase Arf-like 3 (Arl3) functions in the exocytic transport of cargo to the fusion site
(Kakihara et al., 2008).
The fly tracheal system also contains terminal cells that connect to the tubular network
via an invagination around a circular adherens junction. Previously, the terminal cell lumen was
thought to form by coalescence of intracellular vesicles, however recent data suggests that the
lumen is formed by addition of apical membrane at the trunk cell junction site (Gervais and
Casanova, 2010). The initial site of lumen growth into terminal cells is defined by the
accumulation of microtubules (Gervais and Casanova, 2010). Microtubules extend from the
intercellular junction to the cell boundary before the terminal cell elongates and any subcellular
lumen is formed. Tyrosinated tubulin is specifically enriched at the front of the growing lumen,
and may act as a guide for lumenogenesis (Gervais and Casanova, 2010), reminiscent of vesicle
delivery to forming lumen along central spindle microtubules during hollowing in vitro (see
Figure 1.3).
Vesicle transport is also a key step during the formation of a lumen in terminal cells.
Mutations in NSF2, the protein that is required for SNARE complex disassembly, disrupt apical
membrane expansion (Song et al., 2013). Further, Germinal center kinase III is required for
regulating the traffic of material to the apical domain (Song et al., 2013). The Exocyst complex, a
known component of AMIS, is also required for plasma membrane morphogenesis in terminal
cells, presumably by mediating the targeting and tethering of apical transport vesicles.
Importantly, another AMIS component the Par3/6 polarity complex provides membrane
14 localization cues for the Exocyst (Jones and Metzstein, 2011). Rab35 has also been implicated in
lumen formation in vivo (Schottenfeld-Roames and Ghabrial, 2012), although it’s role in
lumenogenesis remains to be defined.
Caenorhabditis elegans excretory cells: Studies utilizing the C. elegans excretory system also
provide significant insights into lumen formation in vivo. The C. elegans excretory system
consists of five epithelial cells that form fluid filled tubules. The excretory cell is polarized with an apical plasma membrane along the luminal surface and contributes to most of the luminal
structure of the system. During development the excretory cell grows in an H shape, with four
processes extending anteriorly and posteriorly along the body of the animal and these processes
continue to grow throughout development. Similar to MDCK cells in 3D tissue culture and fly
terminal cells, the worm apical membrane grows distally from the cell body through the
targeting and fusion of intracellular vesicles (Khan et al., 2013; Kolotuev et al., 2013). The
cytoplasm surrounding the tube contains cyst-like membrane structures called canaliculi. In
response to osmotic stress canaliculi fuse to the luminal membrane to rapidly increase the size
of the apical membrane (Khan et al., 2013; Kolotuev et al., 2013). The small GTPase RAL-1 and the polarity protein Par3 are both necessary for delivery of canaliculi to the lumen and excretory
lumen outgrowth (Armenti et al., 2014).
Danio rerio intestinal cells: Zebrafish have increasingly been used as a vertebrate model to
understand lumen formation, due in part to the ease of in vivo imaging and their amenability to
genetic manipulation. Recent work on de novo lumen formation during zebrafish vasculature
system development has provided valuable insights into the plasticity of lumen formation in vivo
(Blum et al., 2008; Lenard et al., 2013).
The zebrafish intestine has emerged as an equally useful model for investigating
epithelial de novo lumen formation. The zebrafish intestine originates as a solid rod of
15 endodermal cells that differentiate into polarized epithelial cells prior to lumen formation.
Interestingly, the gut lumen does not form through cavitation as the lumen forms without
apoptosis (Ng et al., 2005). Initially, actin rich foci localize to the site of nascent lumens (Horne-
Badovinac et al., 2001). Formation of these actin foci resembles AMIS formation in vitro, since
they contain many of the same proteins, including tight junction components and the Par3/6
complex. Furthermore, the Par3/6 complex component aPKC is required to establish epithelial
cell polarity and defects in aPKC function result in gut with multiple lumens (Horne-Badovinac et
al., 2001), resembling the phenotypes observed in 3D tissue culture systems. After establishing
the site of lumenogenesis the subsequent lumen expansion is driven by fluid accumulation
generated by paracellular and transcellular ion transport. Ion transport in the zebrafish gut
epithelium is regulated, at least in part, by the junctional protein Claudin 15 and Na+/K--ATPase.
Consequently, disruption of their function also results in a multiple lumen defect (Bagnat et al.,
2007).
The mechanisms of lumen extension and coalescence
While in vitro models allow for the study of mechanisms governing lumen initiation,
they are mostly derived from studies of cysts forming from a single cell. In contrast, in animal
systems epithelial lumens often form from simultaneous polarization of multiple cells. Thus, the
coalescence of multiple mini-lumens emerged as a key step during formation of apical tubules
and networks. Lumen expansion and coalescence can be mediated through fluid accumulation and the generation of a turgor force. Turgor is generated by hydrostatic pressure, which builds
through the accumulation of ion channels and pumps at the apical plasma membrane. For example, MDCK cysts use the cystic fibrosis transmembrane conductance regulator for lumen
expansion (Bagnat et al., 2007). In the C. elegans excretory system, the water channel AQP8
16 mediates fluid accumulation and regulates the tube size (Khan et al., 2013; Kolotuev et al.,
2013).
Most of the newest insights into mini-lumen coalescence came from work on lumen formation in the zebrafish gut. Lumenogenesis in the developing zebrafish gut does not require apoptosis (Ng et al., 2005), suggesting that hollowing-like processes drive the formation and remodeling of intestinal epithelia. Consistently, it has been shown that zebrafish gut lumenogenesis is initiated by de novo formation of multiple mini-lumens. Upon maturation, these mini-lumens fuse providing a perfect system to study the machinery mediating lumen coalescence (Ng et al., 2005). Paracellular and transcellular ion transport, regulated by
Claudin15 and the Na+/K+-ATPase, drives fluid accumulation and has been identified as a key determinants of gut lumen expansion (Bagnat et al., 2007). However, lumen expansion resulting from fluid accumulation is not the sole mechanism driving lumen coalescence. Prior to lumen coalescence, enlarged lumens are present along the length of the zebrafish gut, separated by basolateral cell junctions, which lack apical and tight junction markers podocalyxin and ZO1
(Figure 1.4) (Alvers et al., 2014). The expansion of the zebrafish gut lumen occurs through both, breaking of basolateral cell contacts and the expansion of the apical membrane (Figure 1.4). This process presumably required the targeted delivery and fusion of new apical endosomes, while simultaneously removing basolateral plasma membrane proteins by endocytosis (Figure 1.4), although the machinery mediating and regulating this process remains unclear.
Summary and Future Objectives
Polarization of epithelial cells is a seminal step in epithelial tissue morphogenesis during development and epithelial tissue reorganization in adults. The emergence of in vitro 3D tissue culture models and novel imaging techniques led to significant progress in identifying the factors mediating the establishment of epithelial polarity at a single cell level, as well as during
17 formation of multi-cellular epithelial tissues. It is now known that the targeted transport and
fusion of apical endosomes plays pivotal roles in delivering the essential polarity factors and
formation of apical lumen (Bryant and Mostov, 2008). The roles of actin and microtubule
cytoskeleton during lumenogenesis are also well-established. However, many questions remain
unanswered. How are apical endosomes targeted during apical lumen formation? How do cells establish the site of single apical lumen? How are multiple mini-lumens resolve to form intricate
networks of tubes and terminal end buds? Further studies will be required to answer these
pivotal questions.
Recently, the midbody was identified as a unique cellular structure intimately involved
in regulating cell fate and polarization (Li et al., 2014b; Schluter et al., 2009; Wang et al., 2014).
In vitro studies have shown that midbody formation is the first “symmetry-breaking” event that
initiates epithelia polarization and determines the location of nascent apical lumen. Consistent
with these concepts, dramatic increase in cell division were found to precede the initiation of de
novo lumen formation in vivo. Midbody effects are not limited to epithelial cells as midbodies
were shown to regulate mitotic spindle positioning during early C. elegans development (Singh
and Pohl, 2014). Similarly, midbodies were shown to mark the location of axon outgrowth in fly
neurons (Pollarolo et al., 2011). Based on these and similar findings, midbodies are emerging as
important regulators of epithelial polarization, yet the machinery mediating these midbody
functions remains unknown. Further research will be needed to define the midbody function
during tissue morphogenesis.
Most of the work on apical lumen formation has been done using 3D tissue culture
based models. While these are very powerful experimental approaches, morphogenesis and
organogenesis in vivo draws on complex polarity/positional information cues that cannot be
fully replicated in cell culture. Thus, the extent to which proteins that regulate single lumen
18 formation in 3D tissue culture systems contribute to lumen formation in vivo remains to be determined. Furthermore, while in vitro epithelial cysts start as single cell embedded in extracellular matrix, in vivo most epithelial tissues form from hundreds of non-polarized cells that simultaneously undergo polarization to generate apical lumens. How the formation and coalescence of these mini-lumens is organized and regulated in vivo remains essentially unknown. The emergence of novel genomic editing approaches as well as new animal models, such as zebrafish, will allow for the analysis of the machinery of epithelial morphogenesis in complex vertebrate organ systems.
19
Figure 1.1. The proteins that establish and maintain epithelial cell polarity.
20
Figure 1.2. Cavitation (A) and hollowing (B) models of apical lumen formation in vitro. Red circles represent apical endosomes. Abbreviations: AMIS, apical membrane initiation site; TJ, tight junction.
21
Figure 1.3. The role of the midbody during apical membrane initiation site (AMIS) formation and apical endosome recruitment. Red indicates apical endosomes and all proteins known to be associated with them. Blue indicates AMIS-associated proteins. Proteins listed in black are known to be required for lumen formation but have unclear subcellular localization.
22
Figure 1.4. Model depicting apical lumen coalescence during the formation of the zebrafish intestinal lumen. (A) Definition of the apical lumen initiation site and initial formation of the zebrafish gut lumen require localization of apical and tight junction proteins. Both Claudin 15 and Na+/K+ -ATPase are required for the paracellular ion transport necessary for lumen fluid accumulation and initial gut lumen expansion. (B) Gut lumen expansion occurs along adjacent basolateral membranes (red line) through Rab11-mediated apical trafficking (blue) and concurrent with basolateral membrane recycling (red). (C) The mechanisms regulating the coalescence of multiple expanding apical lumens into a single continuous lumen remain unresolved. The Hedgehog pathway protein Smoothened is required for lumen resolution, but not lumen expansion, suggesting lumen expansion and resolution are two distinct processes.
23 CHAPTER II
2 SPATIOTEMPORAL DYNAMICS OF FIP5 AND CINGULIN DURING APICAL LUMEN FORMATION
Abstract
Apical lumen formation is a key step during epithelial morphogenesis. The
establishment of the apical lumen is a complex process that involves coordinated changes in
plasma membrane composition, endocytic transport and cytoskeleton. These changes are
brought about, at least in part, by the targeting and fusion of Rab11/FIP5-containing apical
endosomes with the apical membrane initiation site (AMIS). While the formation of the AMIS
and polarized transport of Rab11/FIP5-containing endosomes is crucial for the formation of a
single apical lumen, the spatiotemporal regulation of this process remains poorly understood.
Here it is demonstrated that midbody formation during cytokinesis is a symmetry-breaking
event that establishes the location of the AMIS.
Additionally, Fluorescent imaging of fixed cells grown in two-dimensional (2D) cultures is
one of the most widely used techniques for observing protein localization and distribution
within cells. Although this technique can also be applied to polarized epithelial cells that form
three-dimensional (3D) cysts when grown in a Matrigel matrix suspension, there are still
significant limitations in imaging cells fixed at a particular point in time. Here, the use of 3D-
time-lapse imaging of live cells to observe the dynamics of AMIS formation and lumen expansion
in polarized epithelial cells is described.
2 Portions of this chapter are published with permission from our previously published articles Li D, Mangan A, Cicchini L, Margolis B, and Prekeris R. FIP5 Phosphorylation During Mitosis Regulates Apical Trafficking and Lumenogenesis. EMBO Reports. 2014: 15(4):428-437, and Mangan A, Prekeris R. 3D-Time-Lapse Analysis of Rab11/FIP5 Complex: Spatiotemporal Dynamics During Apical Lumen Formation. Methods in Molecular Biology. 2015: 1298:181-6.
24 Introduction
The establishment of a solitary apical lumen is a crucial step during epithelial
morphogenesis of hollow organs. A recently proposed model of apical lumen formation (Bryant
et al., 2010; Bryant et al., 2008; Martin-Belmonte et al., 2008) demonstrates that apical proteins,
such as glycoprotein135 (gp135) and Crumbs3a (Crb3a), are initially localized at the outer
surface of forming cysts of MDCK cells in 3D cultures. These apical proteins are then selectively
endocytosed and transported to the apical membrane initiation site (AMIS), which contains several TJ proteins, such as ZO-1 and Cingulin (Bryant et al., 2010; Willenborg et al., 2011).
Cumulative fusion of apical endocytic carriers to the AMIS gives rise to a nascent lumen (Gin et
al., 2010).
It is now well-established that endocytic trafficking of apical components to the AMIS
plays a key role in apical lumen formation. Much work is dedicated to elucidating the molecular
mechanism of this transport process. Recent work has shown that FIP5, a Rab11-interacting
protein, mediates apical lumen formation by interacting with Sorting nexin 18 (SNX18) for the
formation of apical endocytic carriers from apical recycling endosomes (AREs) (Willenborg et al.,
2011). However, what remains largely unknown is the spatiotemporal regulation of AMIS
formation and FIP5-endosome targeting during early stages of lumenogenesis. Here, I describe
studies finding that FIP5-endosomes are transported along central spindle microtubules to the
midbody where the AMIS, as marked by Cingulin, forms during cytokinesis.
In addition, fluorescent microscopy has proven to be a useful tool in studying the
mechanisms and dynamics of lumen formation and many other pathways by allowing
visualization of co-localization and distribution of proteins in cells. While imaging of polarized
epithelial cells grown on two-dimensional (2D) filters is most widely used, it is not as sufficient
for looking at the development of three-dimensional (3D) multicellular structures such as the
25 apical lumen. Instead, epithelial cells can be grown in 3D culture by suspending in Matrigel matrix, which mimics the extracellular matrix and allows formation of 3D cysts with internal apical environments (Bryant et al., 2010; Willenborg et al., 2011). Although the 3D cysts can be fixed and immuno-labeled, this method limits the viewing of cells at a single time point and thus is not adequate for determining the timing and dynamics of apical lumen formation. These limitations can be overcome with the use of live imaging of tagged-proteins in live cells, which enables the following of a single cell though all of the stages of division as well as lumen formation and expansion.
I created a protocol for 3D-time-lapse analysis of apical lumen formation using Madin-
Darby canine kidney (MDCK) cells (Li et al., 2014a, b). Using this technique, I was able to determine that Cingulin is one of the first proteins recruited to the AMIS during late telophase, and that the Rab11/FIP5-endosomes are transported after AMIS formation around the midbody
(Li et al., 2014b). This 3D-time–lapse microscopy approach has greatly expanded our knowledge of the machinery mediating lumen formation by allowing us to elucidate the timing of certain steps in the overall mechanism. Furthermore, this 3D-time-lapse imaging method can now also be used to further expand our understanding of many molecular processes governing epithelial tissue morphogenesis.
3D-Time-lapse Method
Materials
3D Tissue Culture:
Type II Madin-Darby canine kidney (MDCK) cells.
MDCK media: 50mL fetal bovine serum (FBS) and 5mL penicillin-streptomycin (10,000 U/mL) in
500 mL 1X Dulbecco’s Modified Eagle Medium (DMEM, 4.5 g/L glucose, L-glutamine), filter sterilized.
26 1X Phosphate buffered saline (PBS).
1X 0.25% Trypsin-EDTA.
Matrigel growth factor reduced basement membrane matrix (Corning Life Sciences).
Tissue culture (100mm) and 5 cm gridded glass-bottom (35mm) dishes (Ibidi).
Microscopy:
Inverted Axiovert 200M fluorescent microscope (Zeiss) with 63X oil immersion lens and QE
charged-couple device camera (Sensicam).
Slidebook 5.0 (Intelligent Imaging Innovations) 3D rendering and exploration software.
Detailed instructional method
Plate MDCK cells on 100 mm tissue culture plate in 10 mL MDCK media and let grow for
24 hours at 37°C. The plated cells should not be fully confluent since cells need to be in growth
phase. Typically, cells are at 50-70% confluency after 24-hour incubation when initially plated at
30% confluency.
One to two hours before plating cells take an aliquot of Matrigel and set in on ice to
thaw out. It is very important to keep Matrigel cold even while thawing, since it rapidly solidifies at room temperature. Aspirate media and rinse cells with 10 mL PBS. This is a key step, as leaving some of the serum-supplemented media will inhibit Trypsin and will make very difficult
to lift individual MDCK cells. Add 2 mL 0.25% Trypsin-EDTA and let sit at 37°C for 10-15 minutes.
MDCK cells are usually difficult to dislodge, thus, if needed, they can be incubated for 20-25 minutes. Dislodge cells by gently tapping at the side of 100 mm dish. Harvest cells by adding 8 mL MDCK media to plate and transferring cells to 15 ml tube. Sediment cells by centrifugation at
1000xrpm for 3 minutes. Aspirate media and resuspend cells in 1 mL of MDCK media by pipetting
up and down 50-100 times using 1 mL pipette tip. Pipetting up and down many times helps to
separate cells. It is crucial for this assay to embed individual cells in Matrigel.
27 Embedding cell clumps will lead to formation of cell aggregates with multiple lumens. Pipetting
cells 50 times with a 1 mL pipette usually gives a maximum number of individual cells; however,
this number may vary. Thus, it is advisable to initially pipette cells varying number of times,
while monitoring the efficiency of cell separation.
Count cells to determine number of cells/mL and add 20,000 cells to 25 μL MDCK media
in 1.5 mL Eppendorf tube. The optimum number of cells to plate may vary. Plating too many
cells may cause clumping of dividing cells and overlap of fluorescence, making it difficult to
distinguish single cells and observe them over time. Plating too few cells will make it more
difficult to find single cells that are entering metaphase. Testing a range of 10,000-50,000 cells
to find the optimum number of cells for each assay is recommended.
Make a 75% Matrigel solution by adding 75 μL Matrigel to the 25 μL cell solution from
resuspension. The percentage of Matrigel may vary between 25-80%. The volume of media to suspend cells in and volume of Matrigel added can be adjusted to determine ideal conditions. In lower concentrations of Matrigel, cells may sink through the Matrigel matrix (during process of
solidifying the Matrigel) and stick to the bottom of the plate, counteracting the function of the
Matrigel to observe the cells suspended in the matrix. If the Matrigel concentration is too high,
the cells might not be able to suspend in the matrix and instead would just sit on top, which is
again not ideal for 3D imaging. Gently mix by pipetting up and down and immediately plate
cell/Matrigel mix by placing a drop onto center of 5 cm gridded glass-bottom dish. This step needs to be done quickly, since Matrigel-cell mixture solidifies readily at room temperature. Let
the solution solidify in tissue culture incubator at 37°C for 30 minutes.
Add 5 mL MDCK media to dish and let cells acclimate for 6-12 hours at 37°C. Typically
cells will start dividing within 12 hours after embedding, therefore imaging them within a 6-12-
hour time period will ensure that at least some cells will be undergoing first cell division. Mount
28 the dish on a fluorescent microscope and use a gridded chart to label the location of several
individual cells, especially those that may be in metaphase (starting division). Isolated cells will
be more ideal for imaging. As nearby cells divide and form cysts, the fluorescence can interfere
with the cells being followed. Adjust the focus to set top and bottom of each cell and take initial
0.2 μm-step z-stack images. To be sure to include the whole cell in the range of imaging from
top to bottom, it is best to set the focus a little beyond the point when the cell first begins to blur.
Repeat imaging according to time frame for desired observation. This will result in
taking a mini-z-stack for every time point. Generally, it is best to avoid taking more than 50 time-
lapse z-stacks. Taking fewer time-points or fewer images in each mini-z-stack will decrease
photo-damage to cells. Excessive imaging can be a problem, since mitotic cells are especially
sensitive to photo-damage. Shorter exposures will also reduce photo-bleaching. On the other
hand, using long time-lapses increases the risk of missing the critical steps of the observation.
To visualize AMIS formation during cell division, we typically use 10 minute time-lapse.
To visualize lumen formation and expansion, we use 30-minute time-lapses. Finally, if the
motility of individual Rab11/FIP5-endosomes needs to be observed, we use 200-500 ms time-
lapse. Use the grid etched in the glass dish to locate various cells for imaging at different time
points. Taking mini-z-stacks at every time point allows the use of post-acquisition image analysis
to generate three-dimensional images of MDCK cells at each time point during lumen formation.
Alternatively, individual images that best represent the lumen formation dynamics can be
selected and displayed/analyzed for every time point.
29 Results
Midbody and central spindle microtubules mediate AMIS formation and FIP5-endosome transport during telophase
To understand the spatiotemporal dynamics of FIP5-endosome transport during apical lumenogenesis, I first investigated the localization of FIP5 during AMIS formation. MDCK cells were cultured in Matrigel for 24 h and stained for FIP5 and acetylated-tubulin. FIP5 was present at the midbody at late telophase (Figure 2.1A), suggesting that apical endocytic carriers containing
FIP5 (referred to as FIP5-endosomes hereafter) may be involved in apical protein transport to the midbody during cell division. Interestingly, the AMIS marker Cingulin was localized at the cleavage furrow on both sides of the midbody. Three-dimensional reconstruction of the image (square in
Figure 2.1B) showed that central spindle microtubules were surrounded by a ring of Cingulin at the midzone (Figure 2.1C, D), which implies that the AMIS forms around the midbody at late telophase.
To visualize the dynamics of AMIS formation and FIP5-endosome delivery during cell division, time-lapse microscopy was performed using MDCK cells expressing Cingulin-GFP or FIP5-
GFP respectively. Cingulin was cytosolic during metaphase and anaphase, and became enriched in a ring pattern at late telophase around the midbody before the expansion of the forming lumen
(Figure 2.1E). Compared to Cingulin, FIP5-endosomes accumulated around the centrosomes during metaphase and anaphase (Figure 2.1F) (Hobdy-Henderson et al., 2003), and gradually translocated to the midbody during telophase (Figure 2.1G). Interestingly, FIP5 movement often occurred in an asymmetric fashion, with FIP5 translocating from one centrosome first (Figure
2.1G; 19 min and 59 min), followed by apical lumen initiation and the movement of FIP5- endosomes from the other centrosome (Figure 2.1G; 86 min). To test whether FIP5-endosomes are targeted to the midbody, a FIP3 marker known to be present at the recycling endosomes on
30 the central spindle and midbody was used (Schiel et al., 2011; Schiel et al., 2012). FIP5 co-localized
with FIP3 and the apical marker Crb3a during mid- and late-telophase (Figure 2.2A-D), suggesting
that FIP5-endosomes deliver apical proteins along central spindle microtubules to the midbody.
Additionally, this delivery of apical proteins, marked by gp135, appeared to be preceded by the
formation of the AMIS (Figure 2.2E, F; the AMIS is visualized by ZO1 and Cingulin staining).
To further test whether the AMIS forms before the delivery of FIP5-endosomes to the
lumen formation site, cells co-expressing Cingulin-GFP and FIP5-RFP were imaged. Consistent
with the time-lapse data shown above, Cingulin-GFP accumulated at the midbody during early
telophase (Figure 2.2G). In contrast, during initial AMIS formation, FIP5-endosomes could be
detected accumulating around centrosomes (Figure 2.2G). Only after formation of the AMIS,
during late telophase, are FIP5-endosomes translocated from centrosomes to the midbody.
FIP5-endosome movement to the midbody coincided with the formation and expansion of the
apical lumen (Figure 2.2G), suggesting that AMIS formation around the midbody during late
telophase is likely required to ensure the fidelity of the central spindle-dependent targeting of
FIP5-endosomes.
Discussion
Apical lumen formation has been shown to be a critical step during epithelial
morphogenesis and was shown to be dependent on the establishment of a single AMIS at the
site of lumen formation (Apodaca et al., 2012). What remains unclear is what is the “symmetry-
breaking” event that leads to the establishment of the single AMIS. Here it is shown that the
AMIS is formed around the midbody during late telophase and that FIP5-endosomes containing
apical proteins are delivered to the AMIS along central spindle microtubules. These FIP5-
endosomes fuse with the cleavage furrow PM, which is bordered by the AMIS, thus delivering
apical cargo required for the formation and expansion of the nascent apical lumen (Figure 2.3B).
31 Our data show that FIP5-endosomes translocate from centrosomes to the midbody during cell
division. However, the regulation of FIP5-endosome trafficking in this context remains virtually
unknown. It is also demonstrated that GSK-3 phosphorylates FIP5-T276 during metaphase and
anaphase, thus blocking apical carrier formation by inhibiting FIP5 binding to SNX18 (Li et al.,
2014) (Figure 2.3A). It is likely that this inhibition ensures that apical endocytic carriers are not
transported to the PM before the formation of the AMIS. During late telophase, after the
formation of the AMIS and the dephosphorylation of FIP5-T276, apical endocytic carriers can be
transported along central spindle microtubules and targeted to the site of forming apical lumen.
Consistent with this hypothesis, FIP5-T276D overexpression causes the accumulation of FIP5 and
other apical proteins in enlarged endosomes and prevents the formation of a single apical
lumen (Li et al., 2014). These results suggest that the FIP5-T276 phosphorylation/dephosphorylation cycle is required for the fidelity of apical protein transport.
In support of this hypothesis, GSK-3 inhibition also results in the formation of multiple lumens in
polarized MDCK cysts (Li et al., 2014).
While our data demonstrate that the level of FIP5-T276 phosphorylation decreases as
dividing cells progress from metaphase to telophase, it remains unclear how FIP5-T276
phosphorylation is regulated during cell division. Since the total amount of GSK-3 during mitosis
does not change, it is likely that FIP5-T276 phosphorylation is regulated by modulating GSK-3
activity. Indeed, it has been shown that GSK-3 is active during metaphase and that GSK-3 activity
is required for mitotic chromosomal alignment and the formation of the mitotic spindle (Ong
Tone et al., 2010; Tighe et al., 2007; Wakefield et al., 2003). Thus, it is likely that in addition to
regulating chromosomal alignment GSK-3 also functions in inactivating endocytic recycling
during metaphase.
32 In summary, during the process of de novo lumenogenesis, apical lumen formation is
initiated by FIP5-dependent transport of apical membrane components to the AMIS during
cytokinesis (Figure 2.3A). During metaphase/anaphase FIP5-T276 phosphorylation inhibits FIP5
from binding SNX18, thus preventing apical cargo protein from exiting recycling endosomes.
During telophase, after the formation of AMIS around the midbody, FIP5-T276
dephosphorylation allows the SNX18-dependent formation of apical endocytic carriers (FIP5-
endosomes). These FIP5-endosomes contain apical cargo, including Crb3a and gp135 and move
along central spindle microtubules to fuse with the cleavage furrow PM to generate an apical
lumen (Figure 2.3B).
Several recently published studies, along with this work, have begun to decipher the
molecular machinery that regulates and coordinates apical lumen formation during epithelial
morphogenesis. Multiple Rab GTPases, including Rab11, Rab8, Rab3, and Rab27, are required for this complex cellular process (Apodaca et al., 2012). Our work demonstrates the significance
of FIP5, as a Rab11-binding protein, in regulating several steps of apical lumen formation.
Additionally, it is becoming clear that the phosphorylation of FIP5 by GSK-3 facilitates the cross-
talk between endocytic trafficking and other intracellular signaling pathways and activities. This
connects data from several studies to draw a clearer picture of the regulation of the apical
trafficking network. However, many questions remain. It is unclear how and why the AMIS forms
around the midbody during telophase. The mechanism underlying the tethering/sequestering of
FIP5-endosomes at the AMIS remains elusive. Several proteins, such as the Exocyst complex,
Syntaxin3 and Slp2a/4a, recently emerged as potential candidates for the tethering of apical
exocytic carriers to the AMIS, although their relationship with FIP5-dependent transport is
unclear (Galvez-Santisteban et al., 2012). Finally, most of the studies that analyzed the
molecular machinery for apical lumen initiation and maturation have been done using the 3D
33 tissue culture system. Confirmation and further investigation of these mechanisms in animal models will be required to understand the normal and pathological development of epithelial organs in vivo and to find clinical treatment strategies for related human diseases. In addition, more work is needed to determine the upstream activation of lumen formation, including the mechanism of AMIS formation and the specific factors involved in its spatiotemporal regulation.
34
Figure 2.1. Apical membrane initiation site (AMIS) formation around the midbody at late telophase mediates FIP5-endosome targeting during lumen formation. (A, B) Localization of FIP5 (green) at the midbody and Cingulin (CGN, green) around the midbody marked by acetylated tubulin (red) in MDCK cysts. (C,D) 3D reconstruction of the area squared in (B). (E) Dynamics of AMIS formation marked by CGN. (F-G) FIP5 translocation from the centrosomes to the midbody. Arrows point to the midbody or the forming apical lumen. Asterisks mark FIP5 associated with centrosomes. M: midbody. L: lumen. Scale bars: 5 µm.
35
Figure 2.2. FIP5-endosomes deliver apical proteins along central spindle microtubules to the midbody-associated AMIS. (A-F) FIP5 co-localized with FIP3 (A-B) and apical marker Crb3a (C-D) during mid- and late telophase. AMIS formation marked by CGN and ZO-1 staining preceded apical protein transport (gp135) to the midbody (E-F). MDCK cells expressing either GFP-FIP3 (A- B) or GFP-Crb3a (CD) were grown in 3D cultures for 24 h. Arrows point to the midbody. Scale bars: 5 µm. (G) MDCK cells co-expressing Cingulin-GFP and FIP5-RFP were embedded in Matrigel and imaged after 24 h incubation. Cells shown in panels are at different stages of cytokinesis and apical lumen formation. Asterisks mark centrosomes, arrows point to the midbody. Scale bars: 5 µm. Images shown are Z-axis projections (maximum intensity projection) created from Z- stacks (35 images, Z-step 0.25 nm).
36
Figure 2.3. Proposed models of FIP5-endosome trafficking and apical lumen initiation. (A) Model of FIP5-dependent apical protein trafficking during lumen initiation. (B) Model of the role of cytokinesis during apical membrane initiation site (AMIS) formation and apical lumen initiation.
37 CHAPTER III
CINGULIN AND ACTIN MEDIATE MIDBODY-DEPENDENT APICAL LUMEN FORMATION DURING
3 POLARIZATION OF EPITHELIAL CELLS
Abstract
Coordinated polarization of epithelial cells is a key step during morphogenesis that leads
to the formation of an apical lumen. Rab11 and its interacting protein FIP5 are necessary for the
targeting of apical endosomes to the midbody and apical membrane initiation site (AMIS) during
lumenogenesis. However, the machinery that mediates AMIS establishment and FIP5-endosome
targeting remains unknown. Here, Cingulin is identified as a FIP5-interacting protein which localizes to the AMIS and functions as a tether mediating FIP5-endosome targeting. The
machinery mediating AMIS recruitment to the midbody was analyzed and determined that both
branched actin and microtubules are required for establishing the site of the nascent lumen. The
Rac1-WAVE/Scar complex is also shown to mediate Cingulin recruitment to the AMIS by
inducing branched actin formation, and Cingulin directly binds to microtubule C-terminal tails
through electrostatic interactions. With this new data, an updated mechanism for apical
endosome targeting and AMIS formation around the midbody during epithelial lumenogenesis is
proposed.
Introduction
The formation of an apical lumen is a key step during epithelial tissue morphogenesis
and function, and it is now well established that Rab-dependent endosome transport is
responsible for driving individual cell polarity as well as de novo lumen formation (Blasky et al.,
3 Portions of this chapter are published with permission from our previously published article Mangan A, Sietsema D, Li D, Moore JK, Citi S, Prekeris R. Cingulin and Actin Mediate Midbody- Dependent Apical Lumen Formation During Polarization of Epithelial Cells. Nature Communications. 2016: 3;7:12426.
38 2015; Bryant et al., 2010, Bryant et al., 2008; Roignot et al., 2013). Specifically, the Rab11 family
of GTPases were shown to regulate the transport of vesicles carrying apical cargo to the site of
the forming lumen, known as the apical membrane initiation site (AMIS) (Blasky et al., 2015;
Bryant et al., 2010; Datta et al., 2011; Li et al., 2014a,b; Willenborg et al., 2011). The AMIS is a
transient structure that contains many proteins including the Par3/Par6 polarity complex, the
Exocyst complex, and tight junction proteins such as ZO-1 and Cingulin Blasky et al., 2015;
Bryant et al., 2010; Datta et al., 2011; Li et al., 2014b; Willenborg et al., 2011). De novo
formation of a single AMIS is an essential cellular step leading to the proper initiation and
expansion of a single apical lumen (Blasky et al., 2015; Bryant et al., 2010; Li et al., 2014b;
Willenborg et al., 2011). Recent work from our and other laboratories has shown that midbody
formation and midbody-dependent AMIS recruitment during telophase is the first symmetry-
breaking event that determines the time and site of apical lumen formation (Blasky et al., 2015;
Li et al., 2014b). However, the factors involved in AMIS recruitment to the midbody are still
unknown and are the focus of this study.
In addition to midbody-dependent AMIS formation, apical endosome targeting and
fusion at the AMIS is also an important step in apical lumen formation. Previous studies have
begun to identify the mechanisms of apical endosome budding and targeting and have shown
that apical endosome transport is governed by Rab11 GTPase bound to its effector protein
known as Rab11 Family Interacting Protein-5 (FIP5) (Li et al., 2014a, b; Willenborg et al., 2011).
The sequential interactions of Rab11/FIP5 targeting complex with Sorting Nexin-18 (SNX18) and
Kinesin-2 regulate apical endosome formation and transport along central spindle microtubules during the initial steps of lumenogenesis (Li et al., 2014a, b; Willenborg et al., 2011). Although it
is known that these FIP5 vesicles fuse with the plasma membrane at the AMIS, the specific
mechanisms of targeting and tethering of Rab11/FIP5 endosomes to the AMIS are not fully
39 understood. While several proteins, such as synaptotagmin-like proteins Slp2 and Slp4 as well as the Exocyst complex were shown to be required for single lumen formation (Galvez-Santisteban
et al., 2012), it is unlikely that they alone can target endosome transport to the AMIS, since most
of these factors localize and function at other subcellular locations in addition to the AMIS
and/or midbody, thus limiting their ability to serve as AMIS-specific tethers for incoming apical
vesicles.
Here, the machinery that mediates AMIS formation at the midbody is investigated, as
well as the targeting/tethering of apical endosomes during lumenogenesis. Cingulin (CGN) (Citi
et al., 1988) is identified as a FIP5-binding protein and CGN serves as the tethering factor that
ensures the fidelity of apical endosome targeting to the AMIS. CGN also binds to the C-terminal
tails of midbody microtubules, and this CGN and microtubule interaction may play a major role
in recruiting the AMIS to the midbody during late telophase. Finally, a novel and midbody-
dependent role of Rac1-WAVE/Scar-induced actin polymerization during the initial steps of
apical lumen formation is described. In conclusion, a new model of apical lumen formation is
proposed that includes the interaction of polarized endocytic membrane transport, midbody
microtubules, and branched actin cytoskeleton to form a coincidence detection system that
regulates the timing and fidelity of a single apical lumen.
Results
Cingulin is a FIP5 binding protein concentrated at the AMIS
During de novo lumen formation, the AMIS is established at the midbody during late
telophase, marking the site of a future apical lumen (Figure 3.1a) (Blasky et al., 2015; Li et al.,
2014b). Following AMIS formation, Rab11/FIP5 apical endosomes are transported to the AMIS
(Figure 3.1a) (Blasky et al., 2015; Li et al., 2014b). What is not known are the mechanisms that
target Rab11/FIP5 vesicles to the AMIS. To identify these targeting factors I immunoprecipitated
40 FIP5 from polarized Madin-Darby canine kidney (MDCK) cells (Figure 3.2a). Many of the
identified proteins (Figure 3.1b) are already known to regulate apical vesicle transport,
confirming the efficacy of the immunoprecipitation. In fact, SNX18, dynamin-2, and Arp2/3 are
all known to form a complex that is essential for the budding of FIP5 apical endosomes
(Willenborg et al., 2011). Myosin Vb and FIP1 are other known components of apical endosomes
(Baetz et al., 2013; Carson et al., 2013; Jin et al., 2011; Li et al., 2007; Ossipova et al., 2015), and
clathrin is a general vesicle coating factor (Brodsky et al., 2001; Morris et al., 1989).
Interestingly, CGN was also identified as a putative FIP5-binding protein (Figure 3.1b and Figure
3.2a). CGN is a known AMIS marker that is concentrated almost exclusively around the midbody
during late telophase (Li et al., 2014b; Willenborg et al., 2011), thus making it a strong candidate
to serve as a tethering factor mediating Rab11/FIP5 apical endosome targeting.
To further test whether CGN binds to FIP5 I immunoprecipitated GFP-CGN from MDCK
cells stably expressing GFP-CGN. Consistent with our proteomics data, FIP5 co-
immunoprecipitated with GFP-CGN (Figure 3.1c). Since it was previously shown that FIP5-T276
phosphorylation inhibits FIP5 (Li et al., 2014b), I next co-immunoprecipitated CGN with either
FIP5-T276A-GFP or FIP5-T276D-GFP and found that FIP5 phosphorylation mimetic FIP5-T276D
did not bind to CGN (Figure 3.2b). Finally, purified recombinant 6His-FIP5 and 6His-CGN were
also co-immunoprecipitated with anti-FIP5 antibody (Figure 3.1d), further demonstrating that
CGN directly binds to FIP5.
CGN contains three main domains: a large globular head, coiled-coil rod, and short
globular tail (Figure 3.1e) (Cordenonsi et al., 1999). The head domain is known to bind ZO1 and
mediate CGN recruitment to tight junctions (TJs) (Cordenonsi et al., 1999; Stevenson et al.,
1989) and the coiled-coil rod is required for CGN homo-dimerization and mediates binding to
GEF-H1 (Figure 3.1e) (Cordenonsi et al., 1999; Citi et al., 2000). To map the FIP5-binding motif a
41 GST pull-down assay was used and showed that FIP5 binds to the N-terminal portion of the
coiled-coil rod (amino acid region 355-579) (Figure 3.1e). Since CGN(355-579) is located just
after the large globular head domain, I refer to it as the CGN-coiled-coil-1 region (Figure 3.1e).
CGN is required for single apical lumen formation
To assess the role of CGN in lumen formation, I created a doxycycline (dox) inducible
MDCK-shCGN cell line (Figure 3.2c, d), and tested the effects of CGN knock-down (KD) on apical
lumen formation. After embedding MDCK cells in Matrigel for 4 days, dox- cells produced single-
lumen cysts (Figure 3.3a, d) with CGN, FIP5, gp135 and actin outlining the apical domain (Figure
3.3a, Figure 3.4 a, c, i). In contrast, CGN knock-down led to increased formation of cysts with
multiple lumens (Figure 3.3 b, d, Figure 3.4 b, d, e, j). These multiple lumens are likely the result
of apical cargo mistargeting, since CGN depletion did not have any effect on the angle and
positioning of the mitotic spindle (Figure 3.5c-f). Furthermore, even the dox+ cells that
contained a single lumen had defects in apical surface formation. As shown in Figure 3.3c and
Figure 3.4e, MDCK-shCGN cells appear to have a large apical plasma membrane surface that
extends and folds into the luminal space. Significantly, the addition of shRNA resistant GFP-CGN
to dox+ cells rescued the single lumen phenotype as well as the integrity of the apical surface
(Figure 3.3d, Figure 3.4 f, t-test).
Our data suggest that CGN regulates formation of a single apical lumen by affecting
targeting of Rab11/FIP5 endosomes. To test this, I analyzed MDCK-shCGN cells embedded in
Matrigel for 12 hours. Under these conditions, most embedded cells are either in late telophase,
or just completed their first division and formed a nascent mini-lumen between two daughter
cells7. To determine whether CGN knock-down affects Rab11/FIP5 endosome transport, I
stained cells with anti-FIP5 antibodies. As previously reported, FIP5 and gp135 are
predominantly concentrated at the AMIS (Figure 3.3i, k, arrows; Figure 3.4g) as marked by either
42 tubulin or actin. In contrast, in the dox+ treated cells FIP5-endosomes are distributed throughout the daughter cells, indicating that CGN is required for the targeting of FIP5 (Figure
3.3j, l). Consistently, gp135, a well-established FIP5-endosome cargo protein, was also scattered throughout the cytosol (Figure 3.4h). Additionally, in some cells gp135 was observed to be accumulating in enlarged cellular structures that may represent ectopic nascent lumens (Figure
3.4h, arrow).
Surprisingly, CGN knock-down also disrupted actin cytoskeleton localization (Figure 3.3f- h, l). Typically, during lumenogenesis actin clearly marks the surface of the apical lumen (Figure
3.3e, k). In contrast, in dox+ treated cells actin accumulation at the apical plasma membrane is lost, and actin is instead present in distinct foci along the entire plasma membrane (Figure 3.3f- h, l). In some cells, large intracellular structures containing enriched actin were observed (Figure
3.4g, h, asterisks). These intracellular structures are likely either enlarged apical endosomes or ectopic apical lumen-like structures, since they also contain FIP5 (Figure 3.4j, asterisk). CGN knock-down also changes the appearance of actin in 2D cultures, where stress fiber formation is induced at the basal side of the epithelial monolayer (Figure 3.5a, b).
The WAVE/Scar complex is present at the AMIS
Although it was previously shown that CGN and the AMIS are recruited to the midbody during late telophase (Li et al., 2014b), the mechanisms mediating the establishment of the
AMIS at the midbody were unknown. Thus, synchronized MDCK cells at telophase were used to immuno-precipitate CGN. Isolated proteins were then analyzed by mass spectroscopy and compared to those pulled down in an IgG control. Among the proteins identified only in anti-
CGN and not in the control (Figure 3.6a), was identified ZO1, a known CGN-binding protein.
Consistent with our finding that CGN is FIP5-binding protein FIP5 and SNX18 were also identified from anti-CGN antibody precipitates (Figure 3.6a).
43 In addition to expected CGN-interacting proteins Nap1 and Abi2, two components of the
WAVE/Scar complex, were also identified (Figure 3.6a, b). The WAVE/Scar complex is known to be activated by Rac1 and induces the polymerization of branched actin through the activation of
Arp2/3 (Figure 3.6b) (Patel et al., 2008; Steffen et al., 2004; Stradal et al., 2006). To determine if the WAVE/Scar complex may play a role in the recruitment of CGN to the midbody, the localization of Nap1 and Arp3 during division of MDCK cells embedded in 3D matrix were tested.
As shown in Figure 3.6c, Nap1 co-localizes around the midbody during late telophase and forms a ring structure localized to tight junctions after the formation of the nascent lumen (Figure
3.6d, e), mirroring what was previously observed with CGN (Li et al., 2014). Likewise, Arp3 becomes concentrated at the midbody and co-localizes with CGN during AMIS formation (Figure
3.6f). The actin cytoskeleton was also stained during AMIS formation and lumen expansion, and an increase in actin polymerization around the midbody was detected (Figure 3.6g, h and Figure
3.7a). These actin structures emanate from the AMIS, suggesting that they could be actively playing a role in the establishment of the apical lumen. After formation of the nascent lumen, actin is still closely aligned with CGN, marking the apical surface and tight junctions, respectively
(Figure 3.6i). Based on these findings I hypothesized that the midbody-associated WAVE/Scar complex is helping to recruit the AMIS and may also drive apical lumen formation.
Active Rac1 is required for AMIS formation at the midbody
The WAVE/Scar complex is activated by the binding of the small GTPase Rac1 (Patel et al., 2008; Steffen et al., 2004; Stradel et al., 2006). Our finding that WAVE/Scar and therefore
Rac1 may be involved in regulating AMIS formation during late telophase is somewhat unexpected, since it is well-established that RhoA rather than Rac1 drives cleavage furrow formation and ingression during cytokinesis (Glotzer et al., 2004; Piekny et al., 2005; Piekny et al., 2008). Importantly, it was demonstrated that RhoA is inactivated at late telophase to allow
44 for the disassembly of the contractile actomyosin ring (Shiel et al., 2010). This suggests that during late telophase RhoA inactivation is followed by Rac1 activation, thus allowing a transition to Arp2/3-dependent branched actin cytoskeleton. To that end I analyzed the localization of
Rac1 during lumen formation (Figure 3.8a-d). During anaphase, Rac1 is evenly distributed throughout the cytosol of dividing cells, consistent with Rac1 not being involved in the formation of the actomyosin ring (Figure 3.8a). During late telophase Rac1 becomes concentrated at the AMIS (Figure 3.8b) where it co-localizes with filamentous actin and CGN
(Figure 3.8b,c). Upon formation of the nascent mini-lumen, Rac1 remains enriched at the apical plasma membrane (Figure 3.8d), indicating its possible involvement in maintenance of apical polarity. While sub-cellular Rac1 localization can provide clues about its possible function, ultimately GTP-bound Rac1 is what activates WAVE/Scar and induces actin polymerization. To test the localization of active Rac1 a Rac1 biosensor was used (Komatsu et al., 2011). As shown in the Figure 3.8e, in metaphase activated Rac1 appears to be equally distributed along entire plasma membrane. In contrast, once the cell enters telophase, Rac1 becomes activated at the
AMIS (Figure 3.8f, g).
Although these experiments spatially link Rac1 activation to the establishment of the apical lumen, it does not explain the mechanism governing the recruitment and activation of
Rac1 during late telophase. Previous studies have shown that Arf6 is recruited to the midbody during telophase (Schweitzer et al., 2002; Zhu et al., 2005). It was suggested that Arf6 binds and recruits Tiam1, a known Rac1 GEF (Palacios et al., 2002). Consistent with this hypothesis, Arf6 can be observed at the midbody of the MDCK cells during lumenogenesis (Figure 3.8h). I next wanted to determine if Rac1 plays a functional role in the establishment of the apical lumen. It has been previously shown that Rac1 knock-down leads to formation of inverted cysts with the apical pole of the cells facing the ECM, thus lacking a well-defined apical lumen (O’Brien et al.,
45 2001; O’Brien et al., 2006; Monteleon et al., 2012). However, the use of cell lines expressing
Rac1 shRNA is difficult to interpret, since Rac1 is one of the main regulators of actin
cytoskeleton. As the result, instead of shRNA, an inhibitor that specifically targets the Rac1 GEFs
(Tiam1 and Trio) was used. To ensure that inhibition of Rac1 occurred only during the first
mitotic division, MDCK cells were embedded into Matrigel for 4 hours, followed by a 12-hour
incubation with Rac1 inhibitor. After 16 hours, most of the cells successfully complete the first
division, and the CGN-enriched tight junction ring surrounds the nascent mini-lumen, which is
always found between the two daughter cells (Figure 3.9a). In contrast, cells treated with Rac1
inhibitor failed to form the CGN ring and apical lumen between the two daughter cells (Figure
3.9b). Instead, CGN forms a tight junction ring at the side of one of the daughter cells (Figure
3.9b), despite the fact that cells successfully completed the first mitotic division (Figure 3.9b;
arrows mark actin-rich plasma membrane between two nuclei). Occasionally, a small CGN and
actin ring (Figure 3.9c) were also observed forming on the neck of a small bud-like structure on
one of the daughter cells. Finally, in some cases the cells divide, but fail to polarize and form any
tight junctions of nascent mini-lumens (Figure 3.9d).
To further analyze the role of Rac1 in regulating AMIS formation GFP-CGN dynamics during MDCK cell division were observed by time-lapse microscopy. Consistent with previously
published work (Li et al., 2014b), CGN accumulated at the midbody during telophase (Figure
3.9e, arrow). Upon completion of cell division, CGN formed a ring between two daughter cells,
an indication of apical lumen and tight junction formation (Figure 3.9e). In contrast, treatment
of cells with Rac1 inhibitor blocked accumulation of CGN at the midbody and consequently
prevented apical lumen formation and CGN accumulation at the tight junctions (Figure 3.9f and
Figure 3.10a). Since Rac1 activates the WAVE/Scar protein complex I next checked the
localization of Abi2, the core subunit of the complex. Similar to Nap1, GFP-Abi2 also localized to
46 the midbody during telophase (Figure 3.11a), as well as to tight junctions after completion of the
first division and the establishment of the nascent apical lumen (Figure 3.11c). Incubation of
dividing cells with Rac1 inhibitor blocked GFP-Abi2 recruitment to the midbody (Figure 3.11b).
Interestingly, even recruitment of GFP-Abi2 to the tight junctions was affected, although not
completely inhibited (Figure 3.11d). Taken together, these data indicate that Rac1 and
WAVE/Scar-dependent formation of branched actin filaments at the midbody may also play a
role in the correct placement of tight junctions around the midbody and nascent apical mini-
lumen.
Since Rac1 is required for a recruitment of the AMIS to the midbody, Rac1 inhibition
should lead to defects in targeting of apical proteins. To test this, localization of gp135, an
apical plasma membrane protein delivered to the apical lumen via Rab11 and FIP5-endosomes,
was observed. As previously reported (Li et al., 2014; Willenborg et al., 2011) in control cells gp135 is delivered to the midbody-associated AMIS (Figure 3.11e). These gp135-containing
endosomes eventually fuse to form nascent mini-lumen (Figure 3.11f). Inhibition of Rac1 led to a
failure of gp135 targeting to nascent lumen. Instead gp135-containing vesicles accumulated in
multiple clusters within the cytosol (Figure 3.11g). Significantly, in some cells a CGN ring forms at
the neck of small bud-like structures, and gp135-containing vesicles could also be observed at
those necks (Figure 3.11g, arrows), confirming that CGN does function as a factor mediating
apical endosome targeting/tethering. In some cases, after 16 hours the cells underwent two
rounds of division, thus generating small 4-cell cysts. Typically, at this stage gp135 could already
be observed enriched within small mini-lumen (Figure 3.4f). Rac1 inhibition led to the formation
of multiple mini-lumens surrounded by CGN-containing tight junctions (Figure 3.11i). However,
about 45 % of all the cysts contained an inverted polarity, with gp135-containing plasma
membrane facing the Matrigel (Figure 3.11j). These inverted cysts were similar to the ones
47 observed in cells expressing dominant-negative Rac1 mutants (O’Brien et al., 2001; Monteleon
et al., 2012).
If Rac1 is required for correct placement of the apical lumen and tight junctions, one
would predict that inhibition of Rac1 during the first mitotic division should lead to the
formation of mature cysts containing multiple lumens. To test this prediction, embedded cells
were treated with Rac1 inhibitor for 12 hours, then changed the media and allowed the cells to
form mature cysts (4 days) (Figure 3.12a). As reported previously, untreated cells developed into
well-structured, single-lumen cysts (Figure 3.12b, c) with CGN marking tight junctions and actin
outlining the apical domain (Figure 3.12c). The majority of cells treated with Rac1 inhibitor
(Figure 3.12b) formed cysts containing multiple lumens (Figure 3.12d). It is likely that this multi-
lumenal phenotype is caused by incorrect placement of the AMIS (or multiple AMIS-like
structures) since inhibition of Rac1 during the first cell division does affect mitotic spindle
positioning and angle during subsequent cell divisions (Figure 3.5c-f). Surprisingly, after 4-day
incubation no inverted cysts were observed, suggesting that re-activation of Rac1 (after inhibitor
wash-out) was sufficient to correct polarity inversion but not the multi-luminal phenotype. In
contrast, some cells did not form any apical lumen and had no clear CGN-rich tight junctions
(Figure 3.12b, e).
To further confirm that Rac1 pathway regulates apical lumen formation I also treated
cells with the Arp2/3 inhibitor CK-666 to prevent branched actin polymerization. To inhibit
Arp2/3 only during the first cell division I treated newly embedded cells with Arp2/3 inhibitor for
12 hours, then changed the media and allowed the cells to form mature cysts (Figure 3.7b).
Consistent with the involvement of Rac1 in mediating lumenogenesis, Arp2/3 inhibition led to
the formation of cysts containing multiple lumens (Figure 3.7c-e). Again, cysts with ectopic
accumulations of CGN were observed (Figure 3.7d-e, arrows).
48 To test whether Rac1 is required for apical lumen initiation only during the first mitotic
division, MDCK cells were treated with Rac1 inhibitor 24 hours post-embedding (Figure 3.10b).
Surprisingly, treatment with Rac1 inhibitor after the initiation of a lumen during the first cell
division still led to a multi-luminal defect (Figure 3.10c). However, these multi-luminal cysts
were different from the ones formed by inhibiting Rac1. Typically, these cysts contained a single
central primary lumen with one or more small mini-lumens located at the cyst periphery (Figure
3.10d). This indicates that even after the formation of a primary lumen, Rac1 still plays a role in
subsequent polarized cell divisions, and inhibiting Rac1 leads to the formation of multiple small
secondary lumens. This is also consistent with previously published studies demonstrating that
stable Rac1 knock down leads to very dramatic effects on multiple stages of epithelia
polarization (O’Brien et al., 2001).
CGN binding to tubulin mediates targeting to the midbody
While it is now shown that CGN is required for single lumen formation, the mechanism
ensuring the fidelity of CGN recruitment to the midbody is still unknown. The midbody is
composed of a dense network of microtubules that have been shown to transport endosomes
carrying apical cargo to the AMIS (Li et al., 2014a, b). A recent study suggested that CGN can
bind directly to the microtubules (Yano et al., 2013) suggesting that microtubule binding may
target CGN to the midbody. To test this possibility, an in vitro microtubule-binding assay was used to demonstrate that the CGN-head region (aa1-406), but not tail region (aa1015-1203),
binds directly to polymerized microtubules (Figure 3.13a).
To further analyze the mechanism of CGN and microtubule binding, I focused on the
basic structure of microtubules. Microtubules are formed by polymerization of α-tubulin and β-
tubulin heterodimers. Each α and β-tubulin subunit contains an unstructured C-terminal tail
region (CTT) that extends off the surface of polymerized microtubules (Figure 3.13b). These CTTs
49 contain a region rich in negatively charged amino acid residues, which I refer to as the ‘acidic
patch’ (Figure 3.13c). Tubulin CTTs are also susceptible to post-translational modifications, such as glutamylation (Janke et al., 2014), which adds a chain of glutamate residues on a genetically
encoded glutamate residue within the acidic patch and increases the negative charge of CTTs.
While little is known about the functional consequences of CTT glutamylation, midbody
microtubules are also known to be highly glutamylated (Figure 3.14b) (Janke et al., 2011). To determine whether CTTs may mediate CGN binding, subtilisin was used to cleave the CTTs from microtubules (Figure 3.14a) and compared CGN ability to bind CTT-less microtubules (Figure
3.13d). As shown in Figure 3.13d, subtilisin treatment completely eliminated CGN binding to microtubules (Figure 3.13d, lane 6).
It was also important to determine whether CGN can bind to microtubules in which the
CTTs are genetically ablated. Since mammalian cells have many β-tubulin isoforms, yeast was used, as they contain only one gene for β-tubulin and can be engineered to express mutant β-
tubulin that lacks the CTT region (Aiken et al., 2014). I first tested whether CGN can bind to
microtubules made from purified wild-type yeast tubulins. As shown in Figure 3.13e, despite
differences in the primary sequences between human and yeast β-tubulin CTTs, human CGN can
still bind to yeast microtubules. This further supports the putative involvement of the acidic
patch in binding to CGN, since the presence of the acidic patch is preserved across species,
despite sequence differences between human and yeast β-tubulin (Figure 3.13c). The binding of
CGN to mutant yeast tubulin was also tested (Figure 3.13c) and found that deletion of β-tubulin
CTT leads to about 75% reduction in CGN binding (Figure 3.13e), likely due to the loss of the CTT
acidic patch.
Since CGN binds to the CTT acidic patch I hypothesized that the CGN head domain
should contain a basic patch. Indeed, CGN sequence analysis led to an identification of an
50 arginine and lysine-rich sequence that is present in all vertebrate sequences analyzed (Figure
3.13f). In mammals the core sequence of this basic patch consists of a highly conserved 36-
RRGGRR-41 motif (Figure 3.13f, boxes). To test whether this motif mediates microtubule binding
two separate CGN mutants were created, Rs36/37As and Rs40/41As, and analyzed mutant
binding to microtubules (Figure 3.13g). Consistent with the involvement of this basic patch, both
mutants showed a significant decrease in CGN binding to microtubules (Figure 3.13g, t-test).
This result confirms that the binding is regulated by electrostatic interaction between the
negatively charged tubulin tails and the basic patch of the CGN head region. The addition of
charged modifications, such as glutamylation at the midbody, would enhance this interaction
and in our 3D embedded MDCK cells, an enhancement in both glutamylated tubulin around the
midbody, and focused localization around CGN at the AMIS was observed (Figure 3.14b).
All four arginines were also mutated to alanines (CGN-M1/M2-GFP) and the ability of
these CGN mutants to localize at the lumen formation site was tested. However, since CGN is known to homodimerize, there was concern that even in CGN shRNA-expressing cells, remaining
CGN may still target mutants to the appropriate location. To eliminate this possibility,
CRISPR/Cas9 was used to make an MDCK CGN knock-out cell line (MDCK-KO) (Figure 3.15a-c).
Importantly, MDCK-KO recapitulated the multi-luminal phenotype observed in MDCK cells
expressing CGN-shRNA (Figure 3.15d). MDCK-KO cells were then used to analyze the localization
of CGN-M1/M2-GFP mutants during lumenogenesis. As shown in Figure 3.16a-b, in the majority
of cells (73.42+/-2.6% of cells) wild-type CGN-GFP localized to the tight junctions surrounding
actin rich single apical mini-lumens. In contrast, CGN-M1/M2-GFP expressing cells largely failed
to form a single apical lumen (26.58+/-2.8% of normal apical lumens; statistically different from
wilt-type CGN-GFP at p<0.0001; t-test), likely due to the fact that CGN-GFP mutants did not
localize to the site of lumen formation (Figure 3.16c-e). Interestingly, CGN-M1/M2-GFP
51 localization was similar to CGN localization after treatment with Rac1 inhibitor. In some cells
CGN-GFP mutants formed ectopic tight junction-like structures that were also rich in actin
(Figure 3.16c-d), while in other cells CGN-GFP mutants were predominately cytosolic (Figure
3.16e). Together, this data provides a targeted mechanism where cross-talk between actin and
tubulin leads to the recruitment of CGN during establishment of the AMIS and formation of the
apical lumen.
Discussion
Recent work from many laboratories has demonstrated that de novo lumen formation
relies on the targeted delivery of apical proteins to the AMIS, which forms around the midbody
during cell division. It is now well established that these apical proteins are transported via
Rab11/FIP5-containing apical endosomes, and that targeting of these endosomes to the AMIS is
essential for the formation of a single apical lumen. However, while the mechanisms mediating
apical vesicle targeting are beginning to emerge, two major questions remained unanswered:
what factors are involved in establishing the AMIS at the midbody during the first cell division,
and how are apical Rab11/FIP5-endosomes specifically targeted to the AMIS. In this study it is shown that Cingulin (CGN) binds directly to FIP5 and functions as a targeting/tethering factor for
Rab11/FIP5-endosomes during the early stages of lumen formation (Figure 3.17). Our data
supports a model in which both the branched actin cytoskeleton and midbody microtubules play
a role in initial establishment and maintenance of the AMIS. It is also shown that the C-terminal
tails of microtubules bind to a basic patch located in the globular head region of CGN, and that
this microtubule binding is likely to be required for initial CGN recruitment to the midbody
(Figure 3.17). It is also demonstrated that Rac1 activation at the midbody leads to the
recruitment of the WAVE/Scar complex and polymerization of branched actin filaments (Figure
3.17). Importantly, polymerization of these branched actin filaments is also necessary for the
52 formation of the AMIS at the midbody, and is also required for the formation and expansion of a
single apical lumen (Figure 3.17). How Rac1 is activated at the midbody during late telophase
remains unclear, but it is possible that Arf6 mediates localization of the Rac1 GEF Tiam1 at the
midbody. Indeed, Arf6 was shown to be enriched at the midbody (Zhu et al., 2005) and was also
shown to be required for apical lumen formation (Tushir et al., 2007; Tushir et al., 2010).
While Arf6 likely plays a role in activating Rac1 at the midbody-associated AMIS, there
are also other pathways that can regulate actin dynamics during lumenogenesis. CGN itself
recently emerged as a key hub that regulates the actin cytoskeleton. It was shown that CGN
directly binds to GEF-H1, a GEF for RhoA (Guillemot et al., 2014). Additionally, CGN was also
shown to interact with MgcRacGAP (Guillemot et al., 2014), although it remains controversial
whether MgcRacGAP is a GAP for RhoA or Rac1 (Guillemot et al., 2014; Breznau et al., 2015;
Cannet et al., 2014). Finally, CGNL1 is known to bind to CGN as well as Tiam1, thus providing
another pathway of activating Rac1 at the AMIS (Guillemot et al., 2008). Consistent with these
observations, CGN, RhoA, and Rac1 were shown to be important for tight junction stability in
polarized epithelial cells (Guillemot et al., 2014; Breznau et al., 2015).
In addition to regulating the localization of the AMIS, midbody-associated branched
actin is also likely involved in mediating apical endosome targeting. Myosin-Vb is well known to
be required for apical lumen formation and is present at the Rab11/FIP5-endosomes (Jin et al.,
2011; Li et al., 2007; Ossipova et al., 2015). Consistent with that, Myosin-Vb was also isolated as a protein that is associated with Rab11/FIP5 protein complex (Figure 3.2a). It has been proposed
that Myosin-Va, a closely-related Myosin-Vb isoform, functions as a tether, sequestering
melanosomes at the actin-rich cellular protrusions (Wu et al., 2001). Thus, it is likely that
midbody-associated actin flares may also help to sequester Myosin-Vb containing Rab11/FIP5-
endosomes at the midbody and AMIS.
53 Taken together, a new model of how the AMIS is recruited and maintained at the
midbody during cell division is proposed (Figure 3.17). The requirement of both, branched actin and glutamylated microtubules serves as a coincidence detection system that allows for the
establishment of a single apical lumen. Similarly, targeting of apical endosomes to the AMIS also depends on several factors. It was previously demonstrated that the Exocyst complex and Slps
are required for the formation of a single apical lumen (Jing et al., 2009; Mellman et al., 2008;
Wu et al., 2005). Here it is shown that CGN and branched actin are also required for Rab11/FIP5
targeting. Thus, it is becoming clear that lumen formation is dependent on a combination of
overlapping pathways that occur at the right place and the right time to mediate lumen
formation. This ever-growing complexity of apical lumen formation pathways may also be
needed to allow for tissue specific differences in the dynamics and timing of apical lumen
formation. Indeed, mammary epithelial tissues appear to rely more on apoptosis-dependent
lumen formation (known as cavitation) (Debnath et al., 2005). Consistent with that recent work
has shown that Rac1 may not play a key role in lumen formation during mammary tissue
morphogenesis in vivo (Akhtar et al., 2013).
Although this study has greatly expanded our knowledge of lumen formation, there are
still many questions left to explore. Most of the de novo lumen formation studies have been
conducted using in vitro models, thus it remains to be understood how apical endosome
targeting and AMIS formation machinery function within the context of much more complex in
vivo models. Additionally, during tissue morphogenesis, apical lumen formation relies not only
on initiation of nascent mini-lumen, but also on a coalescence of these mini-lumens to form a
final functioning apical space. While lumen coalescence clearly involves extensive remodeling of
apical and basolateral plasma membrane, the mechanisms governing coalescence remain largely
54 unclear. Thus, future studies will be needed to dissect the roles of CGN and FIP5 during the formation and coalescence of a single apical lumen in vivo.
Methods
Plasmids and antibodies
Rabbit polyclonal anti-CGN antibodies were prepared (Willenborg et al., 2011) using recombinant purified human CGN head (aa1-406) and tail (aa1015-1203) fragments
(Proteintech, Chicago, IL). Antibodies were affinity purified using recombinant CGN head and tail fragments conjugated to Affi-Gel 10 resin (Bio-Rad Laboratories, Hercules, CA) and eluted with
0.1 M glycine buffer, pH 2.5. Previously made rabbit polyclonal anti-FIP5 antibodies were also used (Willenborg et al., 2011; Prekeris et al., 2000). Monoclonal mouse anti-acetylated tubulin antibodies were purchased from Sigma-Aldrich (St. Louis, MO). Rabbit anti-Nap1 antibodies were purchased from NOVUS Biologicals (Littleton, CO). Mouse monoclonal anti-glutamylated tubulin antibodies were purchased from AdipoGen Life Sciences (San Diego, CA). AlexaFluor-
594- and AlexaFluor-488-conjugated anti-rabbit and anti-mouse secondary antibodies were purchased from Jackson ImmunoResearch Laboratories (West Grove, PA). AlexaFluor-568- phalloidin was purchased from Life Technologies (Carlsbad, CA). The IRDye 680RD Donkey anti- mouse and IRDye 800CW donkey anti-rabbit secondary antibodies used for western blotting were purchased from Li-Cor (Lincoln, NB).
The GFP-Nap1 and Arp3-GFP were received from the lab of Giorgio Scita (University of
Milan) (Innocenti et al., 2004). GFP-CGN was received from the lab of Sandra Citi (University of
Geneva) (Paschoud et al., 2011). The Rac1-biosensor was received from the lab of Michiyuki
Matsuda (Kyoto University) (Komatsu et al., 2011). CFP-Rac1 was created by cloning Rac1 cDNA into a pECFP-C1 vector (Clontech, Palo Alto, CA). Antibody concentrations ranged from 0.1 - 0.5 mg/mL, and were used at a 1:100 dilution.
55 CGN knock-out by CRISPR/Cas9 in MDCK cells
The guide sequence targeted to exon 2 of canine CGN (Figure 3.15a) was cloned in
lentiCRISPR vector that also contains Cas9 ORF (Ran et al., 2013). MDCK cells were then
transfected and cells expressing Cas9 and CGN guide were selected using puromycin. Multiple
MDCK clonal cell lines were isolated and tested for the absence of CGN using western blotting. A
cell line that did not contain any detectable endogenous CGN (Figure 3.15b-c) was then chosen
for further analysis. This cell line was then genotyped by amplifying and sequencing the CRISPR
target region and was found to have mutations in both copies of CGN leading to a premature
STOP codon (Figure 3.15a).
Immunoprecipitation and proteomic analysis
Immunoprecipitation of FIP5 and CGN was performed using MDCK cell lysates
(Willenborg et al., 2011). Cell lysates were harvested in phosphate buffered saline (PBS)
containing 1% Triton X-100 and phenylmethylsulfonyl fluoride (PMSF). Lysates were then
incubated overnight at 4oC with anti-FIP5 antibody, anti-CGN antibody or non-specific rabbit IgG
control conjugated to protein A-sepharose beads (Sigma-Aldrich (St. Louis, MO)). The beads were then pelleted and washed. To identify interacting proteins, immunoprecipitats were eluted
with 1% SDS and separated on 7-15% SDS/PAGE gradient gel. Distinct bands present only in anti-
FIP5 immunopreipitate but not IgG control were cut out and analyzed by mass spectroscopy
(Woodman et al., 2014). Briefly, gel pieces were destained in 200 μL of 25 mM ammonium bicarbonate in 50% acetonitrile (ACN) for 15 min and washed twice with 200 μL of 50% ACN.
Disulfide bonds in proteins were reduced by incubation in 10 mM DTT at 60 °C for 30 min,
cysteine residues were alkylated with 20 mM iodoacetamide in the dark at room temperature
for 45 min. After 100 ng of trypsin was added to each sample to each sample and allowed to
rehydrate the gel plugs at 4 °C for 45 min and incubated at 37 °C overnight. Samples were
56 measured on a Velos Orbitrap mass spectrometer (Thermo Fisher Scientific) coupled to an
Eksigent nanoLC-2D system through a nanoelectrospray LC−MS interface. Pep8des were
separated on a house-made 100 μm inner diameter × 150 mm fused silica capillary packed with
Jupiter C18 resin (Phenomex). Data acquisition was performed by using the instrument supplied
Xcalibur (version 2.0.6) software. The mass spectrometer was operated in the positive ion
mode; the peptide ion masses were measured in the Orbitrap mass analyzer, whereas the
peptide fragmentation was performed by using either higher energy collisional dissociation
(HCD) or electron transfer dissociation (ETD) in the linear ion trap analyzer by using default
settings. Ten most intense ions were selected for fragmentation in each scan cycle; fragmented
masses were excluded from further sequencing for 90 s. MS/MS spectra were extracted from
raw data files and converted into Mascot generic format (MGF) files by using a PAVA script
(University of California, San Francisco). These mgf files were then independently searched using
an in-house Mascot server (Version 2.2.06, Matrix Science). Mass tolerances were ±15 ppm for
MS peaks, and ±0.6 Da for MS/MS fragment ions. For all HCD spectra, fragment ion tolerances
were set to 0.05 Da. Trypsin specificity was used, allowing for one missed cleavage. Met
oxidation, protein N-terminal acetylation, and peptide N-terminal pyroglutamic acid formation were allowed for variable modifications while carbamidomethyl of Cys was set as a fixed modification. To identify CGN-binding proteins, immunoprecipitats were eluted from the beads
with 0.1M glycine pH 2.0 and prepped for mass spectroscopy in a similar manner, without the
gel extraction steps and starting with the addition of trypsin.
Protein expression and purification
Full-length 6His-FIP5 and 6His-CGN were produced in baculovirus using the transfer
vector pVL1392 (Willenborg et al., 2011). In brief, 106 Sf9 cells were seeded into a 6-well plate,
and the Bacfectin–DNA mixture was added dropwise. After 5 d, the P1 viral stock was harvested
57 and further amplified to P2 and P3 stages. For protein production, 1L of Sf9 cells at 2 million
cells/ml were infected with 2 ml of P3 viral stock (approximate MOI of 0.5) and harvested after
65 h. Cells were lysed in 50 mM Tris buffer, pH 7.5, containing 300 mM NaCl, and the cleared
lysate was loaded onto a Ni-NTA column. Eluted 6His-FIP5 or 6His-CGN were dialyzed overnight
against buffer (50 mM Tris, pH 7.5, 300 mM NaCl, and 5 mM BME) and frozen in liquid nitrogen.
Yields were typically 3–5 mg/liter with an estimated purity of >75%. GST-CGNaa1-406, GST-
CGNaa355-579, GST-CGNaa571-794, and GST-CGNaa781-1025 fragments were expressed using
the pGEX-4T plasmid (provided by Sandra Citi) (Cordenonsi et al., 1999) and purified using the
BL21-(FE3) RIPL Escherichia coli strain (Willenborg et al., 2011). Briefly, E. coli were lysed using a
French press and then incubated with glutathione agarose beads (Sigma-Aldrich). Beads were
then washed with PBS and the GST-protein eluted with 25 mM glutathione (GE Healthcare).
Final protein concentrations were determined using Bradford protein assay (Bio-Rad
Laboratories).
Glutathione bead pull-down assays
GST pull-down assays were performed (Willenborg et al., 2011) Glutathione beads (50
μL) were coated with 10 μg GST fusion protein or GST control in PBS and incubated with
specified amounts of soluble recombinant protein or MDCK cell lysates (PBS, 1% Triton X-100) in
a final volume of 0.5 ml of reaction buffer (50mM Hepes, pH 7.4, 150mM NaCL, 5mM MgCl2,
0.1% Triton X-100, 0.1%% bovine serum albumin, and 1mM PMSF). Samples were incubated at
25°C for 1 h while rotating, pelleted by centrifugation at 2000g for 3 minutes, rinsed with
reaction buffer (1mL x 3), and bound proteins were eluted with 1% SDS, analyzed by SDS-PAGE,
and stained with Coomassie blue or immunoblotted and scanned using a Li-Core Odyssey
scanner.
58 Cell culture
Parental MDCK-II cells (ATCC, Manassas, VA) were tested for mycoplasma, cultured in
DMEM with glucose, L-glutamine, 10% FBS and supplemented with penicillin and streptomycin.
In 2D cultures, MDCK cells were grown on collagen-coated Transwell filters for 4 days. 3D cultures of MDCK cells (Willenborg et al., 2011) actively dividing were mixed in media to create a
25% Matrigel solution that was plated on 8-well slides (about 12 μL/well). The Matrigel–cell mixture was allowed to harden for 30 min at 37°C, and 400 µl of medium was added. The cells were incubated for the indicated period of time and the media was changed every other day. 3D cell cultures were stained according to a modified previously published protocol (Debnath et al.,
2005). In brief, 3D cultures were fixed with 3% paraformaldehyde for 20 min, permeabilized with PBS and 0.5% Triton X-100 for 10 min, and quenched three times for 15 min each wash with a glycine/PBS solution (130 mM NaCl, 7 mM Na2HPO4, 3.5 mM NaH2PO4, and 100 mM glycine). Cells were incubated in primary block (10% FBS, 130 mM NaCl, 7 mM Na2HPO4, 3.5 mM
NaH2PO4, 7.7 mM NaN3, 0.1% BSA, 0.2% Triton X-100, and 0.05% Tween-20) for 4 h, followed by incubation in secondary block (primary block with 20 µg/ml goat anti–mouse F(ab’)2 fragments) for 1 h. After washing, cells were left overnight in primary block with primary antibody and
Hoerscht nuclear stain. Cells were then washed and incubated for 1 h with secondary antibody in primary block. Cells were washed, dried for 1 h, and mounted with VectaShield.
Immunofluorescent, time-lapse, and FRET microscopy
All fixed cells were imaged with an inverted Axiovert 200M microscope (Carl Zeiss) with a 63× oil immersion lens and QE charge-coupled device camera (Sensicam). Z-stack images were taken at a step size of 100–500 nm. Image processing was performed using 3D rendering and exploration software Slidebook 5.0 (Intelligent Imaging Innovations).
59 For live-imaging, about 50,000 GFP-CGN MDCK cells were embedded in a 25% Matrigel
solution and plated on a glass bottom dish, and 200 μM Rac1 inhibitor was added to treated
samples. For imaging, the dishes were placed in a heat- and humidity-controlled chamber (37°C,
5%CO2/95%air) on the stage of an inverted Zeiss LSM510 Meta confocal microscope. Cells were
brought in focus using a 10X objective, and z-stack images (2 μm steps) were acquired every 30 minutes for a 12-24 hour period using a 63X oil objective.
For Florescence resonance energy transfer (FRET) analysis cells were transfected with
Raichu-Rac1 biosensor and embedded into 3D Matrigel matrix. After 24 hour cells were then
imaged and corrected FRET (cFRET) was calculated using Intelligent Imaging Innovation three-
dimensional rendering and exploration software (Sorkin et al., 2000) with the equation
cFRET=FRET-0.4 X CFP-0.037 X YFP.
MDCK cell lines
Tet-inducible shRNA cell lines were created (Willenborg et al., 2011) using canine CGN-
shRNA sequences (5’-GATCCCCAGAGCATGTTCCAGAAGAATTCAAGAGATCTTTCTGGAACATGCTC
TTTTTTA-3’) cloned into the pHUSH retroviral expression vector, transfected into MDCK cells
using Lipofectamine 2000 (Invitrogen), and grown in media supplemented with tet-free FBS
(Takara Bio Inc.) and 1 µg/ml of puromycin. Selected colonies were then grown in the presence
of 1 µg/ml of doxycycline for 72 h and tested for knock-down by western blotting. The GFP-CGN
MDCK cell line was generated as previously described (Paschoud et al., 2008).
Rac1 inhibition studies
The Rac1 inhibitor (NSC 23766) was purchased from Santa Cruz Biotechnology. Cells embedded in 3D Matrigel were allowed to adjust to media for 2 hours and treated with 200 µM
inhibitor. For 4 day cysts, the inhibitor was removed after 12 hour treatment by replacing
media.
60 Yeast tubulin purification
Saccharomyces cerevisiae (budding yeast) strains yJM0596/ YEF473A MATa ura3-52 lys2-801 leu2-∆1 his3-∆200 trp1-∆63 (wild type), and yJM0282 MATa tub2-430∆ +331::TRP1 ura3-52 lys2-801 leu2-∆1 his3-∆200 trp1-∆63 (Beta tailless) (Aiken et al., 2014) were used as a
tubulin source. Single colony isolates of each strain were selected and cultured individually in
5ml of rich yeast extract-peptone-dextrose (YPD) media overnight at 30°C. 300μl of the
saturated overnight culture was used to inoculate 10 liters of rich media (YPD) and grown to log
phase at 30°C ~24 hours after inoculation. The culture was harvested by centrifugation and
pelleted (3500 rpm, 15 min at 4°C, J-6B; Beckman Coulter, Brea, CA). This yielded between 120-
140 grams of wet cell pellet. Pellets were frozen at -80° until ready to purify. Purification was performed 57 with the addition of an anion exchange chromatography step. After the budding
yeast tubulin was eluted off the TOG column it was dialyzed overnight in 2 liters of BRB80 pH
6.9, 50μM GTP. After dialyzing the purified budding yeast tubulin anion exchange
chromatography was preformed using HiTrap Q HP (1 ml = 1 column volume [CV]; GE
Healthcare, Buckinghamshire, United Kingdom) pre-equilibrated in BRB80 pH 6.8 (wash buffer)
at 1 CV/min. The tubulin was eluted using 1M NaCl in wash buffer. Peak fractions were
determined by Bradford assay and pooled and dialyzed in BRB80, 50μM GTP pH 6.8. The
concentration of the tubulin was determined by measuring the absorbance at 280 nm using a
NanoDrop spectrophotometer (NanoDrop Technologies, Wilmington, DE) and the calculated
extinction coefficient and molecular weight. Glycerol was added to 10% before the tubulin was
aliquoted, snap frozen in liquid nitrogen, and stored at −80ºC.
Microtubule binding assays
The microtubule binding assay was performed according to Microtubule Binding Protein
Spin-down Assay Kit (Cytoskeleton). Briefly, 20 µL of taxol-stabilized microtubules (0.5 mg/mL)
61 were incubated with 2 µg test protein and 40 µg BSA in 200 µL general tubulin buffer (80 mM
PIPES pH 7.0, 2 mM MgCl2, 0.5 mM EGTA) for 30 min. The solution was placed on top of 100 µL
cushion buffer (80 mM PIPES pH 7.0, 1 mM MgCl2, 1 mM EGTA, 60% glycerol), spun-down at
100,000 g, supernatant removed, and the pellet resuspended in 1X loading buffer. The binding
was analyzed by SDS-PAGE and stained with Coomassie blue or transferred for western blotting.
For yeast tubulin binding, the microtubules were polymerized at 30 degrees Celsius in a water
bath and the rest of the experiment carried out at the same temperature in the same manner.
Western blots
Some western blots are shown in their entirety in the Figures. Whenever modified, all
uncropped western blots can be found in Figure 3.18.
62
Figure 3.1. Cingulin (CGN) is an AMIS-associated FIP5-binding protein.
63 Figure 3.1. Cingulin (CGN) is an AMIS-associated FIP5-binding protein. (a) Schematic model showing the formation of a nascent apical lumen. Rab11/FIP5 endosomes are transported along central spindle microtubules and fuse to the apical membrane initiation site (AMIS) present at the midbody during late telophase. (b) List of proteins identified in FIP5 immunoprecipitate by mass spectroscopy analysis. Proteins known to be present in the same complex are highlighted together. (c) FIP5 co-immunoprecipitation with GFP-CGN from MDCK lysates. Images shown are the immunoblots after probing with anti-CGN (top gel) or anti-FIP5 (bottom gel) antibodies. (d) Co-immunoprecipitation of purified 6His-CGN with 6His-FIP5. Images shown are the immunoblots after probing with anti-CGN (top gel) or anti-FIP5 (bottom gel) antibodies. (e) Mapping 6His-FIP5 binding domain using glutathione bead pull-down assays. Image shown is Coomassie stained SDS-PAGE gel.
64
Figure 3.2. CGN is a FIP5-interacting protein. (a) Coomassie-stained SDS-PAGE gel of FIP5 immunoprecipitates. (b) Comparison of GFP-FIP5 mutant co-immunoprecipitation with anti- CGN antibody from MDCK cells transiently expressing either FIP5-T276A-GFP or FIP5-T276D- GFP. (c) MDCK cells stably expressing tet-inducible CGN shRNA were grown in the presence or absence of doxycycline for 4 days. Cell lysates were then analyzed by immunoblotting with antiCGN (left gel) and anti-FIP5 (right gel) antibodies. (d) MDCK cells stably expressing tet- inducible CGN shRNA were grown on Transwell filters in the presence or absence of doxycycline for 4 days. Cells were then fixed and stained with antiCGN antibodies and DAPI.
65
Figure 3.3. Cingulin is required for the formation of a single apical lumen.
66 Figure 3.3. Cingulin is required for the formation of a single apical lumen. MDCK cells stably expressing CGN shRNA were embedded in 3D Matrigel and allowed to grow for 4 days (a-d) or 24 hours (e-l) in the presence (dox+) or absence (dox-) of doxycycline. Cells were then fixed and stained with phalloidin-Alexa594 (a-c, e-h, k, and l; red), anti-CGN (a-c and e-h; green), anti- tubulin (i and j; red) or anti-FIP5 (i and j; green) antibodies. Asterisks in (g), (h), and (j) mark ectopic lumen formation sites. Arrows in (k) point to the AMIS. Panel (d) shows quantification of cysts with one, two, or multiple lumens. Data shown are means and standard deviations of three independent experiments. Consecutive images without an individual letter label show the same cell.
67
Figure 3.4. CGN knock-down affects apical lumen formation.
68 Figure 3.4. CGN knock-down affects apical lumen formation. (a-b) MDCK cells stably expressing tet-inducible CGN shRNA were grown in the absence (a) or presence (b) of doxycycline for 4 days. Cells were then fixed and stained with phalloidinAlexa594 (red) and FIP5 (green). (c-e) MDCK cells stably expressing tet-inducible CGN shRNA were grown in the absence (c) or presence (d,e) of doxycycline for 4 days. Cells were then fixed and stained with phalloidin- Alexa594 (red) and anti-β-catenin (green). (f) MDCK cells stably expressing tet-inducible CGN shRNA were transfected with human GFP-CGN (green) and grown in the presence of doxycycline for 4 days. Cells were then fixed and stained with phalloidin-Alexa594 (red). (g-j) MDCK cells stably expressing tet-inducible CGN shRNA were grown in the absence (g,i) or presence (h,j) of doxycycline for either 12 hours (g,h) or 3 days (i,j). Cells were then fixed and stained with anti- gp135 antibodies. Arrow in (h) points to ectopic lumen. Consecutive images without an individual letter label show the same cell.
69
Figure 3.5. CGN depletion affects actin cytoskeleton in MDCK cells.
70 Figure 3.5. CGN depletion affects actin cytoskeleton in MDCK cells. (a-b) MDCK cells stably expressing tet-inducible CGN shRNA were grown on Transwell filters in the presence (b) or absence (a) of doxycycline for 4 days. Cells were then fixed and stained with phalloidin-Alexa594 (red). Shown images are taken at apical (left images), lateral (midimages) and basal (right images) poles of polarized MDCK cells. (c-f) MDCK cells stably expressing tet-inducible CGN shRNA were embedded in Matrigel and grown for 24 hours in the presence (d) or absence (c and e) of doxycycline. During 24 hour incubation majority of cells undergo either 1 or two cell divisions, thus generating 3 or 4 cell organoids. Where indicated (e) cells were treated with Rac1 inhibitor for the first 12 hours (during first cell division). Images marked with the same letter/number show the same cell. To analyze the second cell division angle, cell organoids were randomly chosen and angle measured between long axis of interphase cell and division plane of cells in late cytokinesis (lines in c-e). Panel (f) shows quantitation of angle from cells either depleted of CGN or treated with Rac1 inhibitor.
71
Figure 3.6. Components of the WAVE/Scar complex and branched actin filaments are present at the midbody during lumen formation. (a) List of proteins identified in CGN immunoprecipitate by mass spectroscopy. Proteins known to interact as a complex are highlighted together. (b) Schematic representation of the WAVE/Scar complex. (c-i) MDCK cells were embedded in 3D Matrigel and allowed to grow for 24 hours. Cells were then fixed and stained with phalloidin-Alexa594 (g-i; red), anti-CGN (f, g, and i), anti-acetylated tubulin (c; red and h; green) or anti-Nap1 (c-e; green) antibodies. In panel (f), cell is expressing Arp3-GFP. White arrows identify structures as labeled. Consecutive images without an individual letter label show the same cell.
72
Figure 3.7. Midbody and Cingulin associated actin flares. (a) 3D-rendering images of midbody regions stained with phalloidin-Alexa594 (red) and anti-CGN (green) antibodies. Images shown are derived from cells depicted in Figure 3g. (b) Schematic representation showing timing of 12- hour treatment with Arp2/3 inhibitor CK- 666. (c-e) MDCK cells were embedded in 3D Matrigel and allowed to grow for 24 hours. Where indicated, cells were treated with 200 μM Arp2/3 inhibitor CK-666 (d-e). Cells were then fixed and stained with phalloidin-Alexa594 (red) and anti- CGN (green) antibodies. Panel (c) shows quantitation of cysts with single lumen, multiple lumens or no lumen. n indicates number of cysts analyzed. Arrows point to ectopic CGN accumulations that associate with actin cytoskeleton.
73
Figure 3.8. Rac1 is activated at the AMIS.
74
Figure 3.8. Rac1 is activated at the AMIS. (a-d) MDCK cells transiently expressing CFP-Rac1 (green) were embedded in 3D Matrigel and allowed to grow for 24 hours. Cells were then fixed and stained with phalloidin-Alexa594 (a, b and d; red) or anti-CGN (c; red) antibodies. (e-g) MDCK cells transiently expressing FRET-based Rac1 biosensor were embedded in 3D Matrigel and imaged either at metaphase (e) or telophase (f). Panel (g) shows quantification of FRET signal at the AMIS (area #1) and cell periphery (area #2) during telophase. Data shown are the means and standard deviations derived from 6 cells. (h) MDCK cells transiently expressing CFP- Arf6 (red) were embedded in 3D Matrigel and allowed to grow for 24 hours. Cells were then fixed and stained with anti-acetylated tubulin (red) antibodies. Arrows mark the midbody. Consecutive images without an individual letter label show the same cell.
75
Figure 3.9. Rac1 inhibition affects midbody-associated AMIS formation.
76 Figure 3.9. Rac1 inhibition affects midbody-associated AMIS formation. (a-d) MDCK cells were embedded in 3D Matrigel and allowed to grow for 24 hours. Where indicated, cells were treated with 200 μM Rac1 inhibitor (b-d). Cells were then fixed and stained with phalloidin- Alexa594 (red) and anti-CGN (green) antibodies. White boxes indicate area zoomed in and shown in far right box. White arrows point to nascent lumen, black arrows point to areas of actin concentration. Consecutive images without an individual letter label show the same cell. (e-f) MDCK cells stably expressing GFP-CGN were embedded in 3D Matrigel. Untreated (e) or Rac1 inhibitor treated (f) cells were then imaged using time-lapse microscopy. In all cases, cell progression from metaphase to abscission was analyzed. Arrow in (e) points to the midbody associated GFP-CGN.
77
Figure 3.10. Rac1 is required for apical lumen formation. (a) MDCK cells stably expressing GFP- CGN were embedded in 3D Matrigel and incubated in the presence of Rac1 inhibitor. Cells were then imaged using time-lapse microscopy. (b) Schematic representation showing timing of 12- hour treatment with Rac1 inhibitor. (c-e) MDCK cells were embedded in 3D Matrigel and allowed to grow for 4 days in the presence (e) or absence (d) of Rac1 inhibitor. The timing of inhibitor addition is shown in panel (b). Cells were then fixed and stained with phalloidin- Alexa594 (red) and anti-CGN (green) antibodies. Asterisks show smaller secondary lumen within the same cyst. Consecutive images without an individual letter label show the same cell. Panel (c) shows quantification of cells with single, multiple, or no lumens. Data shown are the means and standard deviations derived from three independent experiments.
78
Figure 3.11. Rac1 is required for the gp135 targeting during apical lumen formation.
79 Figure 3.11. Rac1 is required for the gp135 targeting during apical lumen formation. (a-d) MDCK cells transiently expressing GFP-Abi2 were embedded in 3D Matrigel and allowed to grow for 24 hours. Where indicated, cells were treated with 200 μM Rac1 inhibitor (b and d). Cells (a and b) were then fixed and stained with anti-acetylated tubulin (red) antibodies. White arrows (a and b) point to the midbody. (e-h) MDCK cells were embedded in 3D Matrigel and allowed to grow for 24 hours. Where indicated, cells were treated with 200 μM Rac1 inhibitor (g- h). Cells were then fixed and stained with anti-gp135 (green) and anti-CGN (red) antibodies. White arrow in panel (e) points to the midbody. White arrow in panel (g) points to misplaced CGN. (i-j) MDCK cells were embedded in 3D Matrigel and allowed to grow for 2 days. Cells were treated with 200 μM Rac1 inhibitor for the first 12 hours of the incubation. Cells were then fixed and stained with anti-gp135 (green) and anti-CGN (red) antibodies. White arrow in panel (j) points to misplaced CGN. Consecutive images without an individual letter label show the same cell.
80
Figure 3.12. Rac1 is required for the formation of a single apical lumen.
81 Figure 3.12. Rac1 is required for the formation of a single apical lumen. (a) Schematic representation showing timing of 12-hour treatment with Rac1 inhibitor. (b-e) MDCK cells were embedded in 3D Matrigel and allowed to grow for 4 days in the absence (c) or presence (d and e) of Rac1 inhibitor. The timing of inhibitor addition is shown in panel (a). Cells were then fixed and stained with phalloidin-Alexa594 (red) and anti-CGN (green) antibodies. Consecutive images without an individual letter label show the same cell. Panel (b) shows quantification of cells with single or multiple lumens. Data shown are the means and standard deviations derived from three independent experiments.
82
Figure 3.13. Electrostatic interactions mediate CGN binding to microtubule C-terminal tails (CTTs).
83 Figure 3.13. Electrostatic interactions mediate CGN binding to microtubule C-terminal tails (CTTs). (a) Microtubule binding assay showing amount of pelleted GST-CGN(1-406) and GST- CGN(1015-1203) when incubated with (+) and without (-) taxol-stabilized microtubules (tubulin). Anti-GST western blot (top gel), and Coomassie stained SDS-PAGE gel (bottom gel) are shown. (b) Diagram depicting positioning of α and β tubulin CTTs within polymerized microtubule. (c) Amino acid sequence alignment comparing CTTs of human and yeast β- tubulins. (d) Microtubule binding assay comparing amount of CGN(1-406) that pellets with wild- type tubulin and subtilisin-digested tubulin. Coomassie stained SDS-PAGE gel is shown. (e) Microtubule binding assay comparing CGN(1-406) pelleting with wild-type yeast tubulin and mutant yeast tubulin containing no β-CTT. Anti-tubulin and anti-CGN western blot is shown. (f) Amino acid sequence alignment comparing CGN basic patch in different vertebrate species. Boxes mark the sites mutated to alanine for binding analysis shown in panel (g). (g) Quantification comparing microtubule binding of wild-type CGN(1-406), CGN-M1 (R36/37A) and CGN-M2 (R40/41A). Data shown are the means and standard deviations derived from four independent binding experiments.
84
Figure 3.14. CGN binds to tubulin C-terminal tail domains. (a) Untreated or subtilisin-treated mammalian tubulin was separated on SDS/PAGE gels and subjected to Coomassie staining. Note decrease in -tubulin and -tubulin size in subtilisin treated samples due to the removal of CTTs. (b) MDCK cells were embedded in 3D Matrigel and allowed to grow for 24 hours. Cells were then fixed and stained with anti-glutamylated tubulin (red) and anti-CGN (green) antibodies. Black arrow points to midbody, white arrow to the AMIS. (c) Tubulin purified from wild-type or -tubulin (TUB2) mutant yeast. Note the decrease in - tubulin size in mutant strain due to the removal of CTT.
85
Figure 3.15. Generation of CGN knock-out in MDCK cells using CRISPR/Cas9.
86 Figure 3.15. Generation of CGN knock-out in MDCK cells using CRISPR/Cas9. (a) Schematic representation of CRISPR guide sequence designed to target canine CGN in MDCK cells. Arrows marked with “F” and “R” shows the location of primers used to amplify and sequence the region targeted with CRISPR/Cas9. Allele#1 and allele#2 show the mutation in CGN exon#2 that were introduced in both CGN genes. (b) Western blot analysis comparing CGN levels in cellular lysates isolated from a parental MDCK cells (WT) or MDCK cells co-expressing CGN CRISPR guide and Cas9 (CGN-KO). (c) Comparison of CGN levels in 4 day-old epithelial cysts formed by either parental MDCK cells (WT) or MDCK cells co-expressing CGN CRISPR guide and Cas9 (CGN-KO). (d) Parental MDCK cells (WT) or MDCK cells co-expressing CGN CRISPR guide and Cas9 (CGN-KO) were embedded in 3D Matrigel and grown for 4 days. Cells were then fixed and stained with phalloidin-Alexa596 (red) and anti--catenin antibodies (green). Arrows point to ectopic muni- lumen forming inside the cell. Quantification of the number of cysts with either single, multiple or no lumens are shown in left panel. Data shown are the means and standard deviations derived from three independent experiments.
87
Figure 3.16. Mutation of basic patch disrupts subcellular CGN targeting.
88 Figure 3.16. Mutation of basic patch disrupts subcellular CGN targeting. MDCK-CGN-KO cells were transfected with either wild-type CGN-GFP (a-b) or CGN-M1/M2-GFP mutant (R36/37/40/41A) (c-e). Cells were embedded in Matrigel, incubated for 12 hours, fixed and stained with phalloidin-Alexa596. Arrows in (a) point to actin rich lumen surrounded by CGN- GFP-containing tight junctions. Arrows in (c) point to actin accumulation at the neck of bud-like structures. Arrows in (d) point to ectopic accumulation of actin and CGN rings. Finally, arrows in (e) point to intracellular formation of mini-lumens in CGN-KO cells.
89
Figure 3.17. CGN interaction with midbody microtubules and Rac1-induced branched actin cytoskeleton is required for AMIS formation and apical lumen initiation. Model depicting pathways that lead to CGN recruitment and AMIS formation at the midbody as well as the targeting/tethering of Rab11/FIP5 apical endosomes to form a single apical lumen.
90
Figure 3.18. Uncropped scans of western blots from: (a) Figure 3.1d. (b) Figure 3.13e. (c) Figure 3.2b. (d) Figure 3.15b.
91 CHAPTER IV
CONCLUSIONS AND FUTURE DIRECTIONS
Conclusions
Epithelial cells form a mechanical and chemical barrier in every organ that is essential
for carrying out physiological function. One critical step in the development of these cells into
functional tissues is coordinated cell division and polarization to form an apical side that faces a
lumen and a basolateral side. The lumen provides a protected environment for specified organ
activity, such as water and nutrient absorption in the intestines and filtering of waste by the
kidneys. Conversely, compromised function of epithelial cells is associated with diseases of
many organs, including polycystic kidney disease and intestinal disorders such as microvilli
inclusion disease. Although much is known about the function of epithelial tissues, the cellular
mechanisms that direct lumen formation remained unknown. Here, I have focused on
identifying the initiating factors of epithelial polarization, and how endosomes carrying apical
cargo are targeted and tethered specifically to the site of lumen formation.
Vesicle transport plays an essential role in cell polarization and apical lumen formation
by both providing membrane needed for lumen expansion and a method for delivering apical
specific cargo. Previous studies in our lab have identified the interaction of Rab11 and FIP5 proteins as a key regulator of apical endocytic vesicle transport during lumen formation. FIP5
serves as the guiding protein for vesicles containing apical cargo by both initiating vesicle
budding and allowing for transport along microtubules. In epithelial cells, the site where these
vesicles fuse is known as the apical membrane initiation site (AMIS).
During the process of de novo lumenogenesis, apical lumen formation initiated by FIP5-
dependent transport of apical membrane components to the AMIS during cytokinesis. During
metaphase/anaphase FIP5-T276 phosphorylation inhibits FIP5 from binding SNX18, thus
92 preventing apical cargo protein from exiting recycling endosomes. During telophase, after the
formation of AMIS around the midbody, FIP5-T276 dephosphorylation allows the SNX18-
dependent formation of apical endocytic carriers (FIP5-endosomes). These FIP5-endosomes
contain apical cargo, including Crb3a and gp135 and move along central spindle microtubules to
fuse with the cleavage furrow PM to generate an apical lumen.
Also, Cingulin (CGN) binds directly to FIP5 and functions as a targeting/tethering factor
for Rab11/FIP5-endosomes during the early stages of lumen formation. Our data supports a
model in which both the branched actin cytoskeleton and midbody microtubules play a role in
initial establishment and maintenance of the AMIS. The C-terminal tails of microtubules bind to
a basic patch located in the globular head region of CGN, and that this microtubule binding is
likely to be required for initial CGN recruitment to the midbody. Rac1 activation at the midbody
leads to the recruitment of the WAVE/Scar complex and polymerization of branched actin
filaments. Polymerization of these branched actin filaments is necessary for the formation of the
AMIS at the midbody, and is also required for the formation and expansion of a single apical
lumen.
The requirement of both, branched actin and glutamylated microtubules serves as a
coincidence detection system that allows for the establishment of a single apical lumen.
Similarly, targeting of apical endosomes to the AMIS also depends on several factors. It was previously demonstrated that the Exocyst complex and Slps are required for the formation of a
single apical lumen (Jing et al., 2009; Mellman et al., 2008; Wu et al., 2005). CGN and branched
actin are also required for Rab11/FIP5 targeting. Thus, it is becoming clear that lumen formation
is dependent on a combination of overlapping pathways that occur at the right place and the
right time to mediate lumen formation.
93 Future Directions
Although our work has greatly expanded our knowledge of the mechanisms of epithelial cell polarization and lumen formation, there are still many factors yet to be discovered and explored. For example, although the midbody plays an important role for recruiting the apical membrane initiation site factors and marking the site of the apical lumen, it is still not known how that microtubule bundle signals the onset of the luminal cascade. Although midbody microtubules are highly modified, and that the increased negative charge brought on by polyglutamylation can interact with positively charged patches of proteins such as Cingulin, the factors involved in specifically modifying those central spindle microtubules remain unidentified.
I have begun investigating the role of tubulin tyrosine ligase-like proteins (TTLLs) in this process. Many of the TTLL family of proteins have been shown to glutamylate microtubule C- terminal tails, and TTLL12 was identified in a proteomics analysis of midbodies collected from
Hela cells. Interestingly, TTLL12 is unique among the TTLL family, and has not been shown to have enzymatic activity. It is possible, however, that TTLL12 binds to glutamylated microtubules to stabilize them and protect them from cleavage, or serve as a recruiting factor for other enzymes and factors needed to stabilize the midbody and signal polarization. I have also imaged both Hela and MDCK cells expressing flag-tagged TTLL12, and found that it localizes to the midbody during cell division. This gives even more evidence to believe that TTLL12 is playing a role in marking midbody microtubules to specify their role in polarization and lumen formation.
To deeper study this role I am imaging a GFP-TTLL12 overexpressing cell line, as well as working to create a TTLL12 knockout in MDCK cells, similar to our approach with Cingulin. For the rest of my time in the lab, I will continue to explore the role of TTLL12 at the midbody, as well as other factors that we may discover are involved in the initiation of lumen formation.
94 Another aspect of my project that could be further explored is the polymerization of actin flares around the midbody. Actin is playing many roles during division, and there are many regulators involved in coordinating its activity. For example, RhoA regulates contraction of the actomyosin ring during cytokinesis, and my work shows that Rac1 activation of branched actin polymerization plays an important role in the localization of the apical membrane initiation site.
However, more work is needed to determine the sequence of event that leads to the switching on or turning off each of these systems. In addition, these actin flares around the midbody could also be playing a direct role in pulling in vesicles that are transported to that site. Actin binding proteins such as Myosin2 have been identified as components of vesicles carrying apical cargo, but their role in targeting of these vesicles has yet to be described. In our model, it would make sense that actin flares at the midbody help carry in the vesicles as they are transported along microtubules, further pushing them for fusion at the AMIS.
While 3D tissue culture techniques are very powerful experimental approaches, morphogenesis and organogenesis in vivo is a more complicated system. There are more complex polarity and positional information cues in the in vivo system that cannot be fully replicated in cell culture. Thus, the extent to which proteins that regulate single lumen formation in 3D tissue culture systems contribute to lumen formation in vivo remains to be determined. Furthermore, while in vitro epithelial cysts start as single cell embedded in extracellular matrix, in vivo most epithelial tissues form from hundreds of non-polarized cells that simultaneously undergo polarization to generate apical lumens. How the formation and coalescence of these mini-lumens is organized and regulated in vivo remains essentially unknown. The emergence of novel genomic editing approaches as well as new animal models, such as zebrafish, will allow for the analysis of the machinery of epithelial morphogenesis in complex vertebrate organ systems. Confirmation and further investigation of these mechanisms
95 in animal models will be required to understand the normal and pathological development of
epithelial organs in vivo and to find clinical treatment strategies for related human diseases. Our
lab has also created knock-out models to explore the role of FIP5 and Cingulin in zebrafish
development, which will begin to provide some answers about the in vivo context of lumen
formation.
Our work also currently focuses on the development and mechanisms of epithelial cells,
which form lumens in organs, however, there are other cell types that express some of these
same factors that do not form lumens. For example, Cingulin is known to be expressed in
endothelial cells, where it also plays a role in cell junctions. More interestingly, Cingulin is expressed in nerve cells, which do not form tight junctions or lumens. However, communication
between neurons is dependent on rapid trafficking of signals that are essential for carrying out
function, as well as internal transport along stabilized microtubules. The role of Cingulin in
targeting specific cargo and binding directly to microtubules in epithelial cells may give some
insight into its role in the cells of the nervous system. In addition, the role of Cingulin and tight
junction proteins in deeper epithelial tissue that is not connected directly to the functional
lumenal space has yet to be described. Testing samples taken directly from patients, which
would contain multiple layers of epithelial cells as well as other tissue types, may give more
insight into the role of Cingulin in those cells. Although our work has contributed greatly to our
knowledge of de novo lumen formation, much further research is needed to continue to build
on our established model.
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