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How cells cry: The cytoskeleton as a facilitator of regulated secretion and membrane trafficking in lacrimal gland acinar cells

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How cells cry: The cytoskeleton as a facilitator of regulated secretion and membrane trafficking in lacrimal gland acinar cells

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Content HOW CELLS CRY:
THE CYTOSKELETON AS A FACILITATOR OF REGULATED
SECRETION AND MEMEBRANE TRAFFICKING IN LACRIMAL
GLAND ACINAR CELLS.
by
SILVIA REGINA DA COSTA
A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR IN PHILOSOPHY
(PHARMACEUTICAL SCIENCES)
August 2002
Copyright 2002 Silvia Regina da Costa
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UMI Number: 3094319
Copyright 2002 by
da Costa, Silvia Regina
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UNIVERSITY OF SOUTHERN CALIFORNIA
The Graduate School
University Park
LOS ANGELES, CALIFORNIA 90089^1695
This dissertation, w ritten b y
S i l v i a d i L Q y $ h ~ _____ ___
U nder th e direction o f h£..C. D issertation
C om m ittee, an d approved b y a ll its m em bers,
has been p resen ted to an d a ccep ted b y The
G raduate School, in p a rtia l fu lfillm en t o f
requirem ents fo r th e degree o f
DOCTOR OF PHILOSOPHY
ean o f Graduate Studies
D ate A ugust 6, 2002
DISSERTA TION COMMITTEE
C' t ^
V ' > Chaii
K
I f / 1 ■
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Dedication
To my family, with love and appreciation
for their help, patience and always being there
during the many times I was in need.
This work could not have been done without them.
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Ill
Acknowledgements
First and foremost I am deeply thankful to my advisor, Sarah Hamm-Alvarez,
who has been an inspiration and role model for me. Over the years we have
worked together she has become a dear friend, sharing in the good times and
being there for me during difficult moments. Graduation is a bittersweet event
as it means leaving her lab. I would also like to thank my committee members,
Profs. Austin Mircheff, Curtis Okamoto, Alicia McDonough and Vincent Lee,
for their kind help and guidance. I could not have imagined a better committee!
Thank you for all you done for me. I am particularly thankful to my friends
and lab members for their friendship and scientific help along the way. They
made coming to work a pleasure and actually, quite a lot o f fim. I will deeply
miss working with all of you. I am especially thankful to Francie Yarber for
her kindness, help, professionalism and good humor. I could never have done it
without you Francie. Last, but certainly not least, I would like to thank my
family for their support and love. They have taught me that honesty,
dedication, spirituality, hard work and joie de vivre are all important in life. I
can never thank them enough for all they have done for me. All that I have
achieved in my life, without exception, I owe to them.
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iv
TABLE OF CONTENTS
Dedication ii
Acknowledgements iii
List of Figures and Tables vii
Abstract xi
I. Chapter 1: The Lacrimal Gland, Secretion and the
Cytoskeleton 1
Introduction 1
The Cytoskeleton 2
Microtubules 3
Intermediate Filaments 8
Actin Microfilaments 8
Lacrimal Gland and Secretion 11
Sjogren's Syndrome 14
B E . Chapter 2: Materials and Methods 17
Materials and Supplies 17
Cell Culture 18
Confocal Micro scopy 18
Electron Microscopy 19
F-Actin Quantitation 20
Generation and Use of Recombinant Proteins 21
Electroporation 22
FITC-dextran Uptake 23
Isolation of Subcellular Protein Pools 24
Analysis of Protein Secretion 24
Statistical Analysis 25
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V
HI. Chapter 3: Actin Microfilaments and Clathrin-Mediated
Endocytosis 26
Introduction 26
Clathrin-Mediated Endocytosis 27
Adaptors 28
Dynamin 34
Membrane Composition and Endocytosis 37
Actin Microfilaments and Endocytosis 40
Results 46
Discussion 64
IV. Chapter 4: Actin Regulatory Proteins in Endocytosis 70
Introduction 70
Src Homology 3 (SH3) domains 71
The SH3-WASP Connection 72
The WASP VCA Domain and Arp2/3 75
Arp2/3 Activation 77
SH3 Domains and Dynamin 78
Amphiphysin and Endophilin 79
Phosphoinositides and Synaptojanin 82
Syndapin 85
Actin Binding Protein 1 (Abpl) 88
Results 91
Discussion 113
V. Chapter 5: MFs and Exocytosis in Lacrimal acinar Cells 118
Introduction 118
Actin MFs and Exocytosis 119
Small GTP-Binding Proteins 122
Rabs 123
SNAREs and VAMPs 125
Results 128
Discussion 145
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VL Chapter 6: Endocytosis in Three Acts: A Summary 150
Act I, scene I: Tension Mounts and the
Membrane Invaginates 151
Act I, scene II: The PIP2 Connection 153
Act I, scene III: Being Negative is Not
Always Bad 155
Act II, scene I: The Opening of N-WASP 157
Act III, scene I: To Pinch, and So Much More 160
Relevance of Study to Lacrimal Gland
Function and Disease 163
YU. References 167
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LIST OF FIGURES AND TABLES
v ii
Chapter 1: The Lacrimal Gland, Secretion and the Cytoskeleton 1
Figure 1.1. Microtubules Distribution 5
Figure 1.2. Cross-section of the Lacrimal Gland 12
Chapter 3: Actin Microfilaments and Clathrin-Mediated
Endocytosis 26
Figure 3.1. Clathrin-Mediated Endocytosis 29
Figure 3.2. Dynamin Domain Structure 36
Figure 3.3. Recruitment of Coat-Proteins Following
Carbachol Stimulation 47
Figure 3.4. Actin Microfilaments Distribution in Lacrimal Acini 49
Figure 3.5. Coat-Protein Recruitment to the APM Following
Carbachol 52
Figure 3.6. Disruption of the MF Network by CD Inhibits Coat
Protein Retrieval Following Secretagogue Stimulation 53
Figure 3.7. CD-Treatment Inhibits FITC-Dextran Uptake 55
Figure 3.8. Electron Microscopy of MF and Coat Protein
Distribution 57
Figure 3.9. Accumulation of Clathrin-Coated Pits at APM
Resulting from CD Treatment 58
Figure 3.10. M(3CD Inhibits the Uptake of FITC-Dextran in
Lacrimal Acini 62
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via
Figure 3.11. Apical accumulation of a-adaptin in MPCD-Treated
Acini 63
Table 3.1 Marker protein confirmation of extraction conditions 48
Chapter 4: Actin Regulatory Proteins in Endocytosis 70
Figure 4.1. WASP Domain Structure 74
Figure 4.2. Syndapin Domain Structure 86
Figure 4.3. Actin Binding Proteins Domain Structure 89
Figure 4.4. Western Blot - Syndapin and Actin Binding Protein 1 92
Figure 4.5. SdpI, SdpII and Abpl Distribution in Rabbit
Lacrimal Acini 93
Figure 4.6. Domains Structures of SdpI, SdpII and Abpl Fusion
Proteins 95
Figure 4.7. GST Pull-down Assay 96
Figure 4.8. Distribution of Synaptojanin and N-WASP in Rabbit
Lacrimal Acini 99
Figure 4.9. Introduction of Fusion Proteins into Lacrimal Acini by
Electroporation 101
Figure 4.10. Introduction of SdpI, SdpII and Abpl SH3 Domains
Results in Apical Accumulation of Coat-Proteins 102
Figure 4.11. Electron Microscopy of Control Electroporated Acini 105
Figure 4.12. Apical Coated Pit Accumulation Resulting from
Introduction of SdpI SH3 Domain 106
Figure 4.13. Introduction of SdpII SH3 Domain in Lacrimal acini 107
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ix
Figure 4.14. Effects of Abpl SH3 Domain Introduction on
Clathrin-Coated Pits and the MF Network 108
Figure 4.15. Increased F-Actin Resulting from Introduction of
SH3 Domains 111
Figure 4.16. Inhibition o f Endocytosis by M|3CD Does Not
Increase F-Actin 112
Chapter 5: MFs and Exocytosis in Lacrimal acinar Cells 118
Figure 5.1. CD Treatment Did Not Inhibit the CCH-Stimulated
Apical Release of Rab3D-Positive Secretory
Granules 130
Figure 5.2. Quantitation of Rab3D Distribution 131
Figure 5.3. Introduction of SH3 Domain of SdpI Does Not
Affect Rab3D Distribution in Resting and
Secretagogue-Stimulated Acini 134
Figure 5.4. An Intact MF Network is Not Required for the Apical
Recruitment of VAMP2 Following CCH-Stimulation 136
Figure 5.5. Electroporation of SdpI, SdpII and Abpl SH3
Domains Does Not Alter VAMP2 Distribution
in Lacrimal Acini 137
Figure 5.6. Apical Recruitment of VAMP2 in CCH-Stimulated
Electroporated Acini 139
Figure 5.7. CD-Treatment Does Nor Introduction of SH3
Domains of SdpI, SdpII and Abpl Inhibits the
Release of |3-Hexosaminidase Elicited by
Secretagogue Stimulation 140
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Figure 5 . 8 . Recovery of Apical Rab3D-Positive Vesicles
Following Stimulated Release is Delayed in
CD-Treated Acini
x
143
Chapter 6: Endocytosis in Three Acts: A Summary 150
Figure 6.1. Endophilin and Amphiphysin Signaling Pathways to
Endocytosis 154
Figure 6.2. A Role for Membrane Phospholipids in
Clathrin-Mediated Endocytosis 156
Figure 6.3. Synaptojanin - a Negative Regulator of Endocytosis 159
Figure 6.4. N-WASP: Multiple Signaling Pathways to
Endocytosis 161
Figure 6 .5. The GTPase Dynamin as a Regulator of Endocytosis;
One Protein, Many Signaling Cascades 162
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xi
ABSTRACT
Objective: The aim of this research project has been to understand the relationship
between membrane trafficking and the actin filament network in lacrimal acinar
epithelial cells, the principal secretory cell within the lacrimal gland. Particular
focus has been given to clathrin-mediated apical endocytosis as well as the
contribution of several novel effectors including syndapin I (SdpI), syndapin II
(SdpII), and actin binding protein 1 (Abpl). These proteins are thought to
participate in endocytosis by coordinating interactions between components of the
endocytic machinery (dynamin, synaptojanin-1) and actin filaments. Methods:
The content and distributions of clathrin, a-adaptin, dynamin and actin filaments
were determined by confocal fluorescence microscopy in 3 day cultures of rabbit
lacrimal acini with and without carbachol (CCH) stimulation, exposure to
cytochalasin D (CD) and in the presence of SH3 domain fusion proteins. Fusion
proteins consisting of SH3 domains of SdpI, SdpII and Abpl fused to
glutathione-S-transferase (GST) were expressed in E. coli, purified, and
introduced by electroporation. Clathrin-coated pit and vesicle formation at the
apical plasma membrane was probed in parallel under these conditions by
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x ii
electron microscopy. Results: CD treatment of lacrimal acini caused a major
enrichment of clathrin, a-adaptin and dynamin at the apical surface, which was
associated with an apical accumulation of coated pits by electron microscopy.
This enrichment was also associated with inhibition of endocytosis as measured
by flow cytometry analysis of FITC-dextran uptake. Apical accumulation of
clathrin and accessory proteins was also seen after introduction of GST-SdpI,
GST-SdpII and GST-Abpl. Fusion protein introduction also resulted in increased
F-actin. GST-pull down assays revealed that the SF1 3 domain of SdpI, SdpII and
Abpl associated with dynamin, synaptojanin and N-WASP from lacrimal acinar
lysates. Measurements of basal and CCH-stimulated protein and 1 3 -
hexosaminidase secretion revealed that the apparent inhibition of clathrin-
mediated apical endocytosis elicited by CD or the SH3 domains of SdpI, SdpII
and Abpl did not diminish the acinar exocytic response. Conclusions: Our data
suggest that SdpI, SdpII and Abpl participate in actin-mediated apical
endocytosis in lacrimal acini.
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1
Chapter 1 - The Lacrimal Gland, Secretion and the Cytoskeleton
Introduction
The aim of this research project has been to understand the relationship
between membrane trafficking events and the cytoskeleton, particularly the
microfilament network (MF) in secretory epithelial cells of the lacrimal gland.
This gland is the major exocrine gland responsible for the production and
release of tear proteins (Stern et al. 1998). Lacrimal gland acinar cells
obtained through a primary cell culture from rabbit glands were used in the
study as a model of polarized secretory epithelial cells. The data presented in
this thesis address various aspects of membrane trafficking and are a
continuation of previous work, done by me, on membrane trafficking and the
lacrimal gland (da Costa et al. 1998 and Master’s thesis). My initial research
defined the organization of the cytoskeleton in lacrimal acini and probed its
participation in stimulated secretion, giving special emphasis to the role of
microtubules (MT) as facilitators of secretion. Past studies implicated MTs in
the movement of secretory vesicles (SVs) to the apical membrane as treatment
of acini with the MT-targeted agents taxol and nocodazole inhibited
secretagogue-stimulated secretion (da Costa et al. 1998). The present study
focuses on how MFs facilitate the compensatory endocytosis and the retrieval
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2
of apical plasma membrane (APM) following stimulated secretion in lacrimal
acini.
In the present chapter, background information on the lacrimal gland, secretion
and components of the cytoskeleton will be given. Chapter 2 pertains to the
assay protocols and experimental methodology used to obtain the data herein
presented. Chapter 3 focuses on the role of the actin microfilaments (MF) as
facilitators of clathrin-mediated endocytosis at the apical plasma membrane
(APM), following secretagogue-stimulated secretion. Actin regulatory proteins
as mediators of endocytosis are the topic of Chapter 4 while Chapter 5
changes directions, so to speak, and looks at exocytosis and the role of MFs in
the outward movement of secretory vesicles. Lastly, Chapter 6 summarizes all
the data and proposes a model for MF participation in the movement of
materials to and from the apical plasma membrane (APM) of lacrimal acini.
The cytoskeleton: Cells exist in an ever-changing environment requiring
constant adaptation. The cytoskeleton, composed of a variety of protein
filaments, is a flexible and dynamic part of this adaptive process and is
composed of microtubules (MT), MFs and intermediate filaments (IFs), each
formed by a unique type o f cytoskeletal protein. These filamentous networks
are involved in a variety of functions within the cell such as cell motility,
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3
maintenance of cell structure and cytoplasmic viscosity (reviewed in Lodish et
al. 2000). They mediate membrane trafficking, endocytosis, secretion,
anchoring of cell organelles, cell attachment and interaction with the
extracellular matrix and help establish cell polarity. Maintenance of cell
polarity is fundamental to cell survival and constitutes the ability of a single
cell to establish subdomains or compartments within its structure - essential to
symmetric and asymmetric cell divisions, cell migration and vectorial transport
of materials within a cell (reviewed in Pedersen et al. 2001). The apical
cytoskeleton has been implicated in numerous studies regarding lumenal
organization and protein sorting in polarized epithelial cells (reviewed in Fath
et al., 1993; Mays et al., 1994) including acinar cells (O’Konski et al., 1990; da
Costa et al. 1998).
Microtubules: Composed of a and P tubulin dimers, MTs are ubiquitous
constituents of eukaryotic cells. These cytoskeletal structures are involved in a
variety of cell functions such as directed membrane trafficking, endocytosis,
anchoring of cell organelles and secretion (Skoufias and Scholey, 1993). In
mitotic cells, kinetochore MTs are instrumental in aligning the chromosomes at
the metaphase plate and in their separation towards the spindle poles (reviewed
in Heald and Walczak, 1999). Each polymer is formed of 13 protofilaments
which associate to form a 25 nm diameter hollow cylindrical structure, making
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the MT the largest of the cytoskeletal filaments. The a and (5 tubulin
monomers associate in a head to tail fashion, giving the microtubule an
intrinsic polarity. Consequently, MTs have distinct biochemical ends known as
the (+)- and (-)-ends, referring to the rate of growth or relative ratios of
polymerization/depolymerization at each end. In most undifferentiated non
polarized cultured cells, the MT (-)-ends are anchored at the centrosome, or
MT organizing center (MTOC), located near the nucleus. In unpolarized cells,
MT (+)-ends radiate outwards from the MTOC to the cell periphery (Schroer
and Sheetz, 1991). However, the MT array differs in specialized cells,
exhibiting a parallel organization in neuronal axons, an antiparallel
organization in dendrites and in polarized cells such as lacrimal gland acinar
cells reveling extension from the apical surface to the basal-lateral surface
(Figure 1.1.), In epithelial cells, MT (-)-ends are apical and the (+)-ends are
basolateral.
One of the key factors in the cellular adaptive response is the ability of
microtubules to polymerize and depolymerize, exhibiting a property known as
“dynamic instability” (Schulze and Kirschner, 1988). This property enables
MTs to form vast filament networks, or to break them down, according to cell
needs. MTs can undergo posttranslational modifications, known as acetylation
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5
MT Distribution
BLM
APM
Polarized Cell
Radial Array
Cell
Body
Axon
Dendrites
Figure 1.1. MT distribution in specialized and non-specialized cells.
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6
and tyrosination (Luduena, 1998; Idriss, 2000). Acetylated MTs are considered
“stable”, referring to their resistance to depolymerization after nocodazole or
cold treatment (LeDizet and Piperno, 1991). Detyrosinated MTs, formed when
the terminal tyrosine of a-tubulin is enzymatically removed, are also “stable”
MTs. Stable MTs have a much longer half-life: the in vivo half-life of MTs
without these modifications is approximately 10 min while stable MTs have
half-lives measured in hours to days (Kreis, 1987). However, other studies
(Webster et al., 1990) have shown that detyrosination alone “does not directly
confer stability on to MTs”, suggesting that other mechanisms are involved in
MT stability prior to posttranslational modifications. It is believed that
functionally, MTs enriched in acetylated or detyrosinated tubulin are
responsible for establishing or maintaining membrane organization of cellular
organelles, such as the Golgi complex (Thyberg et al., 1999). Tyrosinated, or
dynamic MTs, in which a tyrosine residue remains at the C-terminus of the a -
tubulin, are found in interphase cells as well as in the metaphase spindle.
Tyrosinated MTs comprise a distinct subset of the total MT population and,
because of their unique distribution in many cells types (Gundersen et al.,
1984), including lacrimal gland acinar cells, functional differences between
stable and dynamic MTs have been assumed.
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7
Since MTs mediate a variety of cellular functions in interphase cells, including
the formation of a network that supports membrane vesicle movements
(reviewed in Cole and Lippincott-Schwartz, 1995), previous work on exocrine
secretion has explored a possible role for MTs. A substantial precedent exists
for the hypothesis that MTs facilitate regulated secretion at a step or steps in
the process and they have been implicated in the stimulated release of secreted
proteins in lacrimal, pancreas and parotid glands (Robin et al., 1995; da Costa
et al. 1998; Malberti et al. 1998; Ueda et al. 2000). A role for MT-based motor
proteins and vesicle transport in regulated secretion has also been
demonstrated in unpolarized cells including T-cells (Burchardt et al., 1993)
and melanocytes (Rogers et al., 1997). Motor proteins are mechanochemical
proteins that utilize energy derived from nucleotide hydrolysis to drive
vesicular movement in a unidirectional manner along a filament network.
Kinesin and cytoplasmic dynein constitute families of MT-based motor
proteins while myosins utilize the actin MF array as tracks for vesicle
transport. Motor proteins have been implicated in a variety of cellular
functions, such membrane trafficking, transport and localization of mRNA and
have also been suggested to function as mediators between the MT and MF
networks, among others (extensive and excellent reviews include: Hamm-
Alvarez, 1998; DePina and Langford, 1999; Goode et al. 2000; Stebbings,
2001 and Apodaca, 2001).
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Intermediate filaments: As the name states, IFs are intermediate in size
between MTs microtubules and actin MFs, and self-assemble into cytoskeletal
filaments with a diameter of approximately 10 nm (reviewed in Coulombe et
al. 2001). They are composed of a family of filamentous proteins, namely,
cytokeratin, vimentin, desmin, glial filament proteins, and neurofilaments and
are responsible for regulating cell shape, chromosome organization and
structural stability. O f these, cytokeratins, a multigene family of over 30
different proteins, are typically expressed in epithelial cells. Extending from
the nuclear surface to the cell periphery, they are anchored at the desmosomes
and play an important structural role as mechanical scaffolds. A role for IFs
has also been suggested in cellular functions as diverse as neuron growth
(Fuchs and Cleveland, 1998), nuclear envelope assembly (Wilson et al. 2001)
and mediation of epithelial cell architecture (Herrmann and Aebi, 2000).
Defects in the IF organization or filament structure have been implicated in
loss of cellular integrity and identified with a variety of inherited diseases
(Fuchs and Cleveland, 1998).
Actin microfilament (MF): Actin is the most abundant of the cytosolic
proteins existing mostly as monomers (globular proteins) that utilize nucleotide
hydrolysis to polymerize and form a 5-9 nm diameter two-stranded helical
structures known as MFs (reviewed in Cooper and Schafer, 2000). There are
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9
six different isoforms of actin, two skeletal, two cardiac and two non-muscle.
The MF network (F-actin) is in dynamic equilibrium with its protein soluble
pool (G-actin), changing the rates of polymerization/depolymerization
according to cell needs. Similar to their MT counterparts, MFs ends are
structurally distinct with different growth rates: the slow growing minus or
“pointed” end (filament depolymerization end) and the fast growing plus or
“barbed” end (filament polymerizing end). The directional growth at the
barbed end gives rise to what is known as actin “treadmilling”, a function also
associated with movement at the leading edge of cells (Carlier and Pantaloni,
1997). Actin MFs form cellular stress fibers, cell microspikes and are the basic
structural unit of microvilli. They are responsible for a variety of functions
including maintenance of cytoplasmic viscosity, maintenance of cell structure
and they also provide the mechanical basis for cell movement. Reorganization
of the cortical actin cytoskeleton in response to extracellular stimuli is a key
event in cell migration. MFs anchor cells to the substrata, through focal
adhesions. Dynamic changes in the MF array provides the structural stability
needed for membrane extension in phagocytosis (reviewed in Castellano et al.
2001). During cell division, actin MFs are also involved in cytokinesis, or the
“pinching o ff’ of the two daughter cells. Of special interest to this project, MFs
have been also implicated in endocytosis (see Chapter 3) and exocytosis
(reviewed in Schafer, 2002; Schafer, 2002; Chapter 5). As well, MFs serve as
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10
“roads” for a family of motor proteins known as unconventional myosins,
which, like the MT-based motor proteins kinesin and dynein, utilize the energy
derived from ATP hydrolysis to move their “cargo” along the filament. Actin-
based motor proteins have been implicated in membrane trafficking events as
well targeting of transported membranes to correct end-point organelles via
interaction with Rabs (see Chapter 5, reviewed in Hammer and Wu, 2002).
Filament growth is regulated by capping proteins, CapZ 15 and tropomodulin
(Tmod), located at the pointed and barbed ends, respectively (Fowler et al.
1993; Weber et al. 1994). MF dynamics is also regulated by a highly integrated
and complex signaling cascades responsible for transmitting extracellular
stimuli to the actin filaments (Hall, 1998). The signaling cascade is governed
by Rho subfamily of small GTPases (see Chapter 5 for a review on GTPases).
Over 20 Rho family members have been identified (Ridley, 2001) and are
known to play a critical role in the formation and organization of cortical actin
networks (Lanzetti et al. 2001). Rho family members co-localize with actin-
rich structures and include, among others: Cdc42, implicated in filopodia
formation; Racl, associated with lamellipodia and cell membrane ruffles and
Rho A, implicated in stress fiber regulation (Hall, 1998). A role for Rho family
members has also been demonstrated in membrane trafficking: constitutively
active Racl inhibites endocytosis in polarized (Jou et al. 2000) and non
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11
polarized cells (Lamaze et al. 1996) and Cdc42 has been implicated in
basolateral secretion and endocytic membrane trafficking (Kroschewski et al.
1999). The exact mechanism of actin of Rho family members on endocytosis is
not completely understood - indirect regulation of endocytosis could result
from their role in actin remodeling and polymerization. However, a more direct
association has been suggested in recent research that demonstrated that Racl
interacts directly with synaptojanin (Malecz et al. 2000; see Chapter 4).
The Lacrimal Gland and Secretion: Located on the outer and upper
conjunctival sac, the lacrimal gland is composed of various lobes and, in
humans, is about the size and shape of an almond. It is the major exocrine
gland responsible for tear fluid production, releasing water, electrolytes and
proteins to bathe the cornea. Other glands which also aid in the lubrication of
the eye are the tarsal or meibomian glands, located at the margin if the eyelid,
the glands of Wolfring, in the subconjuctival tissue above the meibomian
glands and the glands of Krause, proximal to the lid margin. Secretion occurs
from each lobe onto ducts that empty their fluid contents onto the surfaces of
the superior conjunctival fornix (reviewed in Walcott, 1994). Lacrimal gland
acini are formed by polarized epithelial cells that are joined at the apical
membrane forming the lumen, through which stimulated secretion occurs
(Figure 1.2.), Lacrimal gland acinar secretion is primarily controlled by the
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12
Basal-Lateral
Membrane ,
Apical j
Miembiane,
Lumen
Figure 1.2. Horizontal cross-section of the lacrimal gland
acinus. Cultured acinar cells, isolated from rabbit, regain their
polarized, differentiated state displaying distinct apical and
basolateral membrane domains.
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13
autonomic nervous system (Ruskell, 1975). Steroid hormones also regulate the
functional status of the lacrimal gland, that is to say, its ability to respond to
secretory stimulation. Stimulation of secretion is obtained through various
signal transduction pathways, including the Ca2 + /DAG/cGMP dependent
pathway as well as the cAMP dependent pathway (Dartt, 1994). The acinar
cells of the lacrimal gland secrete an isotonic NaCl and protein-rich fluid. The
parasympathetic innervation responsible for stimulating release of this fluid in
vivo can be mimicked in vitro by a number of agents, including carbachol
(Meneray et al., 1994: Gierow et al., 1996). Exposure to the muscarinic agonist
carbachol (CCH) is known to stimulate both Ca2 + /calmodulin-protein kinase
and protein kinase C (Zoukhri et al., 1994: Hodges et al. 1994: Zoukhri et al.,
1997) and to activate phospholipase D (Zoukhri et al., 1995). The
phosphorylation and consequent activation of specific protein targets of these
second messenger pathways are believed to play a direct role in protein
secretion (reviewed in Dartt, 1994). However, much still needs to be elucidated
about the identity of the target proteins which respond to cholinergic
stimulation and changes in intracellular phosphorylation by facilitating the
release of secretory fluid and proteins.
We utilized the carbachol-stimulated release of (3 -hexosaminidase as our model
for stimulated protein secretion. Previous work with rabbits has shown that
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14
total amounts of P-hexosaminidase in secreted lacrimal gland fluid increase in
direct proportion to the secreted fluid volume when rabbits are stimulated with
pilocarpine (Gierow et al., 1995). Moreover, reconstituted lacimal acini release
P-hexosaminidase into the culture medium in response to cholinergic
stimulation (Gierow et al., 1995; da Costa et al. 1998). As well, our previous
work with analytical subcellular fractionation methods has shown that
membrane-associated kinesin codistributed with an apparent post-Golgi
secretory compartment containing P-hexosaminidase in resting acini (Hamm-
Alvarez et al., 1997). Exposure of acini to carbachol resulted in corresponding
parallel redistributions of membrane-associated kinesin and P-hexosaminidase
that appeared to reflect a return of material to the Golgi complex from the
secretory pathway via the apical plasma membrane, as reported previously by
Herzog and Farquahar (1976). Although we cannot eliminate the possibility
that some P-hexosaminidase may exit the cell through basal-lateral pathways
during stimulation, these data support an exodus through the apical secretory
pathway.
Sjogren’s syndrome and dry eye disease. Sjogren’s syndrome (SS) is an
autoimmune disorder characterized by lymphocytic infiltration and progressive
destruction of the lacrimal and salivary glands, leading to autoantibody
production (Fox et al. 1984; Fox, 1992). Regrettably, identification and
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15
treatment of SS patients is difficult given the fact that there is no
internationally established set of clinical and immunopathological markers or
assessment tools that adequately represent all aspects of the disease (Asmussen
and Bowman, 2001). Nevertheless, generalized criteria for the assessment of
disease status (Asmussen et al. 1997) and damage (Sutcliffe et al. 1998) have
been developed: SS is characterized by a variety of systemic and oral
complications, including Sicca complex symptoms (dry eye, mouth skin, etc),
parotid and salivary gland enlargement, with some patients displaying mild
symptoms of rheumatoid arthritis (morning stiffness, but without joint
deformity). Additionally, there have been reports of increased surgical
complications during anesthesia and the post-operative period in SS patients
(Fox, 1992). The most common complaint is ocular foreign body sensation
usually described as an uncomfortable “sandy” feeling in the eye, eye fatigue,
or a burning itchy feeling in the eye, among others (Fox, 1992). SS is classified
as primary or secondary; in the latter case it is commonly found in patients
with rheumatoid arthritis or other connective order diseases (Moutsopoulos et
al. 1980; Asmussen and Bowman, 2001). Many of the available treatments are
still symptomatic, such as artificial tears. Agents that modulate the immune
system have also been used, such as topical ocular use of corticosteroids or
cyclosporin (Gunduz and Ozdemir, 1994; Stern et al. 1998). Other
immunosuppressant treatments are in clinical trials, such as the use of
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16
interferon-alpha (Ship et al. 1999). Additionally, treatment with muscarinic
agonists such as pilocarpine and cevimeline have already been approved
treatment of some of the symptoms experiences by SS patients (Fox, 2001).
These agents increase secretory function by stimulating M l and M3 receptors
present on salivary and lacrimal glands
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17
Chapter 2 - Materials and Methods
Materials and Supplies: Carbachol, rhodamine-phalloidin, methyl-|3-
cyclodextrin, cytochalasin D, FITC-conjugated goat anti-mouse and goat anti
rabbit secondary antibodies and mouse monoclonals antibody to a-adaptin,
were obtained from Sigma Chemical Co (St. Louis, MO). The mouse
monoclonal antibody to dynamin was obtained from Transduction Laboratories
(Lexington, KY). The hybridoma producing the X-22 antibody to clathrin
heavy chain was obtained from the Developmental Studies Hybridoma Bank
and antibody was affinity-purified using Gamma-bind Sepharose (Pharmacia).
Rabbit polyclonal antibodies to syndapins I, syndapin II and N-WASP were
generated and purified as previously described (Qualmann et al., 1999;
Qualmann and Kelly, 2000). FITC-dextran and mounting medium were
obtained from Molecular Probes (Eugene, OR). Goat anti-rat and anti-mouse
horseradish peroxidase-conjugated secondary antibodies were obtained from
Amersham (Arlington Heights, IL). Matrigel was obtained from Collaborative
Biochemicals (Bedford, MA). The sheep polyclonal antibody to the polymeric
immunoglobulin A receptor (plgAR) was generated against rabbit secretory
component (SC) from bile. The mouse monoclonal antibody to synaptojanin I
was kindly provided by Dr. Pietro DeCamilli (Yale University). All other
chemicals were reagent grade and obtained from standard suppliers.
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18
Cell Culture: Female New Zealand white rabbits weighing between 1.8 and
2.2 kg were obtained from Irish Farms (Norco, CA). Lacrimal gland acinar
cells were isolated as described previously (Gierow et al., 1996: Hamm-
Alvarez et al., 1997) and cultured for 3 days. All animal studies were done in
accordance with the Guiding Principles for Use of Animals in Research.
Experimental treatments included cytochalasin D (CD, 5 pM, 60 minutes,
37°C) or methyl-P-cyclodextrin (MpCD, 1 mM, 30 min) with and without
CCH (100 pM, 5-15 min).
Confocal Microscopy: Rabbit lacrimal acini were cultured for three days on
Matrigel-coated coverslips then processed for confocal fluorescence
microscopy. For detection of a-adaptin, dynamin, syndapin II, synaptojanin-I
and N-WASP, acini rinsed in DPBS were fixed and permeabilized with ice
cold ethanol (-20°C, 5 min) (Mendell and Whitaker, 1978; Yang, 1997) prior to
blocking with 1% bovine serum in DPBS. For detection of clathrin, acini were
exposed to 0.1 M PIPES, pH 7.0, 5 mM M gS04, 10 mM EGTA, 2 mM DTT,
4% polyethylene glycol supplemented with protease inhibitor cocktail
(daCosta et al, 1998) and containing 0.02% saponin for 12 min at 37°C before
fixation in 4% paraformaldehyde, quenching in 50 mM NH4CI and blocking
with 1% bovine serum in DPBS. All samples were then exposed to
appropriate primary or secondary FITC-conjugated antibodies or affinity label.
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19
Samples were imaged using a Nikon PCM Confocal System equipped with
Argon ion and green HeNe lasers attached to a Nikon TE300 Quantum
inverted microscope and Simple PCI software.
Electron Microscopy: Samples were cultured for 3 days on Matrigel-coated
Biocoat Transwell plates. Acini were rinsed with fresh PCM then treated with
CD (5 mM, 60min), CCH (100 mM, 15 min), or CD+CCH. Acini were then
rinsed with 0.1M cacodylate (CaCO) buffer , and fixed in half strength
Karnovsky's fixative (2 hours, RT). The acini were washed in CaCO buffer (3
X 10 minute washes). The cells were post-fixed with 1% 0 s0 4 in 0.1M
cacodylate buffer (2 Hrs, RT). The acini were washed in CaCO buffer (3 X 10
minute washes). Cells were then incubated with 1% tannic acid in 0.1M
cacodylate buffer (1 hour, RT), and dehydrated in a graded ethanol series (25,
50, 75, & 95% respectively). The cells were en bloc stained with 2% unranyl
acetate (UA) in 50% EtOH during the first change of 50% EtOH. Acini were
embedded in LR White, and cured for two days at 60°C. Subsequent thin
sections were stained with 2% aqueous UA and Sato's modified lead stain.
Sections were viewed on a JEOL 1200 EX electron microscope operating at
80KV. Images were compiled using Adobe Photoshop 5.0 for Windows 98
(Adobe Systems Inc, Mountain View CA).
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20
F-actin quantitation: For quantitation of F-actin labeling intensity, acini of
comparable size and thickness and labeled with rhodamine-phalloidin were
selected for z-scanning. All images were acquired at equal gain and contrast
intensities as well as pinhole thickness and zoom (2x zoom magnification,
100% laser strength). Initial gain and contrast levels were established using a
control and treated or electroporated sample to ensure that fluorescence levels
were not saturated. Selected acini were scanned to determine the z-scan range.
Nine to ten z-plane sections, from the bottom to the top of the acinus, were
acquired (thickness of each section plane, 0.05 pm). Image fluorescence
intensity was subsequently analyzed utilizing Metamorph data analysis
software as follows: of the 10 acquired z-plane sections, only the 7 middle
ones are utilized with the first and last sections discarded to avoid imaging the
very small surface areas of the acinus (top) and/or stress fibers and non
specific staining of slide/Matrigel surface (bottom). Quantitation of the
fluorescence intensity of each of the 7 sections is done individually. As the size
of the acinus varies in each section, the region of interest (ROI) for data
acquisition determined by drawing a box around the acinus. The fluorescence
intensity (F.I.) of the ROI is then normalized to the box size resulting in a
value to each plane given in units of F.I./pixel2. The total intensity of the
acinus equals '^ F .I .lp ix e l2.
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21
Generation and use o f recombinant proteins: Plasmids (pGEX-5X) encoding
the SH3 domains of syndapins I (SdpI-SH3, residues 376-441) and II (SdpII-
SH3, residues 419-488) as fusion proteins with glutathione-S-transferase
(GST) were generated as previously described (Qualmann et al., 1999;
Qualmann and Kelly, 2000). A third construct, a point mutation in the SH3
domain of syndapin I (SdpI-SH3 (mut), P434L) was also utilized; this mutation
has previously been shown to abolish the binding of Sdpl-SH3 to dynamin
(Qualmann et al., 1999). GST for additional control experiments was
expressed from the plasmid pGEX-2T. GST and GST-fusion proteins were
expressed in Escherichia coli BL21 cells and purified using glutathione
agarose beads (Sigma-Aldrich) as described previously (Qualmann et al., 1999;
Qualmann and Kelly, 2000).
For isolation of lacrimal acinar proteins with binding affinity for syndapin SH3
domains, a 50% slurry of glutathione-uniflow resin (Clontech) was incubated
with 75 p,g of either GST or GST-fusion protein (or beads alone, as control) in
500 pi of DPBS for 5 hrs at 4°C (with end-over-end rotation) then rinsed
extensively in DPBS and binding buffer (10 mM HEPES, pH 7.4, 150 mM
NaCl, 1 mM EGTA, 0.1 mM MgCb, 1% TX-100 plus protease inhibitor
cocktail). Rabbit lacrimal acini (3.0 x 107 cells/sample) were resuspended in
binding buffer, lysed by freeze/thaw and processed to obtain a high speed
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22
supernatant by first low speed (14,000 rpm, 10 min, 4°C) and then high speed
(100,000 x g, 15 min, 4°C) centrifugation. This high speed supernatant was
incubated with beads or beads plus GST or GST fusion proteins overnight with
end-over end-rotation at 4°C, and subsequently rinsed extensively with binding
buffer. Proteins were eluted from beads into sample buffer, separated by SDS-
PAGE, transferred to nitrocellulose and proteins of interest analyzed by
Western Blotting.
Electroporation: Fusion proteins were introduced into rabbit lacrimal acini by
electroporation. Briefly, acini cultured for 3 days were gently pelleted and
resuspended in fresh medium at a concentration of (3.0-4.5 xlO7 cells/0.5 ml
medium) and placed into a cuvette for electroporation (BioRad Gene Pulser
Cuvette, 0.4 cm electrode gap, 50 Volts, 960 pF capacitance). Pulse lengths
ranged from 16-20 msec. Electroporation efficiency (>85%-90% in each assay)
was confirmed in each experiment by electroporation with (3-galactosidase and
subsequent phase microscopy to detect colorimetric production with X-Gal as
substrate. The concentration of GST and GST fusion proteins during
electroporation was 10 pM (174 pg/500 pi of cell suspension). After
electroporation cells were kept on ice for 5-10 min to recover and then were
seeded onto Matrigel-coated coverslips, Transwell dishes or 24-well plates and
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23
incubated for 3 hrs at 37°C prior to fluorescence microscopy, electron
microscopy and secretion assays, respectively.
FITC-Dextran Uptake Assay: Lacrimal acini cultured for 3 days were isolated
by gentle scraping and centrifugation, then resuspended in fresh medium
(8xl06 cells/ml). Acini were treated with and without cytochalasin D (5 pM,
60 min) or methyl-P-cyclodextrin (ImM, 30 min) treatments prior to addition
of FITC-dextran (1.25 mg/ml) and stimulation with and without CCH (15 min,
37°C). Uptake was stopped by addition of excess cold DPBS. Acini were
centrifuged (low speed tissue culture centrifuge) and the pellet was
resuspended in 1 ml of a 50% dispase solution in DPBS and incubated end-
over-end at 4°C for 60 min. Dispase enzymatic digestion was quenched by
addition of 500 pi of 10 mM EDTA. Acini were pelleted and resuspended
briefly in 1 ml of Ham’s supplemented with collagenase (250 U/ml),
hyaluronidase (350 U/ml) and DNAse (40,000 U/ml) prior to centrifigation and
resuspension in DPBS on ice.
Isolation o f subcellular protein pools: Methods for detergent-based extraction
were based on those described by Hollenbeck (1989) utilizing sequential
release of proteins into buffers containing 0.02% saponin, 1% Tx-100 and 1%
SDS. The distributions of proteins of interest (a-adaptin, clathrin, dynamin) as
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24
well as standard proteins (tubulin, actin, plgAR) were assessed by Western
blotting with appropriate primary and horseradish peroxidase conjugated
secondary antibodies. Detection was with the ECL detection system (Pierce)
and bands were quantified by densitometry. Lactate dehydrogenase (LDH)
activity in these samples was assayed using a Lactate Dehydrogenase Kit
(Sigma).
Analysis o f protein secretion: Procedures for secretion assays were based on
those reported previously (da Costa et al., 1998; Zhang et al., 2000). Control
and treated (electroporated, CD or M(3CD) acini seeded in Matrigel-coated 24-
well plates were incubated in fresh medium in the presence and absence of
treatments as appropriate before removal of a small aliquot for measurement of
protein and P-hexosaminidase activity. After treatment with or without CCH
(100 pM, 30 min), a second aliquot of medium was removed for measurement
of protein content and P-hexosaminidase activity. Attached cells were
dissolved with 5 N NaOH for subsequent measurement of protein content per
well. In each assay, protein and p-hexosaminidase release into culture medium
were calculated from 3-6 replicate wells/treatment and normalized to total
cellular protein. The difference between the post-incubation and pre
incubation values from unstimulated acini represents basal release. The
difference between the post-incubation and pre-incubation values from
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25
stimulated acini represents total release (basal plus stimulated). P-
hexosaminidase activity was determined using methyumbelliferyl-a-D-
glucosaminide as substrate (Barrett and Heath, 1977) while protein was
determined with the Micro-BCA protein assay (Pierce) with bovine serum
albumin as standard protein.
Statistical Analysis: Standard t-tests assuming equal variances were utilized
for analysis of Western Blot densitometry. Paired student's t-tests were
utilized for comparison of P-hexosaminidase and protein secretion data values
as well as Metamorph FITC-intensity quantitation values. A value of p<0.05
was considered statistically significant.
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26
Chapter 3 - Actin Microfilaments and Clathrin-Mediated Endocytosis
Introduction
Endocytosis: Endocytosis is a mechanism by which cells internalize cell
surface components, uptake nutrients and fluids and is also important in the
regulation of cellular function, such as cell surface receptor downregulation
and/or propagation of signaling pathways (Li et al. 2001). Endocytosed cell
surface components are transported to early endosomes, then routed according
to cell needs. Receptors involved in “housekeeping” functions can be reutilized
and recycled back to the plasma membrane. Others, such as down-regulated
hormone receptors, are targeted to the late endosome and lysosome
degradation pathway and still others are targeted for transcytosis and move
from the apical to the basal-lateral membranes (or vice-versa). Some endocytic
processes are non-specific, such as pinocytosis, by which cells continuously
take up fluids, as well as materials dissolved in them. Phagocytosis, another
non-specific endocytic mechanism, utilizes cellular extensions that surround
and subsequently envelop and internalize large particles. Endocytosis,
however, can also be highly regulated and specific, such as in the case of
receptor-mediated endocytosis (RME). RME is initiated with the binding of a
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27
ligand (an extracellular molecule) to a receptor on the plasma membrane
surface. The ligand-receptor binding triggers a signaling cascade that results in
the selective internalization of the membrane region containing the ligand-
receptor complex. Common examples of RME include the uptake of
transferrin, insulin and many other hormones. The question of whether
receptor signaling pathways are switched-off after endocytosis or continue
active until the end-point of the endocytic journey is still debated. Recent
studies have suggested that signaling cascades are extended throughout the
endocytic pathway and that some signaling events, such as activation of
extracellular signal-related kinases, seem to require endocytosis, such as the
Notch receptor association with transmembrane protein Delta (Parks et al.
2000). Protein components associated with signal transduction cascades are
thought to remain associated with endocytic vesicles following their release
from the plasma membrane (McPherson et al. 2001).
Clathrin-mediated endocytosis: One of the most common and best-
characterized mechanisms of RME is clathrin-mediated endocytosis (reviewed
in Brodsky et al. 2001). Coated vesicles were discovered almost 25 years ago
by Roth & Porter (1964) in the course of studying uptake of yolk protein by
insect oocytes. Observations of thin sections by electron microscopy studies
revealed membrane invaginations (pits) and pinched-off vesicles with
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28
characteristic electron dense coats, subsequently shown to be composed of
clathrin. A role for clathrin has been established as a facilitator in various steps
of membrane trafficking, including transport from the plasma membrane to
endosomes as well as from the Golgi to endosomes and Golgi to lysosomes
(reviewed in Harter and Reinhard, 2000). Clathrin molecules, or triskelions,
from the Greek, three-legged, come together to form a basket-like structure.
Each leg is formed by one heavy chain (= 180 kD) and one light chain (s 35-
40 kD). Two types of light chains have been identified, however their
functional differences are still unknown. Thirty-six triskelions are needed to
form a clathrin-coated vesicle (CCV). In some cells, such as hepatocytes or
fibroblasts, clathrin-coated pits (CCP) can make up to approximately 2 percent
of the cell surface (Lodish et al. 2000). Clathrin-mediated endocytosis is a
multi-step, complex and highly coordinated process by which adaptor proteins
recruit clathrin to the plasma membrane (Hirst et al. 1998) resulting in the
subsequent membrane invagination and the formation of a CCP (Figure 3.1.,
Step 1-2). It is still unclear whether the initial recruitment of adaptor proteins
to the plasma membrane requires association of adaptors to other proteins
(Takei et al. 1998).
Adaptors: Adaptor proteins form a family of heterotrimeric complexes (=
300kD); they are found in nucleated cells from yeast to humans and are
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29
___ ___ _ „ . . . „ „ „ — ™ ......,
/ ► • » > *
I y 2
► < * Plasma M embrane
3 m M
■ < ' > %
% V. - N 4
\ * ~
\ / 1 \
V' * % . .
f |
A I ' % ! S H
!
/ % 1 \ m
/ \ 1 ~ A M
1 : ' % | * *
1
■ k % .
Clathrin
a-adaptin
Dynamm
Accessory Proteins
Figure 3.1. Clathrin-mediated endocytosis. Recruitment
of clathrin to the APM by a-adaptin (Step 1) results in the
formation of a coated pit (Step 2) which is subsequently
pinched off by the GTPase dynamin (Step 3) to form a
coated vesicle (Step 4).
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30
considered assembly particles, binding to each clathrin heavy chain in a
triskelion and mediating the formation of clathrin cages. Adaptors are also
responsible for determining which proteins are included or excluded from the
forming clathrin vesicle, as they also bind to the cytosolic face of membrane
proteins (reviewed in Kirchhausen, 1999). Initially, two adaptor isoforms were
identified - API and AP2 - each associated with a different trafficking step.
API, originally identified in the mid 1980’s, facilitates traffic from the trans-
Golgi network (TGN) to the endosome, while AP2 complexes were associated
with cargo originating at the plasma membrane as well as endosomes (Ahle
and Ungewickell, 1986; Robinson and Pearse, 1986; Ahle et al. 1988). Various
novel adaptors have recently been identified: p-arrestin (Goodman et al 1996);
AP3, mostly associated with endosomal compartments (Dell’Angelica et al.
1997; Simpson et al. 1997); and AP4, associated with TGN-membranes
(Dell’Angelica et al. 1999). The exact function of these novel adaptors is not
completely understood. However, unlike API and AP2, AP3 and AP4 have not
been shown to interact directly with clathrin in vivo.
Adaptor complexes are composed of four different types of adaptin chains
(Pearse and Robinson, 1990). Each complex contains two large chains, one
medium chain and one small chain (100 kD, 50 kD and 20 kD respectively)
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31
and their primary structures are highly conserved (50-80% identity). AP2 is the
most abundant of adaptor proteins and given the high sequence similarity
between the various isoforms, AP2 has been the most studied of the adaptors
(Kirchhausen, 1999).
P-arrestin, a monomeric adaptor, has been observed to mediate the removal of
P2-adrenergic receptors from the plasma membrane. Data suggest that P-
arrestin is recruited to the cytosolic face of activated P2-adrenergic receptors
where it serves to link the receptor to the N-terminal domain of clathrin
(Goodman et al. 1997). Although P-arrestins can bind to clathrin, they are not
believed to drive the formation of the clathrin coat (Goodman et al. 1996).
Downregulation of P2-adrenergic receptors is inhibited in cells transfected
with a dominant-negative form of P-arrestin, which still binds clathrin but not
the receptor. However endocytosis of transferrin is not prevented in these same
cells (Kirchhausen, 1999). Consequently, arrestins, although able to link cargo
proteins to clathrin, are thought to play only an indirect role in the regulation or
possible nucleation of the coated pit. Although it is widely accepted that API
and AP2 are essential for clathrin-coat formation, there have been challenges
to this dogma. Rad et al (1995) found that selective knockout of the yeast
A PI2 gene (which is responsible for encoding the P chain of API and AP2)
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32
still showed normal phenotypes in cells expressing normal clathrin. Also,
recent studies showed that simultaneous disruption of all AP subunits did not
generate a clathrin-negative phenotype and had no effects on the formation of
clathrin-coated vesicle in mutant yeast (Huang et al. 1999).
Additionally, studies have suggested that other proteins such as A PI80 (also
named NP185 and F I-20) (Ahle and Ungewickell 1986; Morris et al. 1993),
epsin 2 and amphyphysin (Ramjaun and McPherson, 1998), syndapins
(Qualmann et al. 1999; Qualmann and Kelly, 2000), actin binding protein 1
(Kessels et al. 2000), among many others, may also facilitate clathrin-mediated
endocytosis, although the exact mechanisms are still not well understood.
These proteins associate either with clathrin, with adaptors or with actin
microfilaments (MFs) and are thought to modulate clathrin cage
assembly/disassembly, vesicle scission and or recycling. Interaction between
the various clathrin-associated and regulatory proteins is mediated by a variety
of protein structural modules; protein-lipid recognition modules (i.e.:
pleckstrin homology (PH) domain) and/or protein-protein recognition domains
(i.e.: Src homology 3 (SH3) domains; coiled-coil domains, proline-rich
domains (PRD); or Epsl5 homology (EH) domains). These protein-protein or
protein-lipid interactions are thought to be regulated by
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33
phosphorylation/dephosphorylation reactions (reviewed in Slepnev and de
Camilli, 2000; also see Chapter 4, Introduction).
Membrane proteins destined to be internalized via clathrin-coated vesicles
(CCVs) typically have one or more sorting signal sequences in their
cytoplasmic domain(s) that direct the protein to the CCP. A well-established
function of adaptors is to recognize these sorting signals (Pearse and Robinson
1990). Typically, sorting signals are composed of short peptide sequences (4-6
amino acids), such as the NPXY motif of the LDL receptor, the first identified
sorting signal, where N, P and Y are asparagine, proline and tyrosine,
respectively, and X can be any amino acid, Anderson et al. 1977). Other
sorting signals include the Yppo signal found in the cytosolic tail of the
transferrin receptor (Collawn et al. 1990), which is also widely found in a
number of membrane proteins and is used not only as an endocytioc marker
but in trafficking within the endosomal and secretory pathway (Kirchhausen et
al. 1997). Once the sorting signal is recognized, the receptor-bound ligand is
introduced into the CCP which is subsequently released from the plasma
membrane and becomes a CCV (sizes range from 50-100nm, Figure 3.1., Step
4). This last step, CCP release from the plasma membrane and formation of
CCV, has been shown to be facilitated by the GTPase dynamin, a “pinchase”,
which wraps itself around the neck of the coated-pit and utilizes GTP
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34
hydrolysis to constrict and pinch off the neck of the coated pit, resulting in the
formation of a CCV, which is then internalized (Bottomley et al. 1999).
Dynamin: Dynamin was first isolated from bovine brain in the late 1980’s, in
the search for a novel microtubule-associated motor protein (Paschal et al.
1987). Sequencing of dynamin led to its classification as a novel small
GTPase, given its homology to other larger GTPases (Obar et al. 1990). As
dynamin was identified originally in brain tissue, it was thought to be a
neuron-specific protein. Subsequent studies, however, have identified various
dynamin genes: dynamin 1, expressed in neurons (Orci et al. 1986), dynamin
2, ubiquitous (Serafini et al. 1991); and dynamin 3, expressed in the testis,
brain and lung (Donaldson et al. 1992). A role for dynamin in endocytosis was
first determined when its cDNA was found to have an 81% sequence similarity
with the D. melanogaster shibire gene (van der Bliek and Meyerowitz, 1991;
Chen et al. 1991). Shibire flies are paralytic mutants - paralysis is the result of
depletion of vesicles containing neurotransmitter at the nerve terminal. The
paralysis, which is rapid, reversible and temperature-sensitive at non-
permissive temperatures (Grigliatti et al. 1973), has been shown to be the result
of a block in endocytosis (Kosaka and Ikeda, 1983a and 1983b) with a
concomitant accumulation of clathrin-coated pits found at the plasma
membrane (Kosaka and Ikeda, 1983a). These observations led to the idea that
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35
there was a role for dynamin in the formation of clathrin-coated vesicles. Later
studies confirmed this hypothesis by showing that overexpression of a
dynamin mutant, unable to bind the GTP, resulted in inhibition of endocytosis
(van der Bliek et al. 1993; Herskovits et al. 1993; Damke et al. 1994) and
produced shibire phenotypes (Herskovits et al. 1993). Characterization of
dynamin’s functional role as a “pinchase” also resulted from data suggesting
that dynamin could assemble into rings (Hinshaw and Schmid 1995) and form
helical tubes on lipid bilayers (Sweitzer & Hinshaw 1998, Takei et al 1998)
which resembled structures seen at the coated pits of the shibires mutant
(Kosaka and Ikeda, 1983).
Much research has been done over the years to elucidate the functional role of
dynamin in endocytosis, although the precise mechanism of the pinching-off
reaction and the formation of endocytic vesicles is still unclear. Five dynamin
protein domains have been identified (Figure 3.2.), The N-terminal GTPase
domain is the most highly conserved (reviewed in Warnock and Schmid,
1996). Two dynamin domains function as positive regulators of the GTPase
activity - the C-terminal proline rich domain (PRD) and, as its name indicates,
the GTPase effector domain (GED), a coiled-coil domain also suggested to
play a role in dynamin assembly (Muhlberg et al. 1997). GTPase activity is
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36
Dvnamin
GTPast* M IDD LE
Bruno et.al, Nattire 410 2J1-1J5. M > 01
FH: pleclwtrin homotogy domain:
GEp: GTPasw cfift cfor ifontain;
FRDr pioiSttt/ai gttiliH-ricU domain.;
F M
1
Pt(lIns|4„5)P;
containing lipids
GEO F R 1 >
1
S i B
Dynamin oiigomerUatiem
Adlyatton of GTPase activity
Figure 3.2. Dynamin domain structure: Five dynamin domains have
been characterized: anN-terminal GTP hydrolysis domain (residues 1-
299), a middle domain (residues 300-520); a pleckstrin homology (PH)
domain (residues 521-622); a GTPase effector domain (GED) (residues
623-745); and a C-terminal proline-rich domain (PRD), which binds to
SH3-domain containing proteins (residues 746-864).
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37
downregulated by the pleckstrin homology (PH) domain (Muhlberg et al
1997). The PH domain can bind to phosphoinositides and has also been shown
to be vital for membrane localization of dynamin and RME (Salim et al. 1966).
Finally, the C-terminal PRD domain contains several SH3-binding domains
(with a PXXP motif, in which X is any amino acid, reviewed in McPherson,
1999) and is essential for dynamin function in protein-protein interactions.
Many SH3-domian containing proteins, such as syndapin I, syndapin II, actin
binding protein 1 and endophilin, among others, have been shown to facilitate
clathrin-mediated endocytosis via interactions with dynamin’s PRD (see
Chapter 4). Although much of the research to date has focused on dynamin’s
role in clathrin-mediated endocytosis, it has also been implicated in clathrin-
independent endocytosis (Artalejo et al. 1995) as well as in phagocytosis in
macrophages (Gold et al. 1999) and in caveolae-dependent endocytosis
(Henley et al. 1998).
Membrane composition and endocytosis: Although a wealth of research on
endocytosis has been done to elucidate the role clathrin and the many clathrin-
associated proteins and signaling pathways vital to clathrin-mediated
endocytosis, much has yet to be clarified, including a more detailed
understanding of how cargo is directed to the appropriate sorting vesicle and
how membrane compartments are targeted by adaptors. Additionally, a variety
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38
of clathrin-independent endocytic mechanisms have also been well described
(Sandvig and van Deurs, 1999). One such mechanism utilizes specific
membrane regions (membrane rafts) that contain glycosphingolipids and
cholesterol. Membrane rafts can form small pear-shaped invaginations called
caveolae which are lined with the protein caveolin. Caveolae have been shown
to contain some receptor proteins and facilitate certain types of receptor-
mediated endocytosis. Caveolae membrane domains have also been implicated
in cell signaling (Roch-Arveiller and Couderc, 2000).
Glycosphingolipids are distributed mainly at the surface o f the cell and
facilitate many cellular functions, such as regulation of cellular interactions
with its environment, cell-to-cell recognition and communication. They are
essential for the development and growth of organisms, for normal cellular
function (Bloch, 1991) and have been implicated in a number of serious
diseases, such as cancer as well as with infections, both viral and microbial
(Schuette et al. 1999). Studies in fibroblasts have suggested that 90% of
cholesterol is associated with the cell surface, with the remaining 10%
associated with internalized plasma membrane (Lange et al. 1991). Cholesterol
was originally suggested to play a role in endocytosis in the late 70’s (Heiniger
et al. 1976). Recent studies have implicated cholesterol in clathrin-mediated
endocytosis (Rodal et al. 1999) as well as in clathrin-independent endocytosis
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39
and shown it to be essential in the structure and invagination of caveolae
(Parton et al. 1994; Parton, 1996). Additionally, it has also been shown that
glycosphingolipid and cholesterol microdomains have the capacity to be
internalized and may also play an important role in signaling (Parton et al.
1994, Verkade et al. 1999). It has been suggested that the internalization of the
interleukin 2 receptor is mediated through these membrane raft microdomains
(Lamaze et al. 2001). Cholesterol has also been shown to be required for
proper functioning of the acetylcholine receptor (Criado et al. 1982). Other
studies suggest functional relationships between membrane cholesterol,
membrane transport and the actin cytoskeleton, mediated by annexin II
(Gruenberg and Emans, 1993).
Much of the research done to further elucidate the role of cholesterol and
membrane composition in endocytosis has utilized methyl-P-cyclodextrin
(MPCD), a membrane impermeable cyclic oligosaccharide which contains a
hydrophobic core and has been shown to extract cholesterol from the plasma
membrane (Ohtani et al. 1989, Kilsdonk et al. 1995). Subtil et al (1999)
showed that cholesterol depletion by MPCD inhibited endocytosis of
transferrin and that this inhibition was associated with the accumulation of
clathrin-coated pits at the site of endocytosis. Furthermore, Subtil et al (1999)
suggested that cholesterol depletion resulted in changes in the biophysical
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40
properties of the membrane, such as thickness, membrane tension and its
ability to bend. Ultimately these changes have been hypothesized to inhibit the
cell’s ability to the induce membrane curvature, a necessary step in the
formation of the coated pit and coated vesicle (Rodal et al. 1999; Subtil et al.
1999). In the present study, we have utilized M(3CD treatment of lacrimal
gland acinar cells to investigate the relationship between endocytosis and the
actin MF cytoskeleton.
Actin microfilaments and endocytosis: Much more than a simple scaffold, the
cytoskeleton is vital for many cellular functions, such as cell motility, mitosis,
muscle contraction and maintenance of cell structure, among others (see
Chapter 2, Introduction). Much of the research linking MFs to endocytosis
was established from studies of yeast (Munn, 2001). The need for cytoskeletal
participation seems to be evolutionarily conserved, as mutations in actin and/or
actin binding proteins have resulted in impaired endocytosis in yeast (Kiibler
and Riezman, 1993) as well as in some mammalian cells. However, although
insights from yeast are valid, it is important to be cautious when extrapolating
observations from yeast to mammalian cells, as substantial differences exit in
the endocytic mechanisms between them. Among the most relevant is the fact
that no dynamin counterpart has been found to facilitate endocytosis in yeast
nor has clathrin been shown to be vital for cell growth, membrane trafficking,
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41
secretion or receptor endocytosis in yeast (Geli and Reizman, 1998; Payne,
1990).
An intact MF network has been reported to be necessary for efficient
endocytosis in non-polarized mammalian cells (reviewed in Apodaca, 2001;
Sandvig and van Deurs, 1990; Durrbach et al. 1996) as well as in specialized
cells such as neurons (Schiavo and Stenbeck, 1998; Onofri et al. 2000) or
secretory epithelia such as the pancreas (Freedman et al. 1999) or the lacrimal
gland (present study). Additionally, the link between MFs and endocytosis has
also been established in endothelial cells (Alexander et al. 1998), MDCK cells
(Gottlieb et al. 1993), hepatocytes (Durrbach et al. 1996) as well as in Caco2
and T84 cell lines (Shurety et al. 1996; Song et al. 1999). Three main cell-
permeant toxins have been used to investigate the role o f actin MFs in
endocytosis: (a) cytochalasin D (CD), which inhibits MF polymerization by
capping the growing end of the filament (see Chapter 1, Introduction),
preventing further filament assembly and resulting in filament shortening
(Sampath and Pollard, 1991); (b) latrunculin, which binds to and sequesters
actin monomers, also results in filament depolymerization (Coue et al. 1987)
and (c) jasplakinolide, a marine toxin, which was initially implicated in
stabilization of actin filament assembly by interfering with actin treadmilling
(Bubb et al. 1994). Recent studies, however, suggested that while
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42
jasplakinolide may stabilize actin filaments in vitro, in vivo studies have
shown it to disrupt filaments by inducing polymerization of G-actin into
amorphous masses (Bubb et al. 2000). These data confirmed previous studies,
in lacrimal gland epithelial cells, of the depolymerizing effects of
jasplikinolide on F-actin (da Costa et al. 1998). It is not surprising, given their
varied effects on the actin network, that use of these MF-targeted in
establishing a link between endocytosis and MF has yielded mixed results,
depending on the cell type or assay utilized (Fugimoto et al. 2000).
In polarized epithelial cells, the role for MFs as facilitators of endocytosis is
more intricate, as macromolecules can be internalized from apical as well as
basal-lateral domains (Bomsel et al. 1989), utilizing both clathrin-dependent as
well as clathrin-independent mechanisms (Eker et al. 1994). Elucidating the
precise mechanism by which MFs facilitate endocytosis in polarized epithelia
recently became even more complex given recent data which suggested that
MFs may specifically facilitate apical, but not necessarily basal-lateral
membrane endocytosis (Jackman et al. 1994; Shurety et al. 1996).
The MF network has been hypothesized to facilitate one or more of the many
steps associated with endocytosis, reviewed in Qualmann et al (2000), as
follows:
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43
(a) Given the many accessory and regulatory proteins recently shown to
facilitate receptor-mediated endocytosis, it has been suggested that the
MF array may serve as a physical barrier, anchoring components of the
endocytic machinery, preventing their diffusion from the site where
they will be needed. In support of this hypothesis, Gaidarov et al
(1999) have shown an increase in the lateral-mobility of GFP-labeling
clathrin-coated pits in cells treated with latrunculin B.
(b) Contrary to the physical barrier theory, others have suggested a
negative role for MFs and a need for disassembly or rearrangement of
the actin cytoskeleton at the site of endocytosis, thereby opening a path,
so to speak, for endocytic vesicles to be internalized. Studies suggest
that the immediate vicinity of the clathrin-coated pit has very few actin
MFs (Fujimoto et al. 2000). Additionally, the need for cytoskeletal
rearrangement has also been suggested for viral entry into cells
(Pelkmans et al. 2002). However, other data suggest that MF
disassembly or rearrangement is not vital, as stabilization of the MF
array by jasplakinolide treatment had no effects on apical endocytosis
in MDCK cells (Shurety et al. 1998).
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44
(c) Actin MFs have been implicated in the initial steps of membrane
invagination, a necessary step which provides curvature to the
membrane, facilitating the formation and subsequent pinching off of
coated-pits by components of the endocytic machinery. It has been
suggested that MFs, together with the MF-dependent motor protein
myosin VI may generate actin-dependent forces for membrane
invagination or vesicle movement during the initial steps of endocytosis
(reviewed in Schafer, 2002).
(d) The pinching-off of the clathrin-coated vesicle, freeing it from the
plasma membrane, has been shown to be facilitated by dynamin (see
above, Dynamin). Various novel proteins have been identified which
can mediate association of dynamin to the actin cytoskeleton. Actin
binding protein 1 (Abpl) binds to dynamin’s PRD through its C-
terminal SH3 domain while its N-terminal domain has been shown to
associate with F-actin (Kessels et al. 2000). Membrane trafficking
studies have suggested that the dynamin-binding protein lasp-1, may
mediate the dynamin-MF interaction in apical endocytosis in parietal
cells (Okamoto et al. 2002). Additionally, it has also been suggested
that actin polymerization at the neck of the coated-pit could be used to
generate the force necessary for the membrane fusion event, although
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45
recent actin-depletion studies in CHO, A31 and Cos-7 cells argue
against this hypothesis, suggesting that actin filaments may play an
accessory but not an obligatory role in receptor-mediated endocytosis
and in endocytic coated vesicle formation (Fujimoto et al. 2000).
(e) Finally, it has been suggested that MF comet tails may provide the
driving force necessary for vesicle movement and internalization (Lee
and De Camilli, 2002). Comet tails were originally identified in studies
of mechanisms of propagation of pathogens such as Listeria
monocytogenes in infected cells (reviewed in Wu et al. 2000). Comet
tail facilitation of endocytosis has been suggested in recent studies
which found comet tails associated with endosomes and clathrin-coated
and secretory vesicles (Frischknecht et al. 1999; Merryfield et al.
1999). Additionally, dynamin and dynamin-binding proteins have also
been implicated in comet tail formation (Orth et al. 2002).
It is important to note that the above-mentioned hypotheses are not mutually
exclusive, as MFs may facilitate one or more of the endocytic steps utilizing
different mechanisms concomitantly. Additionally, different cell types may
utilize MFs, dynamin and other regulatory proteins in ways unique ways
bestsuited to the cell’s functional needs.
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46
Results:
Secretagogue stimulation results in the recruitment o f clathrin-associated
proteins from cytosolic to membrane protein pools at the apical plasma
membrane. Selective detergent extraction was used to isolate cytosolic
(saponin extractable), and membrane (Triton-X soluble and insoluble) protein
pools from cultured lacrimal gland acinar cells (see Methods). The effects of
secretagogue stimulation on the distribution of coat proteins associated with
clathrin-mediated endocytosis was analyzed by measuring the recovery of
clathrin, a-adaptin and dynamin within the three extracted protein pools.
Western Blot and densitometry analysis of protein distribution revealed that
secretagogue stimulation (CCH, 100 pM, 5 min) resulted in the recruitment of
clathrin, a-adaptin and dynamin from the soluble to the membrane pool
(Figure 3.3.), Extraction conditions were confirmed by measurement, within
each pool, of the marker proteins polymeric immunoglobulin A (IgA) receptor
(plgAR), a-tubulin, actin and lactate dehydrogenase (LDH) (Table 3.1.),
Treatments did not alter distribution of marker proteins. The distribution of
clathrin, a-adaptin and dynamin in control and secretagogue stimulated acini
(CCH, 100 pM) were further characterized by confocal microscopy. Lacrimal
gland acinar cells were cultured for three days on Matrigel-coated coverslips
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47
a-Adaptin
60
50 -
40 -
30
20
10
0
n
Cytosolic
lid
□ CON
■ CCH
TX-100
soluble
TX-100
resistant
60
■ a 5 0
g 40
30
o 20
S 5 10
0
Clathrin
ri 1 1
O l
□ CON
■ CCH
Cytosolic TX-100
soluble
TX-100
resistant
Dynamin
■ a 50
g 40-
l- 30
o 20 -
S 5 10 -
0 - -
Cytosolic TX-100 TX-100
soluble resistant
□ CON
■ CCH
Figure 3.3. Recruitment o f coat-proteins following secretagogue
stimulation. Soluble, Triton X-100-resistant and Triton-X-100 soluble
proteins were extracted from control and CCH stimulated acini (see
Methods) and investigated for the recovery of clathrin, a-adaptin and
dynamin. Secretagogue stimulation resulted in a loss of coat-protein
association with the soluble protein pool and a concomitant significant
increase in association with the membrane (TX-100 soluble) pool.
(n=7, p < 0.05).
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48
Control CCH 100 pM
Saponin Tx-100 SDS Saponin Tx-100 SDS
Tubulin 17.22 + 2.40 4.56 + 2.45 78.10 + 3.80 19.98 + 1.81 5.92 + 4.01 74.10 + 3.82
Actin 35.26 + 2.93 8.56 + 2.84 56.21 + 5.18 29.78 + 1.53 10.98 + 3.61 59.24 + 2.67
LDH 59.39 + 4.18 33.96 + 4.19 6.65 + 2.14 58.10 + 2.67 39.16 + 2.46 2.73 + 0.79
plgAR 3.52 + 0.85 79.62 + 5.39 16.85 + 5.05 3.05 + 0.64 77.67 + 4.44 19.23 + 4.68
Table 3.1. Marker protein confirmation o f extraction conditions.
Western blot analysis of the distribution of the marker proteins (actin,
plgAR, a-tubulin and LDH) was used to confirm extraction conditions
(see Methods, n=7). CCH stimulation had no effects on the recovery of
marker proteins in soluble, TX-100 soluble and TX-100 resistant
protein pools.
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49
Microvilli
Figure 3.4. Actin M F distribution in lacrimal acini. Lacrimal acini were
cultured then processed for confocal microscopy (see Methods).
Rhodamine-phalloidin staining of MFs reveals thick lumenal actin rings,
associated with the apical actin terminal web and microvilli. MFs
staining can also be seen at individual cell basal-lateral membrane (thin
lines). Treatment with CD resulted in fragmentation of the MF network
as seen by the discontinuous apical and basal-lateral MF staining. (*,
apical lumen). Bar = 10pm.
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50
then processed for immunofluorescence and confocal fluorescence microscopy
analysis (see Methods). Staining of the MF array by rhodamine-phalloidin was
used to resolve the location of apical/lumenal regions. The presence of the
actin terminal web and lumenal microvilli (enriched in MFs) at the apical
plasma membrane (APM) was seen, in the confocal images, as a thick MF ring
around the apical lumen (*), while thin lines of actin staining denote actin
associated with the basal lateral membrane (Figure 3.4.), Carbachol
stimulation (CCH, 5 minutes) resulted in a dramatic redistribution of clathrin,
a-adaptin and dynamin (green) to the APM as seen by the increased staining
intensity of all three proteins at the apical region (Figure 3.5., arrow).
Stimulation with PMA (1 pM, 5 min) resulted in a similar apical recruitment of
coat proteins (data not shown). A time-course analysis of CCH stimulation (5,
15, 30 and 60 minutes) revealed that by 15 minutes of stimulation, the apical
staining of all three proteins had returned to what resembled the staining seen
in resting acini, suggesting that secretagogue stimulation elicited an immediate
rapid recruitment of coat proteins to the APM followed by coat protein
retrieval from the APM at later time-points.
Treatment with CD blocks the retrieval o f clathrin-associated coat-proteins
from the APM and inhibits FITC-dextran uptake. In order to investigate
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whether MFs facilitated the coat-protein retrieval seen at later time-points of
carbachol stimulation, lacrimal acini were treated with the fungal metabolite
cytochalasin D (CD, 5 pM, 60 min, 37°C), prior to secretagogue stimulation
(CCH, 100 pM, 15 min), then processed for confocal fluorescence microscopy
analysis (see Methods). CD has been shown to cap the ends of actin
microfilaments, resulting in filament shortening and fragmentation (Cooper,
1987). Confocal fluorescence microscopy revealed that, as expected, CD
treatment resulted in discontinuities of the apical/lumenal actin terminal web
(arrowheads), as well as fragmentation of basal lateral membrane-associated
actin, as seen by the punctate rhodamine phalloidin staining. (Figure 3.6., *,
apical lumen). Unlike the carbachol-stimulated acini, in which the intensity of
apical coat-protein staining decreased by 15 min of secretagogue stimulation,
pretreatment with CD prior to CCH blocked the retrieval of clathrin, a-adaptin
and dynamin, as suggested by the continued intense apical staining of all three
proteins (arrow). Interestingly, CD treatment alone resulted in apical
accumulation of coat-proteins, suggesting that constitutive endocytosis of
apical membrane following basal secretion may also be facilitated by clathrin.
CD-induced accumulation of coat proteins at the basal-lateral membrane was
also occasionally seen (data not shown), however, given the much larger area
of the basal-lateral membrane compared to that of the apical plasma
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52
Resting CCHSmin CCHlSmin
Figure 3.5. Coat-protein recruitment to the APM following CCH
treatment The distribution of clathrin, a-adaptin and dynamin in
control and CCH-stimulated acini (CCH, 100 pM) was investigated
using confocal microscopy. Early time-points of CCH-stimulation (5
min) resulted the recruitment of all three coat-proteins to the APM, as
seen by the increased apical staining. Continued stimulation (15 min)
resulted in decreased apical coat protein staining resembling control
(resting) acini. (*, apical lumen). Bar s 10pm.
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53
Resting CD CD+CCH15min
Figure 3.6. Disruption o f the MF network by CD inhibits coat
protein retrieval following secretagogue stimulation. Lacrimal acini
were pre-treated with CD (5 pM, 60 min) prior to CCH stimulation
(lOOpM) then processed for confocal microscopy. Disruption of actin
MFs by CD blocked the retrieval of clathrin, a-adaptin and dynamin
at later time-points (see Figure 3.5., 15 min) of secretagogue
stimulation. ( *, apical lumen). Bar = 10pm.
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54
membrane, coat protein accumulation was not as intense and much harder to
visualize by confocal immunofluorescence microscopy. It was our hypothesis
that the coat protein accumulation elicited by CD correlated with a functional
inhibition of endocytosis. To test this idea, we utilized flow cytometry analysis
(FACs) of FITC-dextran uptake in lacrimal acini. Acini were cultured for three
days then stimulated with CCH (15 min, 100 pM, 37°C) with and without CD
pre-treatment (Figure 3.7). FITC-dextran (1.25 mg/ml, 15 min) was added
together with and without CCH for equal time (15 min) in control and CD-
treated acini (see Methods). The increased membrane trafficking elicited by
secretagogue stimulation resulted in a significant ( *, P < 0 .05, n = 4) increase
in FITC-dextran uptake in CCH-treated acini, as compared to control,
unstimulated acini. Pre-treatment with CD blocked the stimulated response
seen with CCH-treatment. Additionally, acini treated with CD showed a
significant reduction in FITC-dextran uptake, compared to control (#, p <
0.05). This paralleled the confocal fluorescence microscopy data in which
unstimulated acini treated with CD showed an accumulation of coat proteins at
the apical membrane. These FACs data suggested CD-treatment disruption of
MF involvement in constitutive secretion. It is important to note that FACs
analysis measured total uptake, making no distinction between basal or apical
uptake. However, given the dramatic accumulation of coat-proteins at the
APM in CD-treated acini (Figure 3 .6 ), these data suggested to us that at least a
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55
cd+cch
Figure 3.7. CD-treatment inhibits FITC-dextran uptake in
lacrimal acinar cells. Control and CCH-stimulated acini were
exposed to FITC-dextran (with and with CD pre-treatment) then
processed for flow cytometry analysis (see Methods). CCH-
stimulation resulted in a significant increase of FITC-dextran
uptake ( *,P<0 .05). Pre-treatment with CD block the secretagogue
response. Additionally, uptake of FITC-dextran in CD-treated
unstimulated acini was significantly lower than control (#, p <
0.05) (n = 4).
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56
component of the inhibition of FITC-dextran uptake was due to inhibition of
apical endocytosis.
Disruption o f the MF network by CD results in the accumulation o f clathrin-
coated pits at the APM. Our results implicated MFs as facilitators of clathrin-
mediated endocytosis at the APM of lacrimal gland acinar cells. However,
confocal microscopy could not resolve at what step of clathrin-coated pit or
clathrin-coated vesicle formation MF involvement became pivotal. To further
elucidate the step at which MF integrity was required, we utilized electron
microscopy to examine the distribution of coated pits and coated vesicles in
resting and stimulated acini, with and without CD pre-treatment (see Methods).
Initial characterization of control samples revealed that the apical cytoplasm of
cultured acinar cells frequently included clusters of pleomorphic, secretory
granules (SG) characteristic of lacrimal acinar cells in situ. In many sections
the apical cytoplasm included a delicate, particulate layer immediately beneath
the apical cell membrane, which in some sections was organized as very
delicate filaments. Additionally a layer of densely aggregated apical MFs could
be seen in control acini (arrowhead, Figure 3.8A.). Coated pits and coated
vesicles were evident within the apical cytoplasm the apical region of control
(Figure 3.8B.) and carbachol-stimulated acini (arrows, Figure 3.8C ). More
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57
SG ■
V ‘ \
/ * > * - *
■ K
*?
mi
*s ^
„ i v i £ i , i*1 -*
y >
% ■ * » ■ • -
#
* V *
***' * *
ccw
Figure 3.8. Electron microscopy o f MF and coat protein
distribution in resting and stimulated acini. Lacrimal acini
were processed for electron microscopy (see Methods) then
analyzed for the distribution of clathrin-coated pits and vesicles
in control (Figure 3.8B) and stimulated (Figure 3.8C.) lacrimal
gland acini. Long continuous apical filaments are shown in
control acini (Figure 3.8A., arrowhead). Occasional coated pits
and vesicles were seen in control and CCH-treated acini
(arrows; L, apical lumen)
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58
CO+CCH
Figure 3.9. Accumulation o f clathrin-coated pits at A PM
resulting from CD treatment., Treatment with CD in the
presence or absence of CCH resulted in a dramatic increase in
apical coated pits (arrows) as seen by electron microscopy.
Electron microscopy analysis of the MF network in control
(Figure 3.8A.) and CD-treated lacrimal acini (Figure 3.9B.)
confirmed the fragmentation of apical MFs by CD (arrowhead).
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59
notable, however, was the dramatic increase in the number of partially formed
coated pits at the APM in CD-treated acini (Figure 3.9A, arrow). Similarly,
coated-pit accumulation was seen in stimulated acini pre-treated with CD. The
effects of CD treatment on the MF network were also confirmed by electron
microscopy. Previous confocal microscopy had shown punctate, discontinuous
MF staining resulting from CD treatment (Figures 3.4. and 3.6.), Compared to
the continuous MF distribution seen in control acini (Figure 3.8A.) electron
microscopy revealed fragments of apically located MFs in the CD-treated
samples (arrow, Figure 3.8B.),
Inhibition o f endocytosis by methyl- (i-cyclodextrin results in the
accumulation o f clathrin, a-adaptin and dynamin at the apical plasma
membrane. Treatment of lacrimal acini with CD demonstrated that an intact
MF network was necessary for coat protein retrieval following secretagogue
stimulation. Additionally, FITC-dextran uptake experiments suggested that the
apical accumulation of clathrin, a-adaptin and dynamin in CD-treated acini
was associated with the inhibition of endocytosis (Figure 3.7.), What these
data did not address, however, was the exact relationship between endocytosis,
the actin cytoskeleton and coat-protein distribution; that is, if disruption of the
MF network resulted in inhibition of endocytosis, was the contrary also true:
would inhibition of endocytosis affect the MF network and/or the distribution
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60
of coat proteins? In order to address this question, we first inhibited
endocytosis by exposing cultured lacrimal gland acinar cells to methyl-P-
cyclodextrin (MPCD). M(3CD has been shown to inhibit endocytosis by
selectively extracting cholesterol from the plasma membrane of cultured cells
(Kilsdonk et al. 1995). Additionally, previous studies have also specifically
suggested a role for cholesterol in clathrin-mediated endocytosis (Rodal et al.
1999). FACs analysis of FITC-dextran uptake was utilized to confirm the
effects of endocytosis inhibition resulting from MPCD treatment. Acini were
cultured for three days then treated with MPCD (1 mM, 30 min) prior to CCH
stimulation (100 pM, 15 min) and incubation with FITC-dextran (1.25 mg/ml,
15 min) (see Methods). As previously seen (Figure 3.7.), CCH stimulation
resulted in a significant increase in FITC-dextran uptake (*, p < 0.05, n = 4,
Figure 3.10.), No significant increase of FITC-dextran uptake was seen in
acini pre-treated with MPCD. This block in the stimulated response suggested
that pretreatment of acini with MPCD resulted in inhibition of the
secretagogue-stimulated response in membrane trafficking as well as an
inhibition of the endocytic response normally elicited by CCH-stimulation.
Confocal microscopy analysis of MpCD-treated acini was then utilized to
investigate whether inhibition of endocytosis would result in Ghanges in coat
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protein distribution. Lacrimal acini were cultured for three days then processed
for immunofluorescence (see Methods). Labeling of actin MFs (top) was used
to locate apical lumens (*, Figure 3.11.), Confocal fluorescence microscopy
indicated that pre-treatment with MPCD (ImM, 30 min) prior to CCH
stimulation (100 pM, 15 min) resulted in apical accumulation of a-adaptin
(green, middle), as seen by the intense staining (arrows), compared to control
(unstimulated) acini. These data, together with the inhibition of FITC-dextran
seen by FACs analysis, suggest that coat protein accumulation at the APM is
associated with inhibition of endocytosis, in lacrimal acinar cells. Additional
confocal microscopy studies (Figure 4.11.) address the effects of MPCD
treatment on the actin MF network.
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62
250 T
'I 200
CON CCH MBCD MBCD+CCH
Figure 3.10 M/3CD inhibits the uptake o f FITC-dextran in
lacrimal acini. Lacrimal acini were cultured for three days then
stimulated with CCH (lOOpiM, 15 min) in the presence of FITC-
dextran (1.25 mg/ml, 15 min), with and without MPCD (ImM, 30
min) pre-treatment. CCH stimulation resulted in a significant (*,
p < 0.05, n=4) increase in FITC-dextran uptake, as measure by
FACs analysis. Pre-treatment with MPCD blocked the CCH-
stimulated response.
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63
MpCD+ CCH
Figure 3.11. Apical accumulation o f a-adaptin in MfiCD-treated
acini. Lacrimal acini were pre-treated with MpCD (1mm, 30 min)
prior to CCH (100 pM, 15 min) stimulation then processed for
confocal microscopy (see Methods). Rhodamine-phalloidin
staining of MFs (top) is used a tool to distinguish apical lumens (*).
Pre-treatment with M3 CD resulted accumulation of a-adaptin at
the APM similar to that seen in CD-treated acini (Figure 3.5.) Bar
= 10pm.
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64
Discussion
Clathrin-mediated endocytosis is a well-established mechanism by which cells
internalize plasma membrane. The studies described herein were initiated to
resolve the role of clathrin and the actin MF network in the retrieval of APM
following secretagogue-stimulated secretion in lacrimal gland acinar cells.
Selective detergent extraction of lacrimal gland acinar lysate revealed that
clathrin and the clathrin-associated proteins dynamin and a-adaptin were
distributed among the three different extracted protein pools (cytosolic or
saponin soluble; membrane or TX-100 soluble and cytoskeletal or SDS
soluble). Western blotting and densitometry analysis of protein distribution
among the three pools suggested that clathrin, dynamin and a-adaptin are
recruited from the saponin-soluble protein to the Triton X-100 soluble protein
pool in response to carbachol stimulation (Figure 3.3.). Confocal
immunofluorescence microscopy was used to further characterize the effects of
secretagogue stimulation of the distribution of clathrin and clathrin-associated
proteins. In resting, unstimulated acini, the distribution of clathrin was
punctate, with minimal association seen with the apical and basolateral (BLM)
membranes. Similar labeling was found with a-adaptin and dynamin (Figure
3.5.), CCH stimulation resulted in a dramatic increase in the apical
accumulation of all three proteins, as seen by their increased
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65
immunofluorescence staining. This suggested to us that CCH treatment results
in the recruitment of clathrin and clathrin-associated proteins from a soluble
(maybe reserve) protein pool, specifically to an apically associated membrane
pool. Although clathrin has been shown to also mediate trafficking at the
basolateral membranes in many epithelial cells, no significant increases in
immunofluorescence staining of clathrin, a-adaptin or dynamin, following
CCH treatment were seen associated with the BLM regions in lacrimal acini.
We cannot, however, rule out this possibility, as coat-protein accumulation at
the BLM is much harder to resolve by confocal microscopy, given the much
larger surface area covered by the BLM as compared to the APM. The
decrease in apical staining of clathrin, dynamin and a-adaptin at later time-
points of secretagogue stimulation suggests a quick internalization of coat
proteins, following the initial apical recruitment. Although similar
secretagogue-induced changes in coat-protein distribution have been shown in
other cell types (Pauloin et al. 1997; Valentijn et al. 1999), the effects of
secretagogue on clathrin and clathrin-associated proteins cannot be
generalized. Recent studies suggest a role for clathrin in membrane trafficking
in resting, but not stimulated parietal cells (Okamoto et al. 2000).
In order to determine whether the MFs facilitated the retrieval of coat proteins
following stimulation, lacrimal acini were pre-treated with the fungal
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66
metabolite cytochalasin D (CD) prior to exposure to CCH. We confirmed that
any differences in the stimulated apical coat-protein recruitment in CD-treated
acini would not be the result of the acini’s inability to properly secrete, as
previous work in our lab has shown that disruption of the MF network by CD
does not inhibit protein release induced by secretagogue stimulation (da Costa
et al. 1998). Confocal microscopy of CD-treated stimulated acini revealed that
MF disruption blocked coat-protein internalization following CCH treatment,
as seen by the continued intense apical staining of coat proteins (Figure 3.6.),
These data suggested that an intact MF network is necessary for clathrin-
mediated endocytosis in stimulated as well as in resting acini in lacrimal acini
as CD treatment of unstimulated acini also resulted in apical accumulation of
coat proteins.
Disruption of actin filaments has been associated with blocks in various stages
of endocytosis in mammalian cells (Durrbach et al. 1996). The involvement of
MFs in early stages of endocytosis (Sandvig and van Deurs, 1990; Gottlieb et
al., 1993; Jackman et al., 1994; Durrbach et al., 1996; Shurety et al., 1996)
as well as in later stages, such as membrane trafficking to the late endosome
and lysosome (van Deurs et al., 1995), or trafficking of ligands to
degradative compartments (Durrbach et al. 1996) have been characterized.
Additionally, depending on the cell type studied, MF disruption has been
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67
shown to selectively and specifically inhibit apical (Gottlieb et al. 1993;
Jackman et al. 1994) or basolateral (Sandvig and van Deurs, 1990)
membrane trafficking. We utilized uptake of FITC-dextran to investigate MF
involvement in endocytosis in resting and stimulated lacrimal acini treated
with and without CD treatment. Secretagogue stimulation resulted, as
expected, in a significant increase in FITC-dextran uptake, given the
increased membrane trafficking elicited by CCH treatment (Figure 3.7.).
Pre-treatment with CD blocked this CCH-stimulated response. Additionally,
FITC-dextran uptake was significantly lower in CD treated, unstimulated
acini, compared to control. Although culture conditions of lacrimal acini
made it impossible to distinguish apical from basolateral uptake, given the
CD-induced coat-protein accumulation at the APM of lacrimal acini seen with
confocal microscopy, these data suggested to us that at least a component of
the inhibition of FITC-dextran uptake in CD-treated acini was due to apical
endocytosis.
We further characterized the relationship between an intact MF array and
endocytosis of clathrin-coated vesicles by electron microscopy. Electron dense
clathrin-coated pits and vesicles were identified at the APM and subapical
region of control and CCH-stimulated acini (Figure 3.8.), Treatment with CD
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68
resulted in a dramatic accumulation of coated pits at the APM in resting and
stimulated acini (Figure 3.9.), with few or no clathrin-coated vesicles seen.
These data suggested to us that disruption of the MF array blocks clathrin-
mediated endocytosis at the coated pit stage and that MFs facilitate the
formation of clathrin-coated vesicles.
The effects of CD treatment on lacrimal acini suggested that MF disruption
inhibited endocytosis. It did not, however, address the question of whether the
reverse was also true: did inhibition of endocytosis alter the MF network. To
address this question, acini were treated with M|3CD, which has been shown to
inhibit endocytosis by extracting cholesterol from the plasma membrane (see
Introduction). FITC-dextran uptake studies of lacrimal acini confirmed
inhibition of endocytosis in M(3CD-treated acini (Figure 3.10.), Although
confocal microscopy data suggests apical accumulation of coat proteins in
stimulated acini, pre-treated with M0CD, no changes were seen in the MF
network resulting from M0CD treatment, as will be discussed further in
Chapter 4.
These data, together with those of CD-treated acini, suggested to us that the
apical coat protein accumulation seen by confocal fluorescence microscopy
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69
was associated with inhibition of endocytosis. Additionally, our data
indicated that, in lacrimal gland acinar cells, an intact MF network was
necessary for clathrin-mediated endocytosis. The mechanisms which may
facilitate the interaction between actin MFs and clathrin, as well as the
regulatory and accessory proteins which may mediate this endocytic process
are investigated in C hapter 4.
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70
Chapter 4 - Actin Regulatory Proteins in Endocytosis
Introduction:
Much of the initial research on clathrin-mediated endocytosis focused on the
role of adaptors and dynamin, essential components of this endocytic process.
However, years of research, much of it done in neurons, has shown that coat-
protein formation and internalization is a highly complex process involving a
variety of accessory proteins and signaling cascades responsible for activating
and regulating receptor-mediated endocytosis. Additionally, accessory proteins
mediate the interaction between coat proteins and the cytoskeleton, particularly
the actin MF array, a key player in coat-protein formation and in membrane
trafficking. This chapter will characterize the function of some of these
accessory proteins in lacrimal gland epithelial cells, particularly, the
association between accessory proteins and the GTPase dynamin. Although
dynamin has been implicated directly in the scission of coat pits in the initial
stages of endocytosis (see Chapter 3), it has also been shown to play a broader
role in endocytosis by virtue of interactions of its carboxy-terminal proline rich
domain (PRD) with a variety of novel Src homology 3 (SH3)-containing
proteins (reviewed in Hinshaw, 2000).
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71
Src homology 3 (SH3) domains: The SH3 domain is one of the best
characterized member of an ever growing family of protein interaction
modules and was one of the first modular protein interaction domains
identified almost a decade ago (Ren et al. 1993). SH3 domains (50-70
residues) are found in many proteins that mediate protein-protein interactions
and are important in cytoskeletal architecture and intracellular tyrosine kinase
mediated signaling (Musacchio et al., 1994; Pawson, 1995; Cohen et al., 1995).
Additionally, SH3 domains have been implicated in various cellular processes,
such as regulation of enzymes and mediation of the assembly of multiprotein
complexes. They have been identified in literally hundreds of proteins, and
have been characterized in species ranging from yeast to humans (reviewed in
Mayer, 2001).
Signal transduction is accomplished through binding of the core conserved
binding SH3 domain motif, identified as PxxP (where P is proline and X is any
amino acid), to PRD motifs of ligand proteins (Ren et al., 1993; Yu et al.,
1994). Given their association with such a vast number of biological events
and the wealth of genetic, biochemical and structural data available on SH3
domains, it is not surprising to note that they have been targeted for use in
therapeutics, for example, as anti-proliferative agents, among others (reviewed
in Vidal et al. 2001). Although SH3 domains have been extensively studied,
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72
much is still unknown about SH3-ligand interactions, such as whether SH3
domains or ligand sites can engage in simultaneous and multiple interactions.
This question is complicated by the fact that, given the many existing different
ligands and SH3 domains, there exist potentially hundreds of possible PRD-
SH3 interactions. Basically, one of the questions that still remains to be
elucidated is how SH3-PRD binding specificity is accomplished. Some have
argued that, given the very low binding affinity and marginal selectivity of
SH3-ligand binding, there must be a role played by additional regulatory
mechanisms (Ladbury and Arold, 2000). Others have suggested that selectivity
is enhanced by multiple interactions, such as is the case with the Grb2 protein
or the Nek adaptor, which has multiple SH3 domains that could theoretically
associate (simultaneously?) with different PxxP sites, activating different
signaling regulatory cascades (Simon and Schreiber, 1995; Adler et al. 2000)
The SH3-WASP connection: Wiskott-Aldrich Syndrome (WAS) is a rare and
severe X-linked immunodeficiency first described in the mid 30’s by Wiscott
(1936) and is characterized by recurrent opportunistic infections, eczema and
thrompocytopenia (reviewed in Snapper and Rosen, 1999). The WAS gene
was first isolated by Derry et al (1994) and the observation that WAS patients
demonstrated both cellular signaling and cytoskeletal abnormalities led to the
realization that WAS and WAS-related proteins played a vital role in
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73
regulation of the actin cytoskeleton and of MF dynamics (reviewed in Mullins,
2000). The WAS protein (WASP) consists of 502 amino-acids and contains
several modular domains (Figure 4.1.), including a WASP homology 1
(WHl)/pleckstrin homology (PH) domain (a PtdIns(4,5)P2-binding site), a
basic residue (BR)-GBD domain which binds to Cdc42 (a Rho family
GTPase), a polyproline domain and a VCA (verproline-cofilin-acidic) C-
terminal region, both of which have been implicated in actin regulation
(Meyer, 2001). WASP’s BR-GBD domain can bind to and inhibit the VCA
region, when it is not bound to Cdc42. Consequently, the BR-GBD domain
functions as a biological switch, inhibiting VCA activity in response to cellular
changes in GTP-Cdc42 levels (Abdul-Manan et al., 1999; Kim et al., 2000;
Miki et al., 1998). A connection between the MF cytoskeleton, WASP and
Cdc42 was initially demonstrated in microinjection studies of aortic
endothelial cells, where it was shown that dominant negative Cdc42, but not
dominant negative Racl or Rho, blocked the WASP-induced formation of
actin clusters (Symons et al. 1996). Recent studies have implicated WASP in
cytoskeletal regulation of lymphocytes (Westerberg et al. 2001), as well as in
cellular functions as varied as secretion (Frantz et al. 2002), signaling (Bauch
et al. 2000), T and B cell activation (Anton et al. 2002) and in extracellular
matrix degradation (Mizutani et al. 2002). Association of WASP with SH3
domains is thought to occur via its extensive proline-rich region. Evidence of
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74
W iskott-Aldrich Synd
—
rom e Pro!
— H -
cm
m m m m m m m mm «
■ ■ ■ ■ ■ ■ ■ ■ M r
J
w K v riii/
i
i y i/ltK .U b ,l^ lr/sii V V i . w < V li Ul A » .
1 \ /
wprwif’
T O t/f f ljW M f ’ borooiogy
(i 8 I> ; V a s * binding i i m
homology dot»atorW A R ; Cbfilta to
Polypro lines: (with SH I domains)
Cdc-CRac
Snapper anil 1
irin homology do
wndtogy domain
SIP domains Actio
town, Anna, : Rct. immuawt 17:9i>S-2t>dS,?‘ J
nM nrBR; Ifasfc residsa's;
.A SI* f (Oiiiolo«v JAerproiin
and acidic residues;
Figure 4.1. WASP domain structure. The WASP protein contains
various modules implicated in endocytosis and MF organization.
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75
SH3-WASP binding has been suggested in a variety of studies, including the
Nek SH2/SH3 adaptor, which has been implicated in the recruitment of
neuronal-WASP (N-WASP) and the Apr2/3 complex in vaccinia virus,
implicated in the formation of comet tails (Frischknecht et al. 1999). Studies in
yeast have also identified SH3-domains of unconventional myosins as potential
N-WASP partners (Li, 1997).
The WASP VCA domain andArp2/3: The VCA domain binds to and activates
the Arp2/3 complex, resulting in MF assembly from de novo nucleation of
actin, that is, the polymerization of monomers to form a new filament.
Additionally, Arp2/3 activation has been shown to induce actin filament
branching (Blanchoin et al., 2000; Mullins et al., 1998; Pantaloni et al., 2000).
Arp2/3, was originally identified in the early 90’s, and contained two
unconventional actins, now knows as Arps, or actin-related proteins
(Machesky et al. 1994). It has a mass of approximately 200 kD and is
composed of seven highly conserved subunits: p40, p35, p21, pl9, pl5, Arp2
and Arp3. The name “Arp2/3” was coined when further analysis of both
Arps revealed that the larger of the two (47kD) showed sequence homology
to Arp3 genes in other species, while the smaller (44kD) resembled Arp2
(Kelleher et al. 1995). The Arp2/3 complex is conserved from yeast to humans
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76
with high sequence identity, between 60-70% (Machesky et al. 1997; Welch et
al. 1997).
Since it’s discovery, exhaustive research on Arp2/3 has shown it to be vital, if
not one of the most important players regulating actin remodeling and
dynamics (reviewed in May, 2001). F-actin filament formation requires
association of globular (or G-actin) monomers into dimers and trimers, a
process that has been shown to be energetically unfavorable, compared to
elongation of pre-existing filaments (Frieden, 1983; Mullins et al. 1997). One
theory suggests that Arp2/3 can overcome this energetic barrier and nucleate
the formation of a new filament as follows: Arp2 and Arp3 residues, located
side-to-side within the Arp2/3 complex, would form a heterodimer, or a
“pseudo” barbed end, resulting in a favorable structural orientation, similar to
the actin MF barbed end, bypassing the kinetically unfavorable dimerization
step (Kelleher et al. 1995). However, this hypothesis cannot be confirmed as
no crystallographic data has been obtained for any of the Arp2/3 subunits.
Additionally, recent data has shown that there is no close interaction between
Arp2 and Arp3 within the complex (Mullins et al. 1997). Others have
suggested that Arp2/3 activation could result in a conformational change that
would bring Arp2 and Arp3 together, however, no experimental evidence has
yet been shown which confirms or rejects this theory. Additionally, Mullins et
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77
al (1998) have shown that Arp2/3 can bind to the pointed end, as well as to the
side of actin filaments, stimulating nucleation and also F-actin branch
formation.
Arp2/3 activation: Although Arp2/3 activation of actin polymerization has
been shown to be weak, its activity is dramatically enhanced by a variety of
regulatory molecules (Mullins et al. 1998). The first identified activator of
Arp2/3 activity was the bacterial protein ActA, a protein originally known as
Scar (for the human homologue of a suppressor of cyclic AMP receptor
mutation in Dictyostelium discoideum), now known as WASF1 (for WASP-
family member 1) (Bear et al. 1998; Welch et al. 1998; Machesky and Insall,
1998). Subsequently, many other activators have been identified, such as
cortactin (Uruno et al. 2001), and type I myosins, in yeast (Lechler et al. 2000).
Activators are thought to interact with Arp2/3 through binding to its acidic
motif, a conserved domain rich in aspartic and glutamic acid residues
(Machesky et al. 1998). Additionally, Abplp (actin binding protein lp) has
also been found to activate Arp2/3. Abplp was the first actin-associated
protein identified in yeast (Drubin et al., 1988). Data suggest that Abplp does
not bind to Arp2/3, but is responsible for recruiting the complex to the sides of
actin filaments to promote nucleation (Goode et al. 2001). Activation of
Arp2/3 by WASP family members requires binding of WASP to G-actin,
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78
through WASP’s WH2 or V (verprolin homology) domain (Rohatgi et al.
1999). It has been suggested that WASP can shuttle actin monomers to Arp2/3,
thereby facilitating actin filament nucleation (Egile et al. 1999). This activity
is similar to that of profilin, a small (14kD) protein, which has numerous
binding partners, including Arp2/3, and which was demonstrated to be able to
bind to G-actin and also shown to “shuttle” actin monomers to the barbed end
of actin filaments (Pollard et al. 2000). Others have suggested the WASP
molecular interactions may be more complex, requiring synergistc activity
between various WASP G-actin binding sites (Marchand et al. 2000).
Elucidating the regulatory mechanisms of Arp2/3 filament nucleation is a
complex undertaking, given the almost bewildering array of molecules known
to interact with Arp2/3 activators (May, 2001). Cdc42, a small GTPase, was
identified as the first Arp2/3-activator binding partner, and shown to interact
with WASP through its conserved GBD domain (Aspenstrom et al. 1996;
Abdul-Manan et al. 1999). Additionally, the acidic phospholipid
phosphatidylinositol 4,5 bisphosphate (PIP2) has been shown to interact with
WASP (Abdul-Manan et al. 1999). Both have been demonstrated to activate
Arp2/3-dependent polymerization of actin in cell extracts (Ma et al. 1998). One
mechanism of Cdc42 and PIP2 regulation of Arp2/3 was demonstrated in terms
of signaling through WASP. WASP is maintained in an inactive state - the
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79
WH2 and acidic C-terminus domains are kept in a closed “hidden”
conformation by binding to WASP GBD domain. In this autoinhibited state,
WASP can bind to Arp2/3, but cannot activate its nucleating activities. Binding
of Cdc42 or PIP2 partially opens this inhibited state however, simultaneous
binding of both molecules is required to fully open the molecule and stimulate
actin polymerization (Rohatgi et al. 1999; Preboda et al. 2000). Although
Ccd42 and PIP2 are the two most characterized regulators of Arp2/3 activators,
many others have been identified (reviewed in May 2001).
SH3 domains and dynamin: The importance of dynamin’s PRD-SH3
interactions cannot be underestimated and is strengthened by the fact that the
list of SH3-domain containing proteins which associate with dynamin in
continuously increasing and includes amphiphysin I and II (David et al. 1996;
Leprince et al. 1997), endophilin (originally called SH3p4; Micheva et al.
1997; Ringstad et al. 1997 and 1999), syndapins I and II (Qualmann et al.
1999; Qualmann and Kelly, 2000), actin binding protein 1 (Kessels et al. 2000)
and synaptojanin (McPherson et al. 1994 and 1996), among many others. PRD
binding to SH3 domain-containing proteins has been shown to stimulated
dynamin’s GTPase activity (Gout et al. 1993) and disruption of dynamin-SH3
domain interactions have been shown to impair endocytosis (Shupliakov et al.
1997). The majority of these dynamin partners were originally characterized in
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80
neuronal tissue. Ongoing studies, however, reveal that these dynamin-
associated proteins, discussed below, facilitate clathrin-mediated endocytosis
in a variety of systems, including lacrimal gland acinar cells (present study).
Amphiphysin and endophilin: One of the best-characterized dynamin-
associated proteins is the SH3-domain containing protein, amphiphysin (125
kD). Studies in nerve terminals have shown it to co-localize with dynamin
(McPherson et al 1994) and synaptojanin, a 145 kD protein with inositol-5 -
phosphatase activity (see, below; McPherson et al. 1994, 1996). It has been
hypothesized that amphiphysin is responsible for directing dynamin to coated
pits by binding both to dynamin and a-adaptin (David et al. 1996). Shupliakov
and colleagues (1997) demonstrated that endocytosis was blocked in cells
microinjected with the SH3 domain of amphiphysin or with a peptide
containing dynamin’s SH3-binding site. This also resulted in an increased
number of coated pits at the plasma membrane of synapses. Recent studies, in
which amphiphysin’s SH3 domain was overexpressed, showed a block in
endocytosis of the GLUT4 glucose transporter (Volchuk et al. 1998),
reinforcing the importance of amphiphysin’s SH3 domain in endocytosis.
Additionally, amphiphysin’s interaction with dynamin and endophilin has also
been implicated in generation of membrane curvature, a necessary step in the
initial stages of coated-pit formation (Huttner and Schmidt, 2002).
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81
Endophilin, originally identified by Ringstad and colleagues (1997), has been
proposed to play a role in both early stages (membrane deformation) and late
stages (vesicle fission and uncoating) of clathrin-mediated endocytosis (Gad et
al., 2000; Schmidt et al., 1999; Ringstad et al. 1999), although much of its
mechanism of action is still unknown. It was demonstrated to bind to dynamin
and synaptojanin, as SH3 domain endophilin mutants lose their ability to bind
to dynamin and synaptojanin (Ringstad et al. 1997; Schmidt et al. 1999).
Depletion of endophilin from rat brain cytosol was shown to inhibit the
formation of synaptic-like vesicles at the plasma membrane of PC 12 cells
(Schmidt et al. 1999). Farsad and colleagues (2001) have recently shown that
purified endophilin can directly bind and evaginate lipid bilayers into narrow
tubules, with a diameter size similar to that seen in the neck of a clathrin-
coated bud. Both endophilin and amphiphysin have been shown to tubulate
lipid bilayers, independently as well as complexed with dynamin (Fardar et al.
2001). Membrane tubulation is one of endophilin’s many possible functions in
clathrin-mediated endocytosis. Future research is still needed, however, to
further characterize endophilin’s role, along with dynamin and amphiphysin, as
part of a protein complex governing various aspects of membrane deformation
and fission.
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Phosphoinositides and synaptojanin: Phosphoinositide phosphatases (PI),
through their PH domains, act as docking sites for signaling molecules, and
have been implicated as regulators of endocytosis. Membrane-bound
phospholipids can be phosphorylated (by kinases) and dephosphorylated (by
phosphatases) at various sites. Sequential phosphorylation of a specific
hydroxyl group within an inositol ring produces phosphatidylinositol 4-
phosphate (PIP) and phosphatidylinositol 4,5-phosphate (PIP2), important in
the generation of two second messengers by phospholipase C activation:
inositol 1,4,5-triphosphate (IP3) and diacylglycerol (DAG). Both are vital in
various cellular functions as IP3 affects cytosolic calcium concentrations by
mobilizing calcium from the endoplasmatic reticulum and DAG activates
protein kinase C (reviewed in Lodish, et al. 2000). PIP2 has been shown to play
a crucial role in various steps of endocytosis, such as synaptic vesicle docking
and fusion with the plasma membrane, synaptic vesicle recycling, recruitment
of synaptic vesicle proteins into clathrin-coated pits as well as the pinching off
of clathrin-coated vesicles (Cremona et al. 1999). Additionally, PIP2 has been
shown to facilitate adaptor binding to clathrin-coated pits (Gaidarov et al.
1999), as well as the recruitment of coats to membranes (Czech 2000; Toker,
1998) and regulation of the localization and GTPase activity of dynamin by
binding to its PH domain (Zheng et al., 1996; Vallis et al., 1999). Conversion
of PIP into PIP2 has been implicated in exocytosis and endocytosis while PIP2
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83
conversion to PIP is required for clathrin-coat shedding and ultimately,
synaptic vesicle recycling (reviewed in Stenmark, 2000). Once the synaptic
vesicle is primed for fusion, rises in cytosolic calcium concentrations result in
vesicle fusion. Membrane recycling, following neurotransmitter release, is
facilitated by clathrin-mediated endocytosis. Once the clathrin-coated
endocytic vesicle is released from the plasma membrane, synaptojanin, a PI-5-
phosphatase, converts PIP2 into PIP, resulting in the release of adaptors from
the clathrin coat. The basis for the regulatory function of PIP and PIP2 is their
rapid turnover rate; consequently, phosphatases, such as synaptojanin, play a
vital role in controlling the various intracellular processes mediated by these
lipids. Data in support of a role for synaptojanin in vesicle uncoating was
demonstrated by Cremona and colleagues (1999) who showed that
synaptojanin knockout mice have elevated levels of PIP2 and show an
accumulation of clathrin-coated vesicles. These data support the hypothesis
that defects in synaptic transmission seen in synaptojanin-deficient mice may
be the result of inhibition of uncoating of clathrin-coated vesicles, which
consequently cannot fuse with the endosome, blocking synaptic vesicle
recycling (Stenmark, 2002).
Additionally, synaptojanin and PIP2 have been implicated in endocytosis by
regulation of actin-dependent dynamics as well as actin rearrangement in
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84
mammalian cells via its binding to a variety of proteins that control actin
dynamics (Sakisaka et al. 1999; Sechi and Wehland, 2000). PIP2 binds to and
suppresses the function of various actin-regulating proteins, such as profilin,
cofilin, gelsolin, and gCap (Lassing and Lindberg, 1985; Janmey and Stossel,
1987; Yu et al. 1990; Yonezawa et al. 1991) and has been implicated in actin
polymerization as well as actin depolymerization events (Sakisaka et al. 1997).
One example of PIP2 function is its binding to profilin: data suggest that PIP2
binds to and sequesters profilin in resting cells, inhibiting its mediation of actin
polymerization. This process is reversed by activation of PLC signaling
cascades, which releases PIP2 and allows profilin-complexes to engage in actin
polymerization (Sohn et al. 1995). Others, however, have suggested other
mechanisms for PIP2 regulation of profilin (Goldschmidt-Clermont et al.
1995). It is hypothesized that synaptojanin binding to actin MFs may facilitate
hydrolysis of PIP2 bound to cytoskeletal components (Sakisaka et al. 1997).
Additionally, novel interactions between inositols and actin regulatory
proteins, such as syndapins, have been described (Hilton et al. 2001). Although
the precise role of actin in endocytosis is still being characterized, the immense
variety of signaling cascades elicited by phosphoinositides and their
association with actin regulatory proteins may represent an additional
mechanism of control of endocytic function and membrane trafficking.
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Syndapin: Syndapin I (synaptic dynamin-associated protein, SdpI) was
originally characterized as a brain-specific protein, enriched in nerve terminals
and hypothesized to act as a molecular link between the actin cytoskeleton and
proteins known to facilitate clathrin-mediated endocytosis, such as dynamin
(Qualmann et al. 1999). Syndapin II (SdpII), was later found to be
ubiquitously expressed and is also thought to coordinate interactions between
components of the endocytic machinery and MFs (Qualmann and Kelly, 2000).
SdpI is a 54 kD protein (Figure 4.2.), with an SH3 C-terminus domain, and
NPF domain, common to all syndapin family proteins. Syndapin function has
been implicated in regulation of actin dynamics (Qualmann and Kelly, 2000)
as well as attachment of the actin cytoskeleton to the membrane (Merilainen et
al. 1997; Qualmann et al. 1999; Ritter et al. 2000) and to cortactin (McNiven et
al. 2000). Since syndapin’s discovery in the late 90’s, it has been related to
other syndapin-like proteins, such as FAP52 (a focal adhesion-associated
phosphoprotein), adding to a list of what is now rapidly becoming the
FRAP52/PACSIN/syndapin family of proteins (Merilainen et al. 1997; Nikki
et al. 2002). In vitro binding studies have demonstrated that the SH3 domain of
syndapin is able to bind to a number of proteins including synaptojanin,
synapsin Ia/Ib and N-WASP, an important regulator of actin cytoskeleton
organization (Qualmann et al. 1999; Modregger et al. 2000). Functional
association of syndapin in endocytosis and its association with components of
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86
Syndapin I
oSltelical s in
N- |-C
Coiled Coils 382 441
Q ualm ann et al. M ol. Biol. C ell. 10 501 513, I9 ‘W.
Syndapin 1 1
N -
-C
Coiled Coils 41.9 4:88
Qualmann and Kelly, J. Cell Biol. 148:10:47-10.6.1, 2000.
Figure 4.2. Syndapin domain structure: Members of
the syndapin family share similar domains structures,
with a highly conserved carboxy-terminus SH3 domain,
several NPF motifs and two predicted coiled-coil
domains.
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87
the endocytic machinery was demonstrated by Simpson and colleagues (1999);
3T3-L1 cells were utilized in the study of biotinylated transferrin endocytosis
and formation of clathrin-coated pits and vesicles. Data suggested that
introduction of SH3 domains of endophilin, intersectin, amphiphysin or
syndapin inhibited endocytosis and the formation of clathrin-coated vesicles.
Although evidence strongly suggested binding of SH3 domains to dynamin as
the likely endocytic partner, association with other proteins which facilitate
clathrin-mediated endocytosis was not ruled out, given that most SH3 domain-
containing proteins have multiple protein-interaction domains. A role for
syndapin in actin reorganization through the N-WASP signaling pathway has
also been suggested. Cortical actin reorganization into filopodia was seen in
cells expressing full-length syndapin - this effect was suppressed by
expression of a dominant negative N-WASP C-terminal fragment (Qualmann
and Kelly, 2000). Given the diversity of syndapin SH3 domain binding
partners, it is hypothesized that selectivity of syndapin action may be achieved
by concomitant binding to more than one component of the endocytic
machinery, forming a multi-protein scaffold, so to speak, to mediate/regulate
endocytic events. Additionally, it has been suggested that different pools of
syndapin could associate with different protein complexes (reviewed in
Lanzetti et al. 2001). Finally, syndapin has also been implicated in Ras and
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88
Rac signaling pathways, small GTPases known to regulate cytoskeleton
organization (Wasiak et al. 2001).
Actin binding protein 1 (Abpl): Abplp is homologous to drebrins and forms a
family of proteins, conserved from yeast to humans (reviewed in Lappalainen
et al. 1998). Originally identified in yeast (Drubin et al. 1988), it contains
multiple functional domains: an ADF/cofilin homology domain (ADF-H), a
helical region, a proline-rich domain and an SH3 domain (Figure 4.3.). Both
Abplp and its mammalian homologue, Abpl, associate with dynamic actin
structures in lamellipodia and filopodia and have consequently been implicated
in regulation o f actin dynamics. Drubin et al. (1988) demonstrated Abpl
involvement in actin cytoskeleton organization, as its overexpression resulted
in a depolarized distribution of cortical actin patches, as well as loss of
regulation of cell growth. Studies of Cos-7 cells have demonstrated the
association between Abpl, Arp2/3 and the GTPase Rac. Rac activation
resulted in the translocation of Abpl from a perinuclear region to the leading
edge of the cells, where it colocalized with Arp2/3 (Kessels et al. 2000).
Additionally, a variety of studies in yeast and mammalian cells have linked
Abpl to endocytosis functionally (reviewed in Qualmann et al., 2000), a
hypothesis strengthened by the demonstration of Abpl binding to proteins
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89
Actin Binding Protein 1 (Abpl)
A D F-H Helical Flexible SH3
Kessesls et al, JC B , 15:351-366, 2001
Figure 4.3. A bpl domain structure. Abpl can bind to
F-actin via N-terminal modules and to proline-rich
domains of proteins implicated in endocytosis via its
SH3 domain.
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90
which facilitate endocytosis, such as dynamin (present study) and
amphiphysin (Goode et al. 2001). A bpl’s ability to bind to dynamin, via its
SH3 domain, and to F-actin, via two distinct N-terminal regions (Kessels et al.
2000), has reinforced the hypothesis that it functions as a bridge, linking actin
MFs to components of the endocytic machinery. GST-pulldown assays
utilizing Abpl-SH3 domains fused with glutathione-S-transferase (GST) have
also recently demonstrated Abpl binding to N-WASP (present study),
implicating it in regulation of actin dynamics through the N-WASP-Arp2/3
signaling pathways (Goode et al. 2001).
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91
Results:
SdpI, SdplI and A bpl: novel components o f the endocytic machinery in
lacrimal acini. The accumulation of coated pits in CD-treated acini seen by
electron microscopy (Figure 3.9.) suggested to us that MFs may facilitate,
together with dynamin, the pinching-off of coated-pits in the formation of
coated vesicles, leading us to investigate possible mediators between them.
Recently, various novel src homology 3 (SH3) domain-containing proteins,
identified in neurons, have been shown to interact with dynamin’s proline rich
domain (PRD) and have been suggested to function as effectors of endocytosis.
SdpI and SdplI have been suggested to function as links between dynamin and
proteins involved in actin dynamics regulation, (such as N-WASP via the
Arp2/3 complex, see Introduction). Similarly, Abpl has been hypothesized to
function as a bridge, binding to dynamin (through its C-terminal SH3 domain)
and to actin MFs (through its N-terminal domain). SDS-PAGE of lacrimal
gland acinar cell lysate and Western blotting utilizing polyclonal antibodies to
SdpI (52 kD), SdplI (56 kD) (Figure 4.4A.) and Abpl (56kD) (Figure 4.4B.)
revealed that all three proteins are expressed in rabbit lacrimal acinar cells.
Non-specific binding of the goat-anti-rabbit secondary antibody is seen at
approximately 50 kD (syndapin Western blot) and at 60kD and 97kD in
Western blots developed for Abpl. We further characterized the distribution of
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92
A.
9 7 k D
66kt)
46kD
SdpI SdplI 2ary
97kD -
Al
66kD -
m m
56kD -
45kD -
ait
,>(M) -
A hp 1 2ary
Rabbit Lacrim al O larN
A bpl
M ouse
Figure 4.4. Western blot. SDS-PAGE of lacrimal gland acinar cell lysate
and subsequent Western blot analysis revealed that Spdl (52kD), SdplI
(56kD) and Abpl (56 kD) are expressed by these cells. Non-specific
binding of the goat-anti rabbit secondary antibody is seen at
approximately 50 kD (syndapin Western blot, Figure 4.4A.) and at
approximately 60kD and 97kD in the Abpl Western (Figure 4.4B.),
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93
B .
Rabbit 2ary
Resting
FTTC Actin
Actin MFs
Stimulated
FITC Actin
' ' W
Figure 4.5. SdpI, SdplI and Abpl distribution in rabbit lacrimal acini.
Lacrimal acini were cultured and processed for confocal microscopy and
immunofluorescence staining with antibodies against SdpI, SdplI and Abpl
(see Methods). Incubation with rabbit 2ary antibody alone showed no
fluorescence signal as seen by confocal microscopy (Figure 4.5A.). A
punctate distribution throughout the cell of SdpI, SdplI and Abpl is seen in
Figure 4.5B. CCH stimulation (lOOpM, 5 min) had no effects on protein
distribution. Bar = 10pm.
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94
SdpI, SdplI and Abpl by confocal immunofluorescence microscopy (Figure
4.5.). Immunostaining with rabbit-FITC conjugated secondary antibody alone
shows no fluorescence signal (Figure 4.5 A.). Staining of MFs with rhodamine-
phalloidin (red) was used to located apical lumens (*). Punctate staining of
SdpI, SdplI and Abpl is seen throughout the cell, including traces at the APM
(Figure 4.5B.), Contrary to what was seen with coat protein distribution,
stimulation with CCH (100 pM, 5 min) did not result in marked redistribution
of SdpI, SdplI or Abpl to the APM region.
Fusion protein SH3 domain interaction with components o f the endocytic
machinery: dynamin, N-WASP and synaptojanin. Our interest was to test,
specifically in lacrimal acini, whether the SH3-domains of SdpI, SdplI and
Abpl associated with proteins known to facilitate clathrin-mediated
endocytosis. One of the known effectors of this endocytic process is the
GTPase dynamin, known to facilitate the pinching off of coated pits. SdpI and
SdplI have been shown to associate with dynamin in neurons (Qualmann and
Kelly, 2000; Qualmann et al. 1999). To address this question, glutathione-S-
transferase (GST) fusion proteins were constructed (see Methods) with the
SH3 domains of SdpI (SdpI-SH3, residues 376-441), SdplI (SdpII-SH3,
residues 419-488) and Abpl (Abpl-SH3, residues 282-433), (Figure 4.6.),
Additionally, an SdpI-SH3 mutant construct was made in which a point
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95
GST-SH3
GST
P434L — Syndapin I (mut)
SH3
Figure 4.6. Domain structures o f SdpI, SdplI and A bpl
fusion proteins. The active SH3 domains of SdpI, SdplI and
Abpl were fused to glutathione-S-transferase (GST) and
subsequently introduced into lacrimal acinar cells. A GST-
Sdpl(mut) fusion protein, containing a point mutation in the
SH3 domain (leucine for proline substitution at amino acid
434) was utilized as a control, as the point mutation results in
an inactive SH3 domain.
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96
V 's ’ # i f # # /
ISOkD
BBIl
Mill
iiiliiiB iiiiP '
Synaptojanin
lOOkl)
Dynamin
66kD
n - : ! n V:ii:;y : py
N-WASP
30kD
GST
Figure 4.7. GST Pulldown Assay: lacrimal gland lysate protein-
specific interactions with SH3 domains o f fusion proteins.. SH3
domain fusion proteins of SdpI, SdplI and Abpl were incubated with
glutathione beads, rinsed, incubated with lacrimal gland acinar cell
lysate then rinsed extensively. Bound protein were eluted from beads
with sample buffer then separated by SDS-PAGE. Western blotting
revealed the direct interaction between the SH3 domains of SdpI, SdplI
and Abpl with dynamin, synaptojanin and N-WASP.
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97
mutation in its SH3 domain (substitution of a leucine residue for a proline at
amino acid 434) caused it to become inactive. Fusion proteins were
subsequently expressed in E. coli and purified. A GST-pulldown assay was
then utilized to characterize the purified fusion protein SH3 domain-specific
interactions with proteins in lacrimal gland lysate. Fusion proteins were
incubated with glutathione beads, rinsed to remove excess unbound protein and
subsequently incubated with lacrimal gland acinar cell lysate. After overnight
incubation and extensive rinsing, proteins bound to beads via SH3 domain
interaction were then eluted with sample buffer and separated by SDS-PAGE
(see Methods). Immunoblotting of membranes revealed that the SH3 domain
of SdpI and SdplI, as well as Abpl, associated with dynamin (Figure 4.7,).
Interactions were specific for the SH3 domain, as no bands were seen with
lysate exposed to the mutated SdpI-SH3 domain (Sdpl(mut)) nor with negative
controls (beads or GST alone). Further investigation of associated proteins in
the GST Pulldown assay revealed that the SH3 domains of SdpI, SdplI and
Abpl also bound two other proteins, synaptojanin and N-WASP, both of
which have also been implicated as facilitators of clathrin-mediated
endocytosis. As in the case of dynamin, association was SH3-domain specific
as no synaptojanin or N-WASP signal was recovered associated with the SdpI
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98
mutant or negative controls. Both synaptojanin and N-WASP were confirmed
to be expressed in lacrimal acini (Figure 4.7., lacrimal gland lysate control).
Given the association of fusion protein SH3 domain with synaptojanin and N-
WASP, confocal immunofluorescence microscopy was utilized to investigate
the distribution, in resting and CCH-stimulated (100 pM, 5 min) lacrimal acini,
of both proteins (Figure 4.8.), Rhodamine-phalloidin staining of MFs was
used to identify apical lumen (*). The punctate distribution throughout the cell
of both synaptojanin and N-WASP was similar to that seen previously with
SdpI, SdplI and Abpl (Figure 4.5B.) Secretagogue stimulation did not result
in apical recruitment of N-WASP and showed a slight apical recruitment of
synaptojanin (Figure 4.8.),
Introduction o f SdpI, SdplI and A bpl SH3-domains results in accumulation
o f coat-proteins and clathrin-coated pits at the apical plasma membrane.
Biochemical confirmation of the direct interaction between fusion protein SH3
domains and components of the clathrin-mediated endocytic machinery
(dynamin, synaptojanin and N-WASP), led us to investigate the effects
resulting from introduction of SdpI, SdplI and Abpl SH3 domains on the
distribution of clathrin and clathrin-associated proteins a-adaptin and dynamin.
Lacrimal acini were cultured for three days, electroporated in the presence of
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99
Resting Stimulated
FITC Actin FITC Actin
Figure 4.8. Distribution o f synaptojanin and N-WASP in rabbit
lacrimal acini. The distribution of synaptojanin and N-WASP was
investigated by confocal microscopy (see Methods). Rhodamin-
phalloidin staining of MF indicated location of apical lumen (*).
CCH stimulation had no effect on the punctate distribution of neither
synaptojanin nor N-WASP. Bar « 10 |um.
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100
fusion proteins and allowed to recover (3 hours, 37°C) prior to processing for
confocal immunofluorescence microscopy (see Methods). Previous work in
our lab had confirmed no differences were found, by confocal and electron
microscopy, between electroporated and non-electroporated acini.
Additionally, cell viability of electroporated samples was similar to that of
their non-electroporated counterparts. Parallel electroporation with (3-
galactosidase and subsequent colorimetric development with X-gal confirmed
an electroporation efficiency of over 90% in all assays, with no major losses in
cell viability. (Figure 4.9A.), Successful introduction of GST-fusion proteins
was also confirmed by SDS-PAGE of lysate from electroporated acini and
subsequent immunoblotting of membranes utilizing an antibody against GST
(Figure 4.9B.), Confocal fluorescence microscopy studies revealed that,
compared to control electroporated acini, introduction of SH3 domains of
Sdpf, SdplI and Abpl resulted in major apical enrichment of a-adaptin, similar
to that seen with CD treatment (Figure 4.10., *, apical lumen). No coat-
protein accumulation was seen in acini electroporated with the SdpI mutant or
with the negative control, GST alone. Similar results in apical protein
accumulation were seen with clathrin and dynamin (data not shown).
Additionally, confocal immunofluorescence microscopy analysis suggested
that introduction of SH3 domains had additional effects on the MF network
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101
3 0 k D
Con GST SdpI (linn ) SdpI SdplI Abpl
Figure 4.9. Introduction o f fusion proteins into lacrimal acini by
electroporation. Lacrimal acini were culture for thee days then
processed for electroporation (see Methods). Parallel electroporation
with (3-galactosidase and subsequent colorimetric development with X-
gal confirmed an electroporation efficiency of over 90% in all assays
(Figure 4.9A.) Successful introduction of GST fusion proteins was
confirmed by SDS-PAGE of electroporated acini lysate and subsequent
Western blotting utilizing an antibody against GST (Figure 4.9B.)
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102
a-adaptin a-adaptin
Figure 4.10. Introduction o f SdpI, SdplI and Abpl SH3 domain
results in apical accumulation o f coat-proteins. SH3 domains of SdpI,
SdplI and Abpl were introduced by electroporation in cultured
lacrimal acini, which were then processed for confocal
immunofluorescence and staining for a-adaptin and MFs (see
Methods). Introduction of SdpI, SdplI and Abpl SH3 domains resulted
in the accumulation of a-adaptin at apical lumen (*), as well as actin
MF bundling (arrows). No effects on coat protein accumulation were
seen with the introduction of the mutated SdpI SH3 domain or GST
alone. Bar « 10 pm.
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103
structure, with increased F-actin staining and MF bundling (Figure 4.10.,
arrows). To further elucidate the effects of the introduction of SdpI, SdplI and
Abpl SH3 domains on apical coat protein accumulation, we utilized electron
microscopy to investigate the effects of introduction of SH3 domains on the
distribution of coated pits and coated vesicles in electroporated acini. Electron
microscopy studies confirmed previous confocal data in that no differences
were seen between non-electroporated (Figure 3.7A.) and control
electroporated acini (Figure 4.11A.). Few microvilli were detected, extending
into the apical lumen (L). The box insert is enlarged (Figure 4.1 IB.) to better
visualize the electron-dense apically located clathrin-coated pits and vesicles
(arrows). No differences in clathrin-coated pits or coated vesicle distribution
were seen in acini in which the fusion protein containing the mutated SH3
domain of SdpI was introduced (Figure 4.11C.) Similar to controls, sparse
microvilli extend into the lumen and occasional clathrin-coated pits and
vesicles are seen at the APM (arrows, insert enlarged, Figure 4.1 ID.)
Introduction of SH3 domains of SdpI (Figures 4.12A-C) and SdplI (Figures
4.13A-C), however, resulted in a dramatic increase in the number of electron
dense coated pits at the APM (Figure 4.12B. arrows), accompanied by a
decrease in the number of clathrin-coated vesicles. These results were similar
to that seen in non-electroporated acini treated with CD (Figure 3.8B.),
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Confocal images had suggested that introduction of SdpI and SdplI domains
resulted in MF bundling (Figure 4.12A. and C, arrowhead). These data were
confirmed by electron microscopy, which showed thick actin MF bundles in
both SdpI (arrowheads, Figure 4.12A and 4.I2B.) and SdplI electroporated
acini (data not shown). Additionally, higher magnification images suggested
that microvilli density at the apical lumen (L) is increased in SdpI (Figure
4.12C.) and SdplI (Figure 4.13C.) electroporated acini, compared to control
acini and those electroporated with the mutated SdpI SH3 domain (Figure
4.11.), Similar results were seen in acini electroporated with the SH3 domain
of Abpl (Figure 4.14.), Higher magnification of the APM indicated that no
differences in coated pit distribution were seen with acini electroporated with
GST alone (Figure 4.14A.) with occasional coated pits (arrow) associated with
the apical lumenal membrane. Similar to that seen with acini electroporated
with the SH3 domain of SdpI and SdplI, introduction of the SH3 Abpl domain
resulted in a dramatic increase of apically located coated pits, (arrows).
Additionally, electroporation with the Abpl-SH3 domain also resulted in an
increased microvillar density (Figure 4.14B.) compared to the GST
electroporated acini.
Introduction o f SH3 domains o f SdpI, SdplI and A bpl result in an increase
in F-actin. Confocal and electron microscopy data suggested that introduction
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105
§ V ^ \ ? * %<:
III A y
wmft
Figure 4.11. Electron microscopy o f control electroporated acini. The
distribution of apically located clathrin-coated pits and vesicles in
lacrimal acini are seen by electron microscopy (Figure 4.11 A.).
Microvilli are seen extending into the apical lumen (L) from the APM.
Arrows indicate coated pits and/or coated vesicles. Insert (box) is
enlarged (Figure 4.1 IB.) to more clearly see coated pits and vesicles
(arrows). Electron microscopy analysis showed no differences from
controls in the distribution of clathrin coated pits and vesicles in acini
electroporated with the mutated SdpI SH3 domain (Figure 4.11C.),
with occasional coated pits and coated vesicles seen associated with the
APM (arrows, insert enlarged, Figure 4.1 ID.).
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106
Figure 4.12. Apical coated pit accumulation resulting from
introduction o f SdpI SH3 domain. Introduction, by
electroporation, of the SH3 domain of SdpI resulted in the
accumulation of coated pits (arrows, Figures 4.12B..) at the APM
of lacrimal acini, as seen by electron microscopy. Actin MF
bundling (arrows, Figure 4.12A.) and an apparent increase in
microvilli density at the apical lumen (L, Figures 4.12C) were also
noted.
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107
Figure 4.13. Introduction o f SdplI SH3 domain in lacrimal acini.
Results similar to those seen in SdpI electroporated acini of apical coat
protein accumulation (arrows) were seen by electron microscopy in acini
exposed to SH3 domains of SdplI.
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108
Figure 4.14. Effects o f Abpl SH3 domain introduction on clathrin-
coated pits and the MF network. Electroporation of acini in the
presence of Abpl SH3 domain resulted in accumulation of coated
pits (arrows) at the APM compared to control electroporation with
GST alone (Figure 4.14A.), Additionally, increased density of apical
microvilli in Abpl electroporated acini is identified by electron
microscopy, compared to GST control (Figure 4.14B.),
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109
of SH3 domains of SdpI, SdplI and Abpl resulted in increased F-actin and MF
bundling, effects not seen with control electroporated acini or with negative
controls, GST alone or the mutated SH3 domain of SdpI. In order to quantitate
the effects of SH3 domain introduction on the MF network, we utilized
Metamorph Imaging Software to measure F-actin fluorescence intensity in
electroporated acini. Lacrimal gland acini were cultured for three days then
processed for immunofluorescence and staining of F-actin with rhodamine-
phalloidin (see Methods). The fluorescence intensity of F-actin in z-section
images of lacrimal acini (10 images per acinus, four acini and 160 images total
per treatment, n = 4 independent assays) was quantitated and normalized to the
area of the acinus in each cross-section image (white outline, Figure 4.15A.),
Initial gain and contrast levels were established using a control and treated or
electroporated sample to ensure that fluorescence levels were not saturated.
Selected acini were scanned to determine the z-scan range. Nine to ten z-plane
sections, from the bottom to the top of the acinus, were acquired (thickness of
each section plane, 0.05 pm). The normalized fluorescence intensity of z-
sections in each acinus was summed, resulting in a final intensity value per
acinus (see Methods). Introduction of SH3 domains of SdpI, SdplI and Abpl
resulted in a significant increase in F-actin as quantitated by Metamorph
analysis of confocal images (*, p<0.05, Figure 4.15B.) confirming the results
suggested by confocal and electron microscopy.
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110
Changes in the MF network resulting from introduction o f SdpI, SdplI and
Abpl SH3 domains are not mediated via inhibition o f endocytosis. The
introduction of SdpI, SdplI and Abpl SH3 domains resulted in dramatic effects
on the MF network. These data led us to question whether the effects of
increased F-actin and MF bundling in electroporated acini could have been the
result of inhibition, by SH3 domains, of endocytosis. Previous FACs analysis
and data (Figure 3.10.) had suggested that MpCD successfully inhibited
endocytosis in lacrimal acini. In order to investigate this possibility, confocal
microscopy and Metamorph z-sectioning quantitation of F-actin was utilized to
visualize whether treatment of acini with MpCD resulted in changes in the MF
network. Lacrimal gland acinar cells were treated with MPCD (1 mM, 30 min)
then processed for confocal immunofluorescence and staining of F-actin with
rhodamine-phalloidin (see Methods). No major changes in the MF network
were seen, by confocal microscopy, in MpCD-treated acini. Z-section images
(four acini per assay, n=3 independent assays, 120 images total per treatment)
were acquired and F-actin staining intensity was quantitated as above (Figure
4.16A.), Metamorph quantitation analysis confirmed confocal microscopy data
in that no significant increases in F-actin resulted from treatment with MpCD
(Figure 4.16B.), These data suggested to us that the introduction of the SH3
domains of SdpI, SdplI and Abpl resulted in changes in the MF network by
mechanisms other than inhibition of endocytosis.
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I l l
Control SdpS(mut)
esr siipum ut) sap)
Figure 4.15. Increased F-actin resulting from the introduction o f SH3
domains. SH3 domains of SdpI, SdplI and Abpl were introduced into
lacrimal acini by electroporation, then processed for confocal
immunofluorescence and staining of F-actin by rhodamine-phalloidin (see
Methods). The effects on F-actin resulting from the introduction of SH3
domains, as seen by confocal microscopy, were quantitated using
Metamorph Quantitation Software of F-actin fluorescence intensity (see
Methods, 10 images per acinus, four acini and 160 images total per
treatment, n = 4 independent assays). Introduction of the SH3 domain of
SdpI, SdplI and Abpl resulted in a significant increase in F-actin as
compared to control electroporated acini. No increases in F-actin were seen
with introduction of the mutated SdpI SH3 domain or with GST alone (*, p
< 0.05, n=4). Bar = 10p.m.
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112
MBCO MBCO+CCH
Figure 4.16. Inhibition o f endocytosis by Mf>CD does not increase F-actin in
lacrimal acini. Lacrimal acini were exposed to M(3CD (ImM, 30 min) then
processed for confocal microscopy and Metamorph F-actin fluorescence
intensity quantitation of confocal microscopy z-sections (see Methods, four
acini per assay, n=3 independent assays, 120 images total per treatment).
Treatment with MPCD did not result in MF bundeling (Figure 4.16A.) or in
an increase in F-actin (Figure 4.16B.) Bar s 10p.m.
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113
Discussion:
Regulation of actin dynamics is vital for many cellular functions and involves
a variety of accessory proteins. In the present chapter, we demonstrate a role
for SdpI, SdpII and Abpl in clathrin-mediated endocytosis in lacrimal gland
acinar cells. Additionally, our data implicate protein SH3 domains in actin MF
organization. Although SDS PAGE and Western blot analysis confirmed the
endogenous presence of SdpI and SdpII in lacrimal gland lysate (Figure 4.4.)
and suggest the presence of Abpl, additional work is in progress to better
resolve A bpl’s 56kD band.
Given the substantial evidence that exists implicating SH3 domains in
regulation o f endocytosis and actin dynamics (see Introduction), we were
interested in investigating SH3 domain function of SdpI, SdpII and Abpl in
endocytosis in lacrimal acini. In order to address this question, the SH3 domain
of SdpI, SdpII and Abpl were expressed in E. coli as fusion proteins with
glutathione-S-transferase (GST) (see Methods). A point mutation in the SH3
domain of SdpI (Sdpl(mut)) fused to GST, which has previously been shown
to abolish the binding of Sdpl-SH3 to dynamin (Qualmann et al., 1999), was
also utilized. GST-pull down assays of fusion proteins demonstrated SH3-
domains of SdpI, SdpII and Abpl binding to endogenous dynamin,
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114
synaptojanin and N-WASP in lacrimal gland acinar cell lysate (Figure 4.3.),
which is, to our knowledge, the first demonstration of SH3 association with
these endocytic proteins in an epithelial cell model. Although binding of SdpI,
SdpII and Abpl to dynamin and synaptojanin had been previously
demonstrated in other systems (Qualmann et al. 1999; Kessels et al. 2000), this
is, to our knowledge, the first demonstration of Abpl-SH3 domain association
with N-WASP. Our data, together with previous research implicating Arp2/3
in Abpl function (Kessels et al. 2000), suggests that Arp2/3 activation by
Abpl is mediated via an N-WASP signaling pathway. Additionally, SH3-
fusion protein domain association with dynamin and synaptojanin suggests that
various components of the endocytic pathway may function in synchronicity,
activating the diverse signaling and regulatory pathways that fully reflect the
complex and well-orchestrated endocytic event which is clathrin-mediated
endocytosis. Hypothetical models addressing the role of syndapins and Abpl
in facilitating endocytosis in lacrimal acini are explored in Chapter 6.
Confirmation of the interaction between fusion protein SH3 domains, dynamin,
synaptojanin and N-WASP led us to further investigate the possible role of
SH3 domains in clathrin-coated vesicle formation. To this end, GST-fusion
proteins were expressed in Escherichia coli and introduced into cultured
lacrimal gland acinar cells by electroporation (see Methods). Similar to what
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115
was seen in CD-treated acini (Figure 3.5.), confocal microscopy revealed that
introduction of the SH-domains of SdpI, SdpII and Abpl resulted in the apical
accumulation o f coat proteins (Figure 4.6.), Electron microscopy analysis of
electroporated samples further confirmed that SH3-domain introduction
resulted in a dramatic increase in the number of apically located coat-pits
(Figure 4.9), with few coated vesicles found at the subapical cytoplasm. No
effects were seen with introduction of the inactive SdpI SH3-domain mutant or
with GST alone. These data strongly suggested to us that clathrin-mediated
endocytosis in lacrimal acini is facilitated by syndapin and Abpl signaling
pathways and implicated them in regulating or facilitating the transition from
coated-pit to coated vesicle formation.
Previous research has implicated accessory proteins in the regulation of actin
MF dynamics. Abpl, with its F-actin binding modules, is suggested to directly
bridge MFs to components of the endocytic pathway. Syndapins have been
implicated in MF regulation through the N-WASP-Arp2/3 signaling pathway
(see Introduction). Much of the present research, however, suggests that
regulation of F-actin by syndapin and Abpl involves regulated activation and
binding of multiple protein domains that ultimately mediate the formation of
multiprotein complexes. In support of this working model, Qualmann and
Kelly (2000) demonstrated that overexpression of full-length syndapins, but
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not the N-terminal or SH3 domains alone, induced filopodia formation and had
strong effects on cortical actin organization. We were consequently surprised
to find that introduction of the SH3 domain of SdpI, SdpII and Abpl resulted
in dramatic effects on the MF array in lacrimal acini. Confocal and electron
microscopy analysis (Figure 4.7. and 4.9.) revealed intense MF bundling in
SH3-domain electroporated acini coupled with an apparent increase in F-actin.
Metamorph analysis and quantitation of confocal images (Figure 4.1.)
confirmed a significant increase in F-actin staining in acini electroporated with
SH3 domains of SdpI, SdpII and Abpl. Furthermore, no such increases or
reorganization in the MF array were seen with acini exposed to the mutated
SdpI SH3 domain. These results lead us to investigate whether the changes
seen in the MF cytoskeleton resulted from the inhibition of endocytosis in
electroporated acini. To address this question, acini were exposed to MfiCD a
membrane impermeable cyclic oligosaccharide shown to extract cholesterol
from the plasma membrane (see Chapter 3, Introduction). Inhibition of
endocytosis in MpCD-treated acini was confirmed by FACs analysis of FITC-
dextran uptake (Figure 4.11). Metamorph quantitation of confocal microscopy
images showed no effects on actin organization or increases in F-actin in
treated acini. We could not confirm whether SH3-domain introduction had an
inhibitory affect on endocytosis, as FITC-dextran uptake studies in
electroporated acini did not yield consistent results. We believe this was due to
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117
the excessive manipulations and digestions o f lacrimal acini required, within
too a short period of time, to perform all needed assays (various resuspensions,
electroporation, exposure to FITC-dextran and digestion of acini for FACs
analysis). Flowever, given the effects of MpCD on inhibition o f endocytosis,
coupled by the lack of effects of the MF network, seen in MpCD-treated acini,
our data suggested to us that changes in F-actin resulting from introduction of
SH3 domains of SdpI, SdpII and Abpl were achieved by mechanisms other
than inhibition of endocytosis and further implicated SH3-domain signaling
pathway activation in MF organization (see Chapter 6).
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Chapter 5 - MFs and exocytosis in lacrimal acinar cells.
Introduction
Intracellular vesicle trafficking is an essential component of endocytosis and
exocytosis. In the previous chapters, we investigated the role of MFs in
endocytosis (Chapter 3) as well as the role of accessory proteins as regulators
of actin dynamics and facilitators of clathrin-mediated endocytosis (Chapter
4). In the present chapter, we focus on exocytic pathways. Although each
chapter has focused on specifically the outward, or inward movement of
membranes, it is important to note that inhibition of endocytic traffic may
indirectly affect exocytic traffic, and vice versa. For example, inhibition of
endocytic apical membrane after secretion may reduce the available membrane
pool available for recycling to form new secretory vesicles, and consequently,
inhibit secretion.
Exocytosis can be constitutive or regulated. In the former, exocytosis occurs at
a steady rate, independent of extracellular signals. In the latter, secretory
proteins are stored in high concentrations at or close to their site of release -
receptor-mediated activation of signaling pathways mobilizes mechanisms that
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119
result in movement and fusion of secretory vesicles (SVs) with the plasma
membrane. This chapter investigates cytoskeletal involvement in regulated
exocytosis, focusing on whether MFs facilitate the outward movement of SVs
in response to secretagogue-stimulation. Additionally, the role of other known
modulators of exocytosis and membrane trafficking, such as rabs and VAMPs
will be reviewed.
Actin MFs and exocytosis: Over the past four decades, cytoskeletal
involvement in exocytosis and vesicular traffic has been studied in many cell
models, such as pancreatic cells (Jungerman et al. 1995; Rosado et al. 2002),
parietal (Okamoto et al. 2001) and lacrimal acinar cells (da Costa et al. 1998),
as well as in the release of neurotransmitters (Doussau and Augustine, 2000)
and hormones (Yoneda et al. 2000; Wilson et al. 2001). In the early 70’s, Lacy
and colleagues (1973) put forth a hypothesis in which the MF network at the
APM was considered as a barrier to secretion, restricting access of secretory
vesicles (SVs) to the plasma membrane. Secretagogue stimulation was
suggested to trigger a transient redistribution of apical MFs, allowing SVs to
move past the MF barrier and fuse with the plasma membrane (Lacy et al.,
1973). This model was supported by observations of MF fragmentation and/or
changes in MF organization that accompanied increased secretion (O’Konski
and Pandol, 1990: Perrin et al., 1992: Jungerman et al., 1995). Studies of
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120
chromaffin cells demonstrated that, in a resting state, chromaffin granules were
connected to the plasma membrane by filamentous structures. Upon
stimulation with nicotine, sites of granule fusion were found to be devoid of
these filaments, suggesting actin disassembly and reinforcing the theory that
cortical actin regulates access to the plasma membrane. (Cheek and Burgoyne,
1986; Burgoyne and Cheek, 1987). Others, however, have questioned whether
the primary role of the cortical actin network is to prevent secretory granule
docking or to regulate granule fusion and exocytosis (Burgoyne and Cheek,
1987; Vitale et al., 1991; Trifaro and Vitale, 1993; Trifaro et al. 2000).
Additionally, other roles have been suggested for MFs and exocytosis -
changes in the organization of the MF network and polymerization of cortical
actin have been implicated in propelling vesicles to their site of fusion
(Waskiewicz and Cooper, 1995; Kibble, Barnard and Burgoyne, 1996; Wacker
et al., 1997; Steyer and Aimers, 1999). MF function as “tracts” for myosin
motor proteins has also been suggested in a variety of membrane trafficking
events (reviewed in Tuxworth and Titus, 2000) as well as the idea that MFs,
through the action of the acto-myosin contractile system, could provide the
force necessary for the extrusion and release of secretory products (Lydowyke
et al. 1994; Segawa and Yamashina, 1989; Doussau and Augustine, 2000).
However, the precise mechanisms as to how MFs facilitate exocytosis have not
been completely resolved, nor are there consistent data demonstrating that
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apical MFs mediate exocytosis in all secretory systems. We, and others (da
Costa et al. 1989; present study; Matthews et al. 1997) have shown that
secretory responses are not inhibited in cells treated with the MF-targeted
agent cytochalasin D (CD). Additionally, there is much evidence linking the
microtubule (MT) filament network and cytoplasmic dynein, a MT-associated
motor protein, in facilitating exocytosis (reviewed in McNiven et al. 2000).
Previous work in our lab has shown that disruption of the MT-network by
nocodazole, or taxol-induced MT stabilization, blocked the release of the
secretory protein P-hexosaminidase into the culture medium of acini exposed
to secretagogue (da Costa et al. 1998). Although we have not previously
demonstrated strong effects of CD on stimulated secretion, the availability of
new probes as discussed below (Rab3D and Vamp2) as well as the possibility
that indirect effects associated with the inhibition of endocytosis might cause
subtle changes prompted a reevaluation of a role for MF in exocytosis. In the
present chapter, we demonstrate that neither the stimulated release of rab3D-
positive granules nor the apical recruitment of VAMP2-positive SVs is
inhibited by CD-disruption of the MF network. We also show that protein
release, in response to CCH stimulation, is not diminished neither by
disruption of the MF array by CD nor by the introduction of the SH3 domain
of the actin regulatory proteins SdpI, SdpII and Abpl (see Chapter 4).
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122
Small GTP-binding proteins: Small GTP-binding proteins (20-40 kD) were
first discovered about two decades ago as the v-Ha-Ras and v-Ki-Ras oncogens
of sarcoma viruses (Shih et al. 1978; Chien et al. 1979) and were later shown
to be related to heterotrimeric G proteins, such as Gs and Gi (Scolnick et al.
1979; Gibbs et al. 1984; Shih et al. 1980). What is today known as the small
GTPase superfamily of proteins consists of over 100 members, characterized
from yeast to humans, and is structurally classified into five subfamilies,
namely, ras, rho, rab, sarl/arf and ran families. Family members share
conserved domains with a 30-55% sequence homology (extensively reviewed
in Takai et al. 2001). They have been characterized as “molecular switches”
and as their name indicates, shuttle between an active GTP-bound state and an
inactive GDP-bound state (Bourne et al. 1990; Takai et al. 1992). Small GTP-
binding proteins are frequently activated by signals originating from membrane
receptors and propagate these signals to downstream effectors. They have been
implicated in the regulation of a truly immense variety of biological functions,
as varied as membrane trafficking, vesicle docking, cell proliferation,
apoptosis, cell adhesion, cytoskeletal organization and gene expression, among
many others (Takai et al. 2001). Rho family members are essential for
cytoskeletal organization and regulation of actin MF dynamics (reviewed in
Chapter 1, Introduction).
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123
Rabs: To date, over 60 mammalian rab proteins have been identified and like
other members of the small GTPase family of proteins, Rab function has been
implicated in an immense diversity of cellular functions (reviewed in Pfeffer,
2001). The cytosolic GDP-bound rab pool at steady state represents anywhere
between 10-50% of a given rab protein (Takai et al. 2001) and cycles between
an inactive cytosolic and an active GTP-membrane-bound state through GTP
hydrolysis, which is regulated by various GTP-regulatory proteins (GEFs -
guanine nucleotide exchange factors; GDIs - GDP dissociation inhibitors and
GAPs - GTPase-activating proteins; Novick and Zerial, 1997). A COOH-
terminal lipid modification of rab proteins has been shown to be essential for
correct rab membrane targeting, although its mechanism of action is still
unknown (Chavrier et al. 1991). A large body of evidence has accumulated
over the years demonstrating rab function in regulation of various stages of
membrane trafficking, including exocytosis (Haddad et al. 2001; Zhao et al.
2002; Valentijn et al. 2000a and 2000b), and more specifically, vesicle
targeting to the acceptor membrane, SV docking and fusion (Nuoffer and
Balch, 1994; Nocivk and Zerial, 1997; Schimmoller et al. 1998; Martinez and
Goud, 1998). Recent research implicated rabs in the recruitment of MT- and
MF-based motor proteins, known to facilitate the transport of vesicles from
their site of formation to their fusion site (Goud, 2002; Hammer and Wu,
2002). Additionally, rabs have been suggested to direct membrane trafficking
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124
by recruiting tethering factors onto transported membranes. Tethering
components, such as Usol (Cao et al. 1998), TRAPP (Sacher et al. 1998) and
exocyst (Guo et al. 1999), are elongated complexes that have been suggested to
act, prior to vesicle docking, interacting with proteins both on the vesicle and
on the docking site (Pfeffer, 2001). Different rab isoforms have been
associated with different intracellular trafficking events, for example: Rabl,
rab2 and rab6 have been localized to the endoplasmic reticulum and the Golgi
apparatus (Haubruck et al. 1989; Chavrier et al. 1990; Goud et al. 1990); rab3
is a marker of secretory granules and synaptic vesicles (Fischer et al. 1990;
Geppert et al. 1994; Valentijn et al. 2000a and 2000b; present study); rab 4 and
rab5 are localized to early endosomes and have been associated with mediating
early steps endocytosis as well as mediating receptor recycling and endosome
fusion events (Chavrier et al. 1990; Gorvel et al. 1991; van der Sluijs et al.
1992; Bucci et al. 1992).
In the present study, we use rab3D as a marker of mature secretory vesicles in
lacrimal gland acinar cells. Rab3D has been localized to dense core granules in
AtT-20 cells and its overexpression has been implicated in inhibition of
regulated secretion (Tabellini et al. 2001). Rab3D has also been associated
with secretory vesicles in pancreas, liver, parotid and lacrimal glands, among
others (Valentijn et al. 1999b and 2000b; Field et al. 2001; Piiper et al. 2001,
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125
present study). Recent studies demonstrating rab3D dissociation from SVs
prior to their fusion with the plasma membrane suggests a role for rab3D in
early steps of the exocytic pathway, such a docking or tethering of SVs
(Valentijn et al. 2000b).
SNAREs and VAMPs: Given the immense variety of membrane trafficking
events, cellular mechanisms that correctly and specifically target proteins to
intracellular compartments are critical for cell function and survival. SNARE
proteins form a family of membrane-associated proteins, with homologues
from yeast to humans which, together with others, such as tethering
complexes, are responsible for the processing and delivery of proteins and
lipids within the exocytic (secretory) and endocytic pathways (reviewed in Lin
and Scheller, 2000). SNAREs were named by Rothman and colleagues
(Rothman and Warren, 1994) in describing a family of proteins able to bind to
soluble factors (NSF, N-ethylmaleimide-sensitive fusion protein and its
membrane-attachment proteins, SNAPs), hence the name “SNARE”, or
“SNAP receptor”. SNARE subfamily components include VAMPs (vesicle
associated membrane proteins, also known as synaptobrevins), syntaxin and
SNAP-25 (synaptosomal-associated protein of 25 kD) and historically
speaking were only recently identified (Sollner et al. 1993; Rothman and
Warren, 1994). Family members are characterized as v-SNAREs (VAMPs) or
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126
t-SNAREs (syntaxin and SNAP-25), depending on their association with
vesicle membranes or target membranes, respectively (Trimble et al. 1988;
Oyler et al. 1989; Bennett et al. 1992). Studies in yeast showed that v- and t-
SNAREs cannot be on the same membrane vesicle for fusion to occur (Nichols
et al 1997). This was an important demonstration given that t-SNAREs,
although mostly found on the plasma membrane, have also been localized to
vesicle membranes (Walch-Solimena et al 1995). In the present study,
antibodies against VAMP2 are utilized for confocal immunofluorescence
microscopy studies.
Since their original identification in neuronal SVs, SNARE mediation of
membrane fusion events has been implicated in various cell types, including
pancreatic (Hansen et al. 1999), parotid and pineal cells (Gaisano et al. 1994;
Fujita-Yoshigaka et al. 1996; Redecker et al. 2000) and lacrimal and parietal
acinar cells (Lehnardt et al. 2000; present study). They play a role in the
regulation of secretagogue-stimulated (Fujita-Yoshigaka et al. 1996) as well as
constitutive secretion (Aalto et al. 1993; Protopopov et al. 1993). Crystal
structure analysis and functional studies have shown that the core SNARE
complex is composed of the syntaxin H3 domain, a VAMP coiled-coil domain,
a SNAP-25 amino-terminal and a SNAP-25 carboxyl terminal helix. Data to
date (reviewed in Lin and Scheller, 2000) suggest the following model of
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127
SNARE-mediated membrane fusion: syntaxin binding to VAMP or SNAP-25
releases syntaxin from a closed conformational state and results in the
formation of a ternary complex responsible for the initial event in membrane
fusion. Changes in cellular calcium concentrations result in further
conformational changes of the SNARE complex, bringing the vesicle and
plasma membrane into contact and driving the fusion reaction forward.
Disassembly of the complex is necessary, after exocytosis, before syntaxin,
SNAP-25 and VAMP can facilitate other fusion events (Lin and Scheller,
2000). Continued research results in an ever-growing SNARE-family of
proteins. Various accessory proteins have been implicated in SNARE
regulation, resulting in added extra levels of specificity in membrane fusion
events. For example, recent data suggest that rab proteins catalyze SNARE
complex assembly and are necessary for the docking of transport vesicles
(Sogaard et al. 1994). Although much has been revealed about SNARE
complex crystal structure and assembly, much still needs to be resolved, such
as how the core complex brings membranes together or how regulatory and
accessory proteins associate with SNAREs. Additionally, a role for SNAREs
as mediators of complex exocytic events, such as secretagogue-stimulated
secretion, must still be further elucidated.
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128
Results.
The stimulated release of Rab3D secretory granules is not inhibited by
disruption of the MF network. Confocal microscopy was utilized to investigate
the distribution of Rab3D, a marker of secretory granules, in resting and
stimulated acini, with and without CD pretreatment (see Methods). Rab3D, a
member of the rab family of small GTPases, has been implicated in exocytosis
(Fischer von Mollard et al., 1994) and is one of the principal rabs associated
with secretory vesicles in various epithelial cells, such as pancreas and lacrimal
gland acinar cells (Ohnishi et al., 1996; Valentijn et al., 1996; see
Introduction). Confocal microscopy indicated that the distribution o f rab3D in
resting acini was heavily concentrated in the subapical cytoplasm (*, apical
lumen), with very little punctate staining seen throughout the cell (Figure 5 .1).
Previous work in our lab suggested that the apical labeling of rab3D in resting
acini is associated with mature pre-formed secretory vesicles. Stimulation with
CCH (100 pM, 5 min) resulted in the dispersal of rab3D apical staining, with a
more intense and uniform labeling seen throughout the cell. No difference was
seen, compared to control, in the subapical localization of rab3D labeling in
CD-treated acini. Additionally, rab3D distribution in stimulated acini, pre
treated with CD, was similar to that in the CCH-stimulated acini, with rab3D
labeling dispersed throughout the cell. This data suggested to us that
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129
disruption of the MF network by CD does not inhibit the release of pre-formed
secretory granules elicited by CCH. To further quantitate the effects of CCH
stimulation associated with rab3D staining, we characterized the distribution of
rab3D, seen by confocal microscopy, as “apical”, “1/2 apical” or “diffuse”,
referring to the intense apical staining seen in control acini, a partial
redistribution or the dispersed staining associated with CCH-stimulation,
respectively (85 to 182 cells scored, n=4 separate assays). Quantitation
confirmed the significant ( *, P < 0 .05) reduction of apical rab3D staining
resulting from CCH stimulation (59% apical in control compared to 16% in
CCH-treated acini), which was accompanied by a significant increase in a
“diffuse” staining distribution pattern (9% apical in control compared to 61%
in CCH-treated acini, Figure 5.2.), The rab3D distribution in resting and
stimulated acini, pretreated with CD, was similar to untreated controls, with
“apical” distribution of 68% and 22% respectively. These data suggested that
CD disruption of the intact MF network did not impair the CCH-stimulated
release of rab3D-positive secretory granules. Preliminary studies o f rab3D
distribution in acini electroporated with the SH3 domain of SdpII and the
mutated SH3 domain of SdpI were also done to investigate the role of actin
regulatory proteins in secretory granule release. Electroporation in and of itself
did not affect the distribution of rab3D in resting or stimulated acini, which
was similar to the non-electroporated counterparts (Figure 5.1.), with intense
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130
Rab3l) Distribution
Figure 5.1. CD treatment did not inhibit the CCH-stimulated
apical release o f Rab3D-positive secretory granules. The
distribution of Rab3D in resting and CCH-stimulated (100|jM, 5
min) lacrimal acini was analyzed by confocal microscopy with
and without CD pre-treatment. Rab3D labeling in CD-treated
acini was similar to control, with intense Rab3D apical labeling.
The decrease in apical association of Rab3D resulting from CCH
stimulation was not inhibited by CD treatment. (*, apical lumen)
Bar = 1 0 pm.
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131
Rab3D-Positive Vesicle Distribution
80
45 50
L d ll
m
I apical
□ 1/2 apical
□ diffuse
control cch cd cd+cch
Figure 5.2. Quantitation o f Rab3D distribution. The distribution of
Rab3D was characterized, in confocal microscopy images, as “apical”,
“1/2 apical” or “diffuse”, in resting and CCH-stimulated acini (100 pM,
5 min), with and without CD pretreatment (5 pM, 60 min). CD pretreated
did not block the significant decrease in “apical” Rab3D staining
resulting from CCH-stimulation (*, p < 0.05, 85 to 182 cells scored, n=4
separate assays).
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132
rab3D staining associated with the apical cytoplasm in the electroporated
control and a dispersed distribution seen in stimulated electroporated acini
(data not shown). Introduction of the SH3 domain of SdpII or the mutated SdpI
SH3 domain did not alter the resting or stimulated rab3D distribution as seen
in Figure 5.3. These preliminary data suggested that the inhibition effects in
endocytosis described in Chapter 4 did not affect Rab3D-positive SVs.
CD treatment does not inhibit the apical recruitment o f VAMP2 in response
to CCH-stimulation. VAMP2 has been implicated as a facilitator of
secretagogue-stimulated exocytosis in various acinar cell models (Fujita-
Yoshigaka et al. 1996; Hansen et al., 1999, see Introduction). Additionally,
research in our lab (Wang et al, submitted) suggests that secretagogue
stimulation results in a redistribution of VAMP2 to the APM. To further
investigate whether there was a role for MFs in facilitating additional
components of the secretory machinery, such as VAMP2, in regulated
exocytosis, the effects of CD treatment on VAMP2 distribution were analyzed
by confocal microscopy. VAMP2 distribution in control unstimulated lacrimal
gland acini (Figure 5.4.) was punctate and dispersed throughout the cell, with
some basal levels of VAMP2 staining associated with the apical lumen (*).
The VAMP2 distribution in CCH stimulated acini (100 pM, 15 min) suggested
that secretagogue stimulation elicited a recruitment of VAMP2 to the apical
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133
cytoplasm, as seen by the increased labeling of VAMP2 associated with the
subapical cytoplasm (arrows). Treatment of acini with CD did not alter
VAMP2 distribution in unstimulated acini nor did it inhibit the secretagogue-
elicited apical recruitment, as seen by the increased apical
immunofluorescence staining of VAMP2 in CD-pretreated, stimulated acini
(arrows, Figure 5.4.),
Similar to our previous studies investigating whether SH3 domains played a
role in Rab3D distribution (Figure 5.3.), we were interested in verifying
whether the inhibition of apical endocytosis caused by the introduction of
SdpI, SdpII or Abpl altered the ability of acini to mobilize VAMP2 in
response to CCH-stimulation. The SH3 domains of SdpI, SdpII and Abpl were
introduced by electroporation into cultured lacrimal gland acinar cells which
were then processed for immunofluorescence staining and confocal analysis of
VAMP2 distribution (see Methods). VAMP2 distribution in control
electroporated acini, similar to the non-electroporated counterparts, was
punctate and dispersed throughout the cell (Figure 5.5., *, apical lumen).
Introduction of the SH3 domains of SdpI, SdpII and Abpl, as well as
introduction of negative controls (mutated SH3 domain of SdpI and GST
alone), did not alter VAMP2 distribution. CCH-stimulation of acini
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134
Rab3D
Resting Stimulated
Figure 5.3. Introduction o f the SH3 domain o f SdpII does not
affect Rab3D distribution in resting and secretagogue-stimulated
acini. Confocal microscopy of electroporated acini suggested
Rab3D distribution in acini electroporated with the SH3 domain of
SdpII or the mutated SH3 domain of SdpI is similar to that in
resting and CCH-stimulated non-electroporated controls (see
Figure 5.1.), (*, apical lumen) Bar = 10pm.
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135
electroporated with SH3 domains elicited a robust subapical recruitment of
VAMP2, as indicated by the increase in VAMP2 immunofluorescence
associated with the apical lumen (*) (Figure 5.6.), These data suggested to us
that the SH3 domains of SdpI, SdpII and Abpl do not impair apical
recruitment of VAMP2 elicited by CCH treatment through either direct or
indirect mechanisms.
The stimulated release o f /3-hexosaminidase is not inhibited by CD-
treatment Confocal microscopy data suggested that MFs did not facilitate the
release of rab3D-positive secretory granules in response to secretagogue
stimulation (Figure 5.1.) or the apical recruitment of VAMP2-positive
secretory granules induced by CCH (Figure 5.4.), Secretion assays, measuring
the release of protein into the culture media of lacrimal acini were done to
further confirm whether disruption of the MF network by CD would inhibit the
secretory response elicited by CCH treatment. P-hexosaminidase is enriched in
secretory membranes of lacrimal acini (Hamm-Alvarez et al., 1997) and
secreted in response to CCH stimulation (Gierow and Mircheff, 1998). The
release of P-hexosaminidase in resting and CCH-stimulated acini (100 pM, 30
min) acini with and without CD pre-treatment (10 pM, 60 min) was measured
in the culture media of lacrimal gland acini. P-hexosaminidase in cell-free
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136
Vamp2
Figure 5.4. An intact MF network is not required for the apical
recruitment o f VAMP2 following CCH-stimulation. Confocal
microscopy of immunofluorescence staining of VAMP2 in lacrimal
gland acini suggests a punctate distribution throughout the cell in
control acini. CCH stimulation (lOOpM, 15 min) results in apical
recruitment of VAMP2, as seen by the increased fluorescence intensity
associated with the apical lumen (arrows). Staining of VAMP2 in CD-
treated acini (5pM, 60 min) is similar to control. CD pre-treatment did
not block the CCH-elicited apical recruitment of VAMP2. (*, apical
lumen) Bar = 10pm.
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137
VAMP2 distribution in control electroporated acini
Figure 5.5. Electroporation ofSdpI, SdpII and Abpl SH3
domains does not alter VAMP2 distribution in lacrimal acini. The
SH3 domains ofSdpI, SdpII and Abpl were introduced by
electroporation into lacrimal acini, which were subsequently
processed for confocal microscopy (see Methods). Contrary to
what was seen with coat proteins (a-adaptin, clathrin and dynamin
(Figure 4.7.) introduction ofSdpI, SdpII and Abpl SH3 domains
did not alter the distribution of VAMP2 in lacrimal acini, which
was similar to controls (GST alone and the mutated SdpI SH3
domain). Bar s 10pm.
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138
media was determined using methyumbelliferyl-a-D-glucosaminide as
substrate (see Methods). Secretion assays revealed that pre-treatment with CD
did not have an inhibitory effect in secretion, as evidenced by the observation
of significant (*, p < 0.05) carbachol-stimulated release of 3-hexosaminidase
in acini with and without CD treatment (Figure 5.7A.), In addition to 3_
hexosaminidase, the total protein released into the culture media of treated
acini was also measured. These results paralleled those of 3-hexosaminidase as
CD pretreatment did not inhibit the secretagogue-stimulated response (Figure
5.7A.), We further quantitated the CCH-stimulated release of 3_
hexosaminidase into culture media of acini electroporated with the SH3
domains of SdpI, SdpII and Abpl, as well as the non-functional SdpI mutant
SH3 domain. Similar to what was seen in CD-treated acini, introduction of
SH3 domains did not inhibit the stimulated response elicited by CCH-
treatment (Figure 5.7B); a significant increase in the release of 3_
hexosaminidase, as well as total protein, was seen in all electroporated samples
stimulated (*, p < 0.05, n=6). Additionally, introduction of SH3 domains had
no effects on constitutive (unstimulated) protein release.
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VAMP2 distribution in CCH-stimulated
electroporated acini
WtplHimU
Figure 5.6. Apical recruitment o f VAMP2 in CCH-
stimulated electroporated acini. Introduction of the
SH3 domains of SdpI, SdpII or Abpl in lacrimal gland
acini (see Methods) did not inhibit the CCH-elicited
apical recruitment of VAMP2 as seen by the intense
fluorescence staining associated with the subapical
region, (arrows, *, apical lumen) Bar = 10pm.
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140
A.
(3 -Hexosaminidase Secretion
300
. 250
200
< 150
100
50
0
JL
■
Cont
■ Resting
□ Stimulated
I
X
CD
Protein Secretion
0.10 -
_ 0.08 -
.= *- 0.06 -
® 0.04
0.02 -
0.00
Cont CD
■ Resting
□ Stimulated
Figure 5.7A. CD-treatment nor introduction o f SH3 domains of
SdpI, SdpII and Abpl inhibits the release o f protein or (3-
hexosaminidase elicited by secretagogue stimulation. The release of
p-hexosaminidase and the release of total protein into the culture
medium of lacrimal gland acini was measured in resting and CCH-
stimulated (100 pM, 30 min) acini, with and without CD
pretreatment (5 pM, 60 min, see Methods). CD did not inhibit the
CCH stimulated release of protein release into the culture medium
(Figure 5.7A., p < 0.05, n=7).
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141
B.
(3-Hexosaminidase Secretion
■ Resting
□ Stimulated
250
200
-j 150
< 100
50
0
X
X. X
i\ ii ■ il il
T
Cont Sdpl(mut) SdpI SdpII Abpl
Protein Secretion
■ Resting
□ Stimulated
0.06
^ 0.04
O )
51.
0.02
0.00
X
I
I
■il il il il
X
X
■ I
Cont Sdpl(mut) SdpI SdpII Abpl
Figure 5.7B. Introduction ofSH3 domains o f SdpI, SdpII
and A bpl does not inhibit stimulated secretion in lacrimal
acini. (n= 6; *, p < 0.05). See Figure 5.7A.
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142
Recovery o f apical Rab3D-positive secretory vesicles is delayed by CD
treatment Secretion assay data suggested that CD treatment did not inhibit the
CCH-stimulated initial secretory response (30 min of secretagogue
stimulation) even though this treatment caused an apparent decrease in apical
clathrin-mediated endocytosis (Chapter 3). These data, however, did not
address the question of whether disruption of the MF network would affect the
recovery of rab3D-postitive granules to the APM following one round of
stimulated apical secretion and release of mature secretory vesicles. To address
this question, acini were stimulated with CCH (100 pM, 5 min, with and
without CD pretreatment at 5 pM, 60 min). The secretagogue was then
removed and acini were allowed to recover for a period of 15 and 30 min. CD
treatment was continued throughout the washout period in CD-pretreated acini.
Samples were then processed for confocal fluorescence microscopy for
immunostaining of rab3D. Confocal microscopy revealed that although acini
were able to recover the intense apical staining typical of rab3D, this recovery
was seen at 15 min post-washout in control acini as compared to 30 minutes in
the CD treated samples (Figure 5.8A., *, apical lumen). Quantitative scoring
of rab3D distribution in confocal images as “apical”, “1/2 apical” or
“dispersed” using the categories described in Figure 5.2. further confirmed the
delayed recovery effect in CD-treated acini (Figure 5.8B) as 65% of the
stimulated acini had recovered an “apical” staining by 15 minutes of washout
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143
Rab3D-positive Secretory Vesicle Recovery after CD Washout
(0
+ ■ >
o
I-
o
, o
■ apical
I 1 1/2 apical
□ diffuse
con+cch+w15 cd+cch+w15 cd+cch+w30
Figure 5.8. Recovery o f apical Rab3D-positive vesicles
followed stimulated release is delayed in CD-treated acini.
Rab3D recovery after CCH-stimulation was analyzed in
control and CD-treated acini. Confocal microscopy images
(Figure 5.8A.) suggested that CD treatment resulted in a
delayed recovery response. Quantitative scoring of Rab3D
distribution in confocal images as “apical”, “1/2 apical” or
“dispersed”, as described in Figure 5.2., further confirmed
the delayed recovery effect in CD-treated acini (Figure 5.8B,
*. n < 0.05, 85 to 182 cells scored, n=4 separate assavsV
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144
as compared to only 35% of the CD-pretreated stimulated acini (85 to 182 cells
scored, n=4 separate assays). An additional 15 minutes of washout was
necessary for the acini exposed to CD to completely recover (66% apical, 30
min washout period).
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145
Discussion
Proper cell functioning requires a coordinated effort between the outward and
the inward movement of secretory and endocytic vesicles. The cytoskeleton -
MTs as well as MFs - has been implicated as mediators of membrane
trafficking events. In Chapters 3 and 4, we demonstrated a role for MFs in
endocytic traffic in lacrimal acinar cells. Given the various models implicating
the actin MF network in exocytosis (see Introduction) we were interested in
investigating whether disruption of MFs with CD facilitated regulated
secretion in polarized epithelial cells. Additionally, we investigated whether
endocytic and exocytic events were tightly coupled in lacrimal acini.
Rab3D has been implicated as a facilitator of secretion and was used, in our
system, as a marker of mature SVs in lacrimal gland acinar cells. Confocal
microscopy revealed that in the resting state, rab3D-positive SVs accumulated
at the APM. Stimulation with secretagogue resulted in a redistribution of
rab3D, from an intense apical staining to a more punctate staining seen
throughout the cell (Figure 5.1.), Disruption of the MF network by CD had no
effects on rab3D redistribution, in the resting or stimulated state. Quantitation
of confocal images, characterizing rab3D immunostaining as “apical”, “ 1/2
apical” or “diffuse” confirmed that the CCH-stimulated release of rab3D-
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146
positive secretory granules did not require an intact MF array. Although our
data did not suggest that MFs played a role in the immediate release of rab3D-
positive SVs, exocytosis is a multi-step process, regulated by various accessory
proteins. MFs could potentially mediate other exocytic steps or serve as a
scaffold for other accessory proteins, such as VAMPs, which in lacrimal acini
are recruited to the APM as part of the secretory response (see Introduction).
Confocal microscopy was used to investigate the distribution of VAMP2 in
resting in stimulated acini in control cells and those treated with CD. VAMP2
distribution in resting acini was punctate and recovered throughout the cell.
Secretagogue-stimulation resulted in the apical recruitment of VAMP2
(Figure 5.4.), seen by the increase in the intensity of VAMP2 apical staining
and implicating VAMP2 as a mediator of apical exocytosis in lacrimal acini.
CD-pretreatment did not inhibit the apical recruitment of VAMP2 suggesting
that the MF array did not directly mediate the CCH-induced VAMP2
redistribution.
Previous data in our lab (Chapter 4) suggested that introduction of the SH3
domain of the actin regulatory proteins SdpI, SdpII and Abpl inhibited the
formation of clathrin-coated vesicles, demonstrating a role for these proteins in
endocytosis. Although no evidence of a direct role for MFs in regulated
exocytosis had been demonstrated in lacrimal acini, we were interested in
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147
verifying whether actin regulatory proteins could mediate exocytic membrane
trafficking events. To this end, preliminary studies were done, utilizing
electroporation to introduce the SH3 domain of SdpII into lacrimal acini. We
found that introduction of the SdpII SH3 domain did not alter the resting
distribution of rab3D, with intense staining still seen associated with the
subapical cytoplasm (Figure 5.3.), Introduction of the SdpII SH3 domain did
not inhibit the apical release of rab3D-labeled SVs following secretagogue
stimulation. Similar results were found in acini electroporated with the non
functional SH3 domain of SdpI. The data confirmed to us that in lacrimal acini,
an intact MF network is not essential for regulated secretion to occur, not did
we find a role for the actin regulatory protein SdpII in exocytosis.
Given these data, an important question still remained to be addressed, namely,
what was the relationship between clathrin-mediated endocytosis and
secretagogue-stimulated exocytosis in lacrimal acini. Did the inhibition of
endocytosis, resulting from CD-treatment (Chapter 3) or from the introduction
of SdpI or II or Abpl SH3 domains (Chapter 4) have a compensatory effect
diminishing secretion? To address this question, biochemical secretion assays
were done in CD-treated acini and those electroporated with the SH3 domains
of actin regulatory proteins. Measurements of basal and CCH-stimulated
protein and P-hexosaminidase secretion revealed that the apparent inhibition of
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148
clathrin-mediated endocytosis elicited by CD-treated as well as the SH3
domains of SdpI, Sdp2 and Abpl did not diminished the exocytic response
(Figure 5.7.), These data suggested to us that in lacrimal acinar cells, clathrin-
mediated endocytosis and CCH-stimulated exocytosis may not be tightly
coupled at least upon one round. Similar results were seen in evanescent wave
microscopy studies of VAMP and dynamin 1 distribution in PC-12 cells.
Tsubo and colleagues (2002) found that in simultaneous observations of GFP-
tagged dynamin and red fluorescent VAMP, over 70% VAMP plasma
membrane distribution in response to exocytic stimulus did not co-localize to
the same site as dynamin. They put forth a “kiss and glide” model of
exocytosis in which lateral mobility and fluidity of motion of molecules within
the plasma membrane is essential. In this model dynamin is recruited to the
plasma membrane in stimulated cells and subsequently forms dynamin
“clusters” which are free to move laterally within the plasma membrane. The
sweeping motion of dynamin clusters results in the collision with empty
vesicles, facilitating membrane retrieval.
We cannot, however, completely disregard a role for MFs in exocytosis in
lacrimal acini. Although secretion assays in CD-treated and electroporated
acini showed no inhibition of stimulated secretion, this secretory response
might reflect the release of pre-formed mature SVs, located at the APM. MFs
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149
could possibly facilitate the recovery of rab3D-positive following one round of
stimulated secretion. To address this question, control and CD-treated acini
were exposed to secretagogue and were then permitted to recover prior to
processing for fluorescence microscopy and immunostaining of rab3D. CD
treatment was continued throughout the washout period in CD-pretreated acini.
Quantitation of rab3D distribution suggested that after 15 minutes of wash-out,
control acini were able to completely recover the intense apical rab3D staining,
similar to that seen in control, unstimulated acini (Figure 5.8.), In CD-treated
acini, this recovery was only seen at 30 minutes of washout. This 15-minute
delay suggested to us that MFs may directly facilitate earlier steps of
exocytosis, for example, in the formation of SVs or their movement to the
APM. Additionally, the inhibition of endocytosis elicited by CD treatment may
indirectly affect SV formation - the block in plasma membrane internalization
in CD-treated acini might result in the decrease of available recycled
membrane used in the formation of new SVs. Also, availability of other agents
that alter actin MFs differently may suggest some role for MFs in exocytosis.
Preliminary research in our lab (Jerdeva, personal communication) suggests
that stabilization of actin filaments with phalloidin inhibits the secretagogue
response as measured by the release of protein and p-hexosaminidase (data not
shown).
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150
Chapter 6 - Endocytosis in Three Acts: A Summary
Clathrin-mediated endocytosis is but a simplified expression for what is, in
reality, a well-orchestrated, highly regulated endocytic event involving an
almost bewildering number of proteins and lipids that interact and activate one
another. The complexity is further enhanced by the fact that a single protein
can up-regulate or down-regulate a signaling pathway, depending on with
which effector it associates. The present chapter presents various models
describing the relationship between many of the components of the endocytic
pathway - some have been demonstrated (actin polymerization induced by N-
WASP activation of Arp2/3); others are hypothesized (a role for synaptojanin
as a negative-regulator of actin polymerization). The focus is on the MF
network and how actin dynamics drives endocytosis. Given the diverse
signaling mechanisms involved in clathrin-mediated endocytosis, we will
address one pathway at a time. Imagine, if you will, endocytosis as a play; the
stage is the lacrimal gland acinar cell and the players are the components of the
endocytic machinery (forgive me Shakespeare). Each “Act” will center around
protein activation of one possible signaling pathway, its effects on actin
dynamics and consequently, on endocytosis. Act I will focus on plasma
membrane-associated proteins and lipids; Act II describes models of actin
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151
polymerization induced by N-WASP-Arp2/3 activation; Act III exposes the
multiple actions elicited by dynamin activation.
As the curtain opens, exocytosis has just been elicited. SVs are fusing with the
apical plasma membrane and the excess SV membrane inserted into the APM
increases membrane tension, activating hitherto dormant signaling mechanisms
that trigger endocytosis.
A ct I, scene I: Tension mounts and the membrane invaginates.
One of the first steps in endocytosis is membrane invagination (see Chapter
3). Key players in this process are endophilin and amphiphysin, proteins that
have been shown to bind to various components of the clathrin-mediated
endocytic machinery. Their role in this play is still hypothetical, as they have
not, as yet, been characterized in lacrimal acini. However, given their
demonstrated importance in other systems (Simpson et al. 1999; Farsad et al.
2001), they are herein permitted a brief, albeit theoretical role (Figure 6.I.),
Endophilin is a lipid-modifying agent demonstrated to modulate membrane
tension and initiate the membrane curvature necessary for clathrin-coated pit
formation (Schmidt et al. 1999). As is the case with most of the players in this
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152
endocytic drama, endophilin is a promiscuous protein, associating with
multiple endocytic partners through, in this case, its SH3 domain. It has been
demonstrated to bind to dynamin and synaptojanin implicating it in fission and
uncoating events (Gad et al. 2000). A role for amphiphysin has also been
suggested in membrane invagination and it has been shown to co-assemble
with dynamin into rings and spirals (Takei et al. 1999). Additionally, it has
been shown to simultaneously bind to dynamin and the coat-protein a-adaptin
(Wigge et al. 1997). It has been suggested that MFs may play a role in the
initial membrane invagination steps (Qualmann and Kelly, 2000). Although
neither endophilin nor amphiphysin has been shown to bind directly to F-actin,
they may indirectly influence MF (de)polymerization by binding to and
activating regulators of actin dynamics, such as dynamin and synaptojanin (see
below). By binding to a-adaptors, endophilin and amphiphysin set the stage
for recruitment of clathrin and coated-pit formation to proceed. More
importantly, as lipid-modifying agents, they cause changes in membrane
composition, such as PIP2 enrichment, that would result in direct and important
consequences for actin polymerization and endocytosis.
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153
A ct I, scene II: the PIP2 connection.
The membrane-phospholipid PIP2 is a key player in endocytosis and is able to
interact and activate multiple components that regulate actin dynamics and
endocytosis (Figure 6.2.), In what pertains to actin MFs, PIP2 promotes actin
polymerization in at least five different ways: (a) uncapping the barbed end of
actin filaments (Takenawa and Itoh, 2001); (b) increasing actin nucleation by
binding to and modulating a variety of actin regulatory proteins such as ezrin
(Defacque et al. 2002); (c) binding to N-WASP BR domain (Miki et al. 1998;
Rohatgi et al. 2000), triggering the N-WASP-Arp2/3 signaling pathway; (d),
activating of Cdc42 signaling pathways (Ma et al. 1998) and (e) regulating
actin cytoskeletal organization (Takenawa et al. 2001; Defacque et al. 2002).
Additionally it can initiate coated-pit formation by binding to recruiting
adaptor proteins (Gaidarov et al. 1999b). PIP2 binding to dynamin has also
been implicated in increasing dynamin’s GTPase activity (Zheng et al. 1996).
Last, but certainly not least, is its ability to bind to dynamin and promote
dynamin insertion into the plasma membrane (Burger et al. 2000). As you may
remember from C hapter 3, dynamin wraps itself around the neck of the
forming coated-vesicle and, by GTP hydrolysis, releases it from the plasma
membrane. However, for PIP2 to mediate dynamin insertion into the
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154
Amphilysin
Binds
dynamin
Binds
a-
C la lh r in
recruitment
see Figure 6.5.
Membrane curvature
Formation of
coated pit
Figure 6.1. Endophilin and amphiphysin signaling pathways to
endocytosis.
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155
membrane, dynamin must first be recruited to the site of endocytosis - a
function fulfilled by endophilin and amphiphysin (see above). It becomes
evident that although the various components of endocytosis are herein
separately described, the various Acts and scenes presented are, by no means,
independent of each other. Rather, it seems cells have created backup
mechanisms in which multiple proteins have redundant functions, activating
diverse signaling pathways which lead to similar end results: actin
polymerization and coated-pit formation, to name two.
Act I, scene III: being “negative ’ is not always bad.
Thus far, clathrin-coated vesicle formation has been triggered and multiple
activated signaling pathways resulted in MF polymerization. At one point,
however, this must stop - too many coated vesicles or filaments is not a good
thing and balance is important. An excess of MFs results in increased
cytoplasmic viscosity (Chapter 3). Also, MF depolymerization has been
demonstrated to be necessary in exocytosis (Chapter 5). To solve this
filamentous problem, enters, in scene III, synaptojanin, who, in the present
drama, has been cast as the negative regulator - mind you, “negative” not in the
“bad” sense, but as a down-regulator of actin polymerization and of
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156
Binding to
other actin
regulatory
proteins
Binds adaptors
(arrestin)
_ Clathrin
recuitment
Phosphatidylinositol 4,5-phosphate (PIP2)
Binds ezrin Uncaps
MF
barbed-Cnd
Binds
N-WASP
BR domain
Arp2/3
activation
Actin polymerization
Facilitates
insertion of
dynamin into
membrane
Activates
Cde42
Small GTPase
activation
Figure 6.2. A role for membrane phospholipids in clathrin-
mediated endocytosis.
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157
endocytosis. Synaptojanin is a phosphoinositide phosphatase (Chapter 4) that
converts PIP2 to PIP. This phosphate removal has a variety of consequences
(Figure 6.3.), namely: (a) by reducing the PIP2 membrane pool, it inhibits
dynamin’s ability to be inserted into the plasma membrane, a necessary step
for its “pinching” activity; (b) it reduces the PIP2 pool available to bind to N-
WASP (essential for N-WASP-Arp2/3 activation of actin polymerization); (c)
it mediates uncoating of the clathrin-coated vesicle. Additionally, it has been
shown to directly bind and sequester profilin, inhibiting its G-actin “shuttle”
activity, consequently inhibiting filament polymerization. Recent studies in
yeast have also demonstrated that synaptojanin may regulate the cellular
distribution if PIP2 (Stefan et al. 2002). Once again, one protein may trigger
various actions, a trend which continues with N-WASP and dynamin.
Act II, scene I: the opening o f N- WASP.
N-WASP has been shown to dramatically increase Arp2/3’s ability to nucleate
actin MFs (Chapter 4). When inactive, it is in a closed conformational state
(the VC A domain is hidden by binding to the GBD domain). Binding of PIP2
and Cdc42 to N-WASP BR and GBD domains, respectively, results in an open
N-WASP conformational state, exposing the VCA domain that can now
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158
activate Arp2/3 (Figure 6.4.), Similar to profilin, the VC A domain also binds
to G-actin and shuttles it to the site of F-actin polymerization. Much however,
must still be resolved about N-WASP activity. Its PRD can bind to a variety of
SH3 domain containing proteins, know to regulate endocytosis, such as
syndapin and Abpl, profilin. The consequences of SH3 to PRD binding,
however, are still unknown. One can envision various hypothetical scenarios:
Abpl does not bind to Arp2/3, but has been implicated in its shuttling to the
side o f actin filaments. Once N-WASP activates Arp2/3 actin nucleation,
binding with Abpl could direct Apr2/3 to a proper site for filament formation.
Profilin and N-WASP VCA domain have redundant G-actin shuttle activity.
Binding of profilin could expedite recruitment of actin monomers for Arp2/3.
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159
Synaptojanin
pip2 — :
'
------► PIP
f
1
Uncoating o f Facilitates Inhibits dy
dathrin-coated cytoskeleton- m em braneii
vesicles bound PIP2
hydrolysis
'
namin Reduces PIP2 Binds to
lsertion available and sequesters
N -W A SP binding profilin
pool
Figure 6.3. Synaptojanin - a negative regulator of endocytosis.
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160
Finally, syndapins have been implicated in a variety of endocytic functions
(Chapter 4). Binding of syndapin to N-WASP may regulate the association
between syndapin and different syndapin binding partners, such as
synaptojanin and the small GTPases, Ras and Rac. Additionally, novel
proteins have been demonstrated to form complexes with N-WASP as well as
G- and F-actin (Ho and Rohatgi, 2001). As well, N-WASP may also play a role
in regulation of actin-based motor proteins as studies have shown it to bind to
the SH3 domain of myosin.
Act III, scene I: To pinch, and so much more!
Without a doubt, dynamin is the protagonist, so to speak, of this endocytic play
given its ability to bind to or activate all of the above-mentioned proteins. In
doing so, it can potentially participate in almost all aspects of clathrin-
mediated endocytosis (Figure 6.5.), Dynamin has been implicated in actin
polymerization through its ability to bind to Cdc42 activating proteins
(Rasmussen et al. 1998) and its downstream effectors: (a) activation of Rac (b)
Cdc42 binding to N-WASP and subsequent Arp2/3 activation. Dynamin’s PH
domain is important in dynamin’s insertion into PIP2 rich membrane regions.
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161
Regulation of motor proteins
Binds
myosin
GBD
N-WASP
Binds Cdc42
VGA.
PRD
PH
Arp2/3
Binds SH3
domain containing
proteins
PIP:
Activation of
small GTPases
Figure 6.2.
Binds Abpl
profilin
Binds
syndapins
Recruits Arp2/3 to M F,
Figure 6.5.
Actin polymerization
Figure 6.4. N-WASP: multiple signaling pathways
to endocytosis.
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162
Other 8113-domam ?
containing proteins " *
or kinases
Binds Cde42
activating kinases
Activated
Cde42 ~
Binds N-WASP
Activates A
PH domain
hinds membrane
PRD
Binds proftiin Binds
Syndapin
’ •Rae
Actiiv polymerkation
Increases
dymnnin
GTPase activity
Binds Synaptojanin
Down-regulation
o f endocytosis
and actin polymerization
Figure 6.5. The GTPase dynamin as a regulator o f endocytosis
- one protein, many signaling cascades.
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163
Similar to N-WASP, dynamin’s PRD can associate with SH3-domain
containing proteins, such as profilin (triggering actin polymerization). Its PRD
domain binds to syndapin (present study), which has been shown to activate
Ras/Rac signaling pathways; furthermore, syndapin has been shown to bind to
synaptojanin (endocytic down-regulator), and N-WASP (actin polymerization).
Dynamin binding to syndapin has also been demonstrated to increase
dynamin’s GTPase activity. Once characterized as simple a “pinchase”,
dynamin function has truly expanded and its importance in clathrin-mediated
endocytosis grows with continued research.
The end? Not quite. As demonstrated by the many diagrams in this chapter,
fully characterizing the many proteins and signaling pathways regulating
clathrin-mediated endocytosis is by no means a trivial pursuit. Although much
has been brought to light, much still needs to be clarified. Future research will
most certainly further elucidate the roles of the many known, and other, still to
be discovered, proteins which regulate this process.
Relevance o f these studies to lacrimal gland function and disease: Previous
research has suggested a role for MFs in endocytosis in many
simple systems although in many cases the studies have been inconclusive.
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164
Recent work using inhibitors of MF assembly has implicated MFs specifically
in apical endocytosis in epithelial cells and in pancreatic acini (Jackman et al,
1994; Shurety et al, 1996 add Gottlieb et al. 1993; Valentijn et al. 1999).
However, this is, to my knowledge, the first demonstration of a role for MFs
and for the actin regulatory proteins syndapins, N-WASP, Abpl and
synaptojanin as facilitators of apical endocytosis in any epithelial cell system.
These findings are important in that they begin to elucidate the complex
pathways involved in apical endocytosis. Here, I have demonstrated, by
confocal microscopy and biochemical analysis, that N-WASP, synaptojanin,
SdpI, SdpII and Abpl are present in lacrimal acini. Additionally, GST-pull
down assays confirmed that the SH3 domain of SdpI, SdpII and Abpl
associate with dynamin, synaptojanin and N-WASP. This is the first
demonstration of the binding of these SH3 domains to components
of the clathrin-mediated endocytic machinery in any epithelial cell, and
is, to my knowledge, the first demonstration of Abpl binding to N-WASP in
any system. Finally, my functional studies suggest that these proteins
function in apical endocytosis in lacrimal acini, confirming that these
proteins have a conserved role from neurons to secretory epithelia. My
findings clearly implicate clathrin-mediated endocytosis, acting in concert with
these novel effectors (MFs, synaptojanin, syndapins, Abpl and N-WASP) in
apical membrane retrieval in lacrimal acini.
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165
Further elucidation of the mechanisms that regulate membrane trafficking
in lacrimal acinar epithelial cells is critical to understanding their function. It is
vital, for cell function, that membranes and proteins be delivered to their
targeted site, be it the apical or basolateral membrane or an internal organelle.
Errors in membrane trafficking and missorting events can result in inadequate
or inefficient cell function and may trigger disease states. We have
hypothesized that such conditions may underlie lacrimal gland autoimmune
diseases. Previous research in our lab (da Costa et al, 1998) demonstrated that
supramaximal stimulation of acini (1 mM CCH) resulted in a decrease in
secretion and release of protein compared to stimulation at maximal, lower,
CCH dose (10 mM). It is our hypothesis that this decrease in protein release
may be the result of the missorting of apically targeted secretory products,
which are sorted to basolateral membrane compartments instead. This theory
has strong implications for diseases such as Sjogren's syndrome. We further
hypothesize that the possible missorting of membranes to basolateral
compartments may be one of the events that triggers the initiation of lacrimal
gland autoimmunity. Others have also recently suggested a role for epithelial
cells in Sjogren's disease (Amft and Bowman, 2001).
Research that further elucidates the signaling pathways and endocytic
mechanisms involved in accurate segregation and sorting of apically- and
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166
basolaterally-targeted materials is also essential at a broader level to
understanding epithelial cell function. Given the fact that polarized epithelial
cells have in common various morphological and functional similarities,
information derived from lacrimal gland acinar cells may provide valuable
insights into the membrane transport mechanisms of other epithelial cells, such
as pancreas or parotid glands, epithelial cells of the gut or the airway
epithelium. This information is relevant, not only in terms of identifying the
source of diseases which may result from deficiencies in membrane transport
(such as Sjogren's in lacrimal gland or acute pancreatitis in pancreatic acinar
epithelia), but especially in terms of treatment of disease. Access of drugs to
many targeted cells occurs via the apical membranes of the gastrointestinal
tract, lung epithelium, corneal epithelium, and other barrier tissues. In many
cases, drug entry is via receptor-mediated endocytosis may be an approach to
improve entry into the targeted cell. Increased understanding of the complex
machinery governing apical endocytosis of drug macromolecules is a key step
in improving drug delivery and targeting.
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167
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Creator da Costa, Silvia Regina (author)

Core Title How cells cry: The cytoskeleton as a facilitator of regulated secretion and membrane trafficking in lacrimal gland acinar cells

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Advisor Hamm-Alvarez, Sarah (committee chair), Lee, Vincent H.L. (committee member), McDonough, Alicia (committee member), Mircheff, Austin K. (committee member), Okamoto, Curtis T. (committee member)

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