Role of the Interaction of proHB-EGF with Heparan Sulfate
Proteoglycans
MASSACHUSETTS INSTITUTE
OF TECHNOLOGY
by
I AUG 16 2010
Robin N. Prince
B.S. Mechanical Engineering
University of Arkansas, 2003
1
S.M. Mechanical Engineering
Massachusetts Institute of Technology, 2005
LIBRARIES
ARCHNES
SUBMITTED TO THE DEPARTMENT OF BIOLOGICAL ENGINEERING IN
PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY IN BIOLOGICAL ENGINEERING
AT THE
MASSACHUSETTS INSTITUTE OF TECHNOLOGY
AUGUST 2009
C 2009 Massachusetts Insitute of Technology, All Rights Reserved
The author hereby grants to MIT permission to reproduce and to distribute
publicly paper and electronic copies of this thesis document in whole or in part
in any medium now known or hereafter created.
Signature of Author:
Department of Biological Engineering
August 14, 2009
Certified By:
Dou's.7
auffenburger
Professor of Biological Engineering
Thesis Supervisor
Certified By:
Richard T. Lee
Professor of Medicine, Brigham and Women's Hospital, Harvard Medical School
Thesis Supervisor
Accepted By:
tRoger
D. Kamm
Professor of Biological Engineering
Thesis Committee Chair
Thesis Committee:
Douglas A. Lauffenburger, Thesis Supervisor, MIT
Richard T. Lee, Thesis Supervisor, BWH & Harvard Medical School
Roger D. Kamm, MIT
Matthew Nugent, Boston University School of Medicine
Role of the Interaction of proHB-EGF with Heparan Sulfate
Proteoglycans
by
Robin N. Prince
Submitted to the Department of Biological Engineering on August 24, 2009
in partial fulfillment of the requirements for the degree of
Doctor of Philosophy in Biological Engineering
Abstract
Heparin-binding epidermal growth factor-like growth factor (HB-EGF) exhibits
activity as a juxtacrine, paracrine, and autocrine ligand for the epidermal growth factor
receptor (EGFR), and possesses the ability to bind heparan sulfate proteoglycans
(HSPGs). The interaction of HB-EGF with HSPGs has been previously studied only with
the soluble (autocrine/paracrine) form of the protein (sHB-EGF), produced after
proteolytic cleavage of the transmembrane form (proHB-EGF) from the cell surface. It
was hypothesized that HSPGs interact with proHB-EGF in ways that could alter behavior
of the transmembrane form of this ligand and consequent processes. Using an engineered
form of proHB-EGF that allowed for independent tracking of the extracellular domain
and the C-terminal tail, proHB-EGF was observed primarily at sites of cell-cell contact.
However, a dramatic change in this localization was observed upon the addition of
exogenous heparin, heparan sulfate, heparinase III or mutation of the heparin-binding
domain of proHB-EGF, suggesting that an interaction with HSPGs is responsible for
localizing proHB-EGF to sites of cell-cell contact. Further studies in wild-type CHO-Ki
cells and heparan sulfate deficient CHOpgsD-677 cells demonstrated that a trans
interaction between proHB-EGF and HSPGs on neighboring cells was responsible for
this localization. Additionally, this interaction inhibited proteolytic processing of the
ligand, as heparin and mutation of the heparin-binding domain increased the amount of
sHB-EGF accumulated in the media.
Acknowledgements
This work would not have been possible without the support and guidance of my
two wonderful advisors, Douglas Lauffenburger and Richard T. Lee. I would like to
thank them for their excellent mentorship, guidance, support, and allowing me to pursue
my own avenues of investigation and interest during my graduate studies. I would like to
thank our collaborators Steve Wiley and Alice Ting, who have provided scientific insight
and reagents that were crucial for this work. I am additionally grateful to my thesis
committee members, Matthew Nugent and Roger Kamm for their guidance and scientific
advice. I also would like to recognize Linda Griffith for her enthusiastic support and
mentorship through my graduate research endeavors.
I would like to thank all members of the Lee, Lauffenburger, and Griffith labs, as
I have learned a great deal being part of them all and have thoroughly enjoyed my time
here. I would particularly like to recognize and thank Jun Yoshioka for providing
excellent training that was crucial in my transition to a biological researcher. Special
thanks go to Eric Shreiter for his help in designing the acceptor peptide HB-EGF
construct, Peng Zou for making mSA-AF568, and Hyung Do Kim for doing monolayer
migration experiments. I would like to thank my funding sources, the NSF graduate
fellowship and the Poitras Predoctoral Fellowship.
To Mom, Dad, Wade, and Grandma, I thank you all for your love, support,
patience and encouragement. To my boyfriend, Ohia, thank you for all your encouraging
advice, your love and support, and your always open ear. Also, thank you to so many
great friends and colleagues I have met in Boston, you have all made my time here so
special. I would additionally like to recognize my undergraduate research mentor Ajay
Malshe at the University of Arkansas, and my high school and mentors at the Arkansas
School for Mathematics and Sciences, as they were crucial in preparing me for this life
path.
Table of Contents
Abstract ...............................................................................................................................
3
Acknowledgements......................................................................................................
4
Table of C ontents..........................................................................................................
5
Chapter One: Introduction .............................................................................................
9
1.1.1 Growth factor signaling ....................................................................................
9
1.1.2 The EGFR system.............................................................................................
9
1.1.3 HB-EGF autocrine signaling and EGFR transactivation..................................... 11
1.1.4 Heparin and heparan sulfate proteoglycans ......................................................
12
1.1.5 Physiologic and pathophysiologic roles of HB-EGF........................................
13
1.1.5.1 HB-EGF in development ...........................................................................
13
1.1.5.2 Diphtheriatoxin .........................................................................................
14
1.1.5.3 HB-EGF in wound healing ........................................................................
15
1.1.5.4 HB-EGF in cancer....................................................................................
15
1.1.5.5 HB-EGF in the cardiovascularsystem ......................................................
16
1.1.5.6 HB-EGF in pregnancy ...............................................................................
17
1.1.5.7 HB-EGF in the kidney ................................................................................
18
Chapter Two: Experimental approach for visualization of HB-EGF ...........................
20
2 .1 Introdu ction ............................................................................................................
20
2.2 Materials and Methods.......................................................................................
23
2 .3 Resu lts....................................................................................................................
30
2 .4 D iscussion ..............................................................................................................
34
2 .5 F igu res.....................................................................................................................
35
Figure 2.5.1: Two methods of BirA (biotin ligase) labeling of the acceptor peptide
pro tein. ......................................................................................................................
35
Figure 2.5.2. Design of a monovalent streptavidin.............................................
36
Figure 2.5.3 Validation of AP-HBEGF constructs...............................................
37
Figure 2.5.4 Final gene map of AP-HBEGF-GFP and bioactivity....................... 38
Figure 2.5.5 AP-HBEGF-GFP is localized to cell-cell contact sites.................... 39
Chapter Three: Polarization of HB-EGF at the wound edge ........................................
41
3 .1 Introdu ction ............................................................................................................
41
3.2 Materials and Methods.......................................................................................
43
3 .3
Resu lts...............................................................................................................
44
3.3.1 ProHB-EGF is missing from the wound edge ................................................
44
3.3.2 HB-EGF loss from wound edge is not due to proteolysis ..............
45
3.3.3 Newly synthesized HB-EGF localizes to cell-cell contact sites only ......... 46
3 .4 D iscu ssion ..............................................................................................................
48
3 .5 F igures....................................................................................................................
49
Figure 3.5.1 HB-EGF is absent from the wound edge....................
49
Figure 3.5.2 Effect of inhibitors on polarization of HB-EGF at wound edge. ........ 50
Figure 3.5.3 Turnover of cell surface HB-EGF at wound edge............................. 51
Chapter Four: The heparin-binding domain mediates localization of proHB-EGF to cellcell contact sites................................................................................................................
53
4 .1 Introdu ction ............................................................................................................
53
54
4.2 M aterials and M ethods......................................................................................
-- - .........- 59
4.3 Results........................................................................
59
proHB-EGF..............
of
localization
the
alter
sulfate
4.3.1 Heparin and heparan
BirA
to
4.3.2 ProHB-EGF in microdomains at cell-cell contact sites is inaccessible
..... 60
and streptavidin.................................................................................
site
4.3.3 Pro-HB-EGF interaction with HSPGs controls cell-cell contact
- ......... 61
..
localization..........................................................................
62
4.3.4 ProHB-EGF interacts with HSPGs in trans ...............................................
4.3.5 Heparin-binding controls amphiregulin localization, but engineered heparin64
binding is insufficient for cell-cell contact localization.........................................
... ..... 66
. ...
4.4 D iscussion.......................................................................
66
4.4 Discussion................................................................................
69
. ---.... - ----- -----..................
4.5 Figures.................................................................-------.....
from
HB-EGF
of
localization
the
changed
Figure 4.5.1. Heparin and heparan sulfate
cell-cell contact sites to a homogenous distribution over the cell surface............ 69
Figure 4.5.2 Localization change of HB-EGF after addition of heparin over time.. 70
Figure 4.5.3 Heparin increases accessibility of cell surface AP-HBEGF-GFP....... 71
Figure 4.5.4 HSPGs target pro-HB-EGF to cell-cell contact sites. ..................... 72
Figure 4.5.5. The heparin-binding domain targets pro-HB-EGF to cell-cell contact
73
sites. .................................................................................---------------------------..............
for
required
are
Figure 4.5.6 HSPGs and the heparin-binding domain of HB-EGF
74
localization to cell-cell contact sites. .....................................................................
75
Figure 4.5.7. HSPGs interact in trans with pro-HB-EGF. ...................................
Figure 4.5.8 ProHB-EGF is preferentially localized to cell-cell contact sites when
76
neighbored by a non-expressing cell....................................................................
Figure 4.5.9 The heparin-binding domain of amphiregulin controls localization to
... 77
cell-cell contact sites ......................................................................................
80
Chapter Five: Role of the heparin-binding domain ......................................................
80
5.1 Introduction................................................................................
... 81
5.2 M aterials and Methods..................................................................................
83
---------------------...................
.
5.3 Results............................................................................
5.3.1 The heparin-binding domain controls cell surface localization.................... 83
86
5.3.2 ProHB-EGF does not cluster EGFR ...............................................................
5.3.3 The heparin-binding domain controls ectodomain shedding........................ 88
89
5.3.4 Wound healing and migration.......................................................................
91
5.4 Discussion.................................................................................
. ---------------------.............. 93
5.5 Figures............................................................................----.
the cell surface
decreases
domain
heparin-binding
Figure 5.5.1 Mutation on the
93
fraction of HB-EGF. ........................................................................
Figure 5.5.2 The heparin-binding domain mutant is primarily localized in the
..... 94
intracellular space. .................................................................................
95
activation....................................................
Figure 5.5.3 EGFR localization and
96
Figure 5.5.4 Heparin reduces ERK activation. .....................................................
97
cleavage.................
pro-HB-EGF
reduces
Figure 5.5.5. Interaction with HSPGs
98
Figure 5.5.6 Wound healing in COS-7 cells.........................................................
99
monolayer.
confluent
a
within
HMECs
Figure 5.5.7 Migration speed of individual
Chapter Six: Conclusions and future directions.............................................................
6.1 Future directions: Use of AP-tagged EGFR ligands............................................
6.2 Future directions: Signaling with heparin-binding domain mutant HB-EGF......
6.3 Future directions: HB-EGF localization change in vivo.....................................
6.4 Future directions: Structure of the HSPG-proHBEGF-CD9 complex................
6.5 Future Directions: Computational analysis.........................................................
Referen ces.......................................................................................................................
101
101
103
103
104
104
106
8
Chapter One: Introduction
1.1.1 Growth factor signaling
Signaling by soluble extracellular ligands is classified by the distance in which the
signal acts and the source of the ligand.
Free ligands can signal in an endocrine,
paracrine, or autocrine manner. Endocrine signaling molecules travel the farthest, from
an endocrine organ to target cells, where the molecules typically travel through blood or
extracellular fluids (Lodish, 2003). However, on a much shorter length scale, paracrine
and autocrine signaling affect cells only in close proximity. In paracrine signaling, a cell
produces a soluble ligand which diffuses to and binds another cell. Autocrine signaling
differs in that the cell which produces the soluble ligand is also activated by that ligand.
However, not all extracellular signals are diffusible. Some growth factors are active in
their membrane bound form before cleavage, which defines juxtacrine signaling.
1.1.2 The EGFR system
The epidermal growth factor receptor (EGFR) system is a well studied example of
cell communication in regards to growth, motility, and development. The EGFR family
consists of four receptor tyrosine kinases:
erbB1/HER1, referred to as the epidermal
growth factor receptor (EGFR), erbB2/HER2, erbB3/HER3, and erbB4/HER4.
Upon
ligand binding, the EGF receptors dimerize and stimulate intracellular signal transduction
pathways which encourage the cell to proliferate, differentiate, or survive (Lodish 2004).
Ligands for the EGFR system include HB-EGF, epidermal growth factor (EGF),
amphiregulin, transforming growth factor-a (TGF-a), neuregulin, betacellulin and
epiregulin. Ligands in the EGFR family are synthesized in membrane anchored proforms and are subsequently cleaved at an extracellular site via a metalloprotease to
release the soluble growth factor (Asakura et al., 2002; Sahin et al., 2004). However, the
ability of some EGFR ligands to activate their corresponding receptors is not dependent
on cleavage; both the pro-membrane and soluble forms can serve as cellular signals
(Massague and Pandiella, 1993). Binding of ligands to receptor tyrosine kinases activates
various intracellular signaling pathways. In particular, EGFR activation signals through
the Ras-MAP kinase pathway. Activation of the EGFR leads to the recruitment of Grb2
and Shc, which leads to activation of the intercellular membrane-bound protein, Ras. Ras
then activates an intracellular kinase cascade including MAP kinase, also known as ERK,
Jun N-terminal Kinase (JNK), and p38, which are active in their phosphorylated form.
Activated ERK dimerizes and can translocate into the nucleus to activate various
transcription factors. In addition to the Ras-MAP kinase pathway, the EGFR has also
been linked to the JAK-STAT pathway.
JAK is a tyrosine kinase and STAT is a
transcription factor.
HB-EGF is expressed primarily in the lung, skeletal muscle, brain and heart
(Abraham et al., 1993) and induces proliferation and migration of smooth muscles cells,
fibroblasts and keratinocytes (Raab and Klagsbrun, 1997; Yahata et al., 2006). Heparinbinding EGF (HB-EGF) was discovered in 1991 in the medium of cultured macrophagelike cells (Higashiyama et al., 1991). HB-EGF has several key features which differ from
traditionally studied EGF. HB-EGF shares the same binding domain to EGFR as EGF
which consists of six conserved cysteines, however, only approximately 40% of the
carboxyl portion protein sequence is homologous to EGF (Higashiyama et al., 1991).
Additionally, as its name suggests, HB-EGF binds strongly to heparin. EGF is selective
for the EGFR, yet HB-EGF binds to EGFR, erbB4/HER4 (Elenius et al., 1997), and
heparan sulfate proteoglycans (Higashiyama et al., 1993) present on the cell surface and
within the extracellular matrix. HB-EGF is an 8.3 kDa protein; however, after heavy 0glycosylation it has an estimated mass of 20-22 kDa on an SDS-PAGE gel.
The ability of HB-EGF to signal in a juxtacrine manner is increased by association
with the tetraspanin protein CD9 (Higashiyama et al., 1995; Iwamoto et al., 1991), which
also links HB-EGF to a31 integrins (Nakamura et al., 1995). In vivo staining shows co-
localization of all three proteins in many tissues (Nakamura et al., 2001).
The
membrane-anchoring domain of HB-EGF is also crucial in juxtacrine signaling, as
domain swapping with the membrane-anchoring domain of EGF, which does not
participate in juxtacrine signaling, led to loss of the ability of HB-EGF to signal in a
juxtacrine manner (Dong et al., 2005). Pro-HB-EGF has also been reported to interact
with the tetraspanins CD63, CD81 and CD82 on the cell surface; however, no known
phenotype has been reported for this interaction (Nakamura et al., 2000). Interestingly,
HB-EGF juxtacrine and autocrine/paracrine signaling have been shown to elicit different
phenotypes, with autocrine/paracrine activity leading to cell proliferation and juxtacrine
activity leading to growth inhibition in some cell lines. DER cells, a hematopoietic cell
line, and human luteinized granulosa cells undergo growth inhibition and apoptosis when
stimulated with proHB-EGF (Iwamoto et al., 1999; Pan et al., 2002). However, proHBEGF juxtacrine signaling has also been reported to protect renal epithelial cells and
Madin-Darby canine kidney epithelial cells from apoptosis (Singh et al., 2007b;
Takemura et al., 1997).
Therefore, the cell fate resulting from HB-EGF juxtacrine
stimulation may be cell-type specific.
Recently, it has been reported that the C-terminal fragment of HB-EGF is also a
signaling molecule. This fragment translocates to the nucleus, where it reverses gene
repression of the transcriptional repressors promyelocytic leukemia zinc finger protein
(PLZF) and Bcl6 (Kinugasa et al., 2007; Nanba et al., 2003). The C-terminal tail of HBEGF can interact with BAG-1, which can increase proHB-EGF proteolysis, increase
resistance to apoptosis, and decrease cell adhesion (Lin et al., 2001). The cytoplasmic
domain of HB-EGF is phosphorylated after treatment with various stimuli, and mutation
of this phosphorylation site does not affect ligand cleavage, but reduces tumorigenicity
(Wang and Dey, 2006).
1.1.3 HB-EGF autocrine signaling and EGFR transactivation
Upon the discovery of autocrine signaling in the EGFR system, it was originally
hypothesized to be a malignant phenotype (Spom and Todaro, 1980).
However,
autocrine signaling has become widely accepted as a major form of cell communication
regulating a host of different processes.
Those specific to HB-EGF include wound
healing in corneal epithelial cells (Xu et al., 2004) and keratinocytes (Piepkorn et al.,
1998), and the response to mechanical stress in bronchial epithelial cells (Chu et al.,
2005). In the HB-EGF/EGFR system, autocrine signaling is initiated with proteolytic
cleavage of HB-EGF with a disintegrin and metalloprotease (ADAM). The specific
ADAM which is responsible for HB-EGF cleavage is a topic of debate and tends to vary
depending on cell type, however ADAMS 10, 12, and 17 have been identified
(Higashiyama and Nanba, 2005).
HB-EGF plays a prominent role in EGFR
transactivation, typically through activation of G-protein coupled receptors (GPCRs).
Activation of various GPCRs with GPCR agonists, such as angiotensin II (Yahata et al.,
2006), endothelin-1 (Chansel et al., 2006), lysophosphatidic acid (Xu et al., 2007), ATP
(Yin and Yu, 2009), histamine (Ancha et al., 2007), interleukin-8 (Itoh et al., 2005), and
extracellular Ca (Yano et al., 2004) has been demonstrated to lead to HB-EGF cleavage.
After proteolytic release, the mature, soluble form of HB-EGF is then free to diffuse and
bind to HSPGs, EGFRs or erbB4 receptors on the cell surface.
This system has the
potential for positive feedback in that EGFR activation can lead to transcription of
additional growth factors and activation of EGFR ligand shedding (Citri and Yarden,
2006).
1.1.4 Heparin and heparan sulfate proteoglycans
HB-EGF was first identified as a growth factor purified from a heparin column
which stimulated fibroblast and smooth muscle cell growth (Higashiyama et al., 1991).
HB-EGF has the ability to bind heparin via a heparin-binding domain that consists of
multiple basic lysine and arginine residues, which interact with negatively charged
heparin (Thompson et al., 1994). Heparin, a highly sulfated glycosaminoglycan, is a
widely used anticoagulant as it activates antithrombin, which inactivates blot clotting
enzymes, such as thrombin and factor Xa (Chuang et al., 2001). In humans, heparin is
produced by mast cells residing in vascularized serosal cavities, and upon degranulation
heparin is released (Kalesnikoff and Galli, 2008).
Proteoglycans are a type of glycoprotein which have glycosaminoglycan (GAG)
side chains.
Four major classes of proteoglycans exist, which are classified by the type
of GAG chains expressed: heparan sulfate, chondroitin sulfate, dermatan sulfate, and
keratan sulfate. Heparan sulfate proteoglycans (HSPGs) consist of repeating units of Dglucouronic or L-iduronic acid with N-acetyl or N-sulfo-D-glucosamine disaccharides
with chains approximately 200 units long (Lodish 2004). As heparan sulfate is similar to
heparin in its structure and sulfation pattern, it also has the ability to bind HB-EGF and
other heparin-binding growth factors.
Likely, it is the interaction of HB-EGF with
HSPGs that is more physiologically relevant, as HSPGs are present in the extracellular
matrix and cell surface glycocalyx of most cells. HSPGs have the ability to modulate the
activity of many heparin-binding growth factors. The heparin-binding domain of HBEGF appears to be inhibitory, however, upon heparin or heparan sulfate binding, the
ability of HB-EGF to activate the EGFR is increased (Higashiyama et al., 1993; Takazaki
et al., 2004).
Heparan sulfate side chains and heparin can be altered via enzymatic cleavage.
There are three major heparin/heparan sulfate lyases purified from Flavobacterium
heparinum (heparinase I, II and III), which cleave specific linkages present in heparin
and/or heparan sulfate (reviewed in (Capila and Linhardt, 2002)) and are commonly used
for experimental purposes. Mammalian heparanase is a pro-enzyme, that is cleaved into
two subunits that associate to form active heparanase (reviewed in (Vlodavsky et al.,
2007)). Additionally, there are the enzymes HSulf-1 and HSulf-2, which are secreted
endosulfatases that degrade heparan sulfate (Morimoto-Tomita et al., 2002).
1.1.5 Physiologic and pathophysiologic roles of HB-EGF
1.1.5.1 HB-EGF in development
HB-EGF is an important growth factor in mammalian development, particularly
for the cardiovascular system.
Most HB-EGF knockout mice die within the first
postnatal week and have grossly enlarged ventricular chambers and cardiac valves
(Iwamoto et al., 2003), while smooth muscle and endothelial cell specific knock-outs
show a similar phenotype (Nanba et al., 2006). Mice expressing only a non-cleavable
form of proHB-EGF also die of severe heart failure, suggesting that autocrine stimulation
is required for proper heart development, while mice expressing the soluble form of HBEGF, lacking the C-terminal and transmembrane domain, develop severe hyperplasia of
the skin and heart (Yamazaki et al., 2003). HB-EGF expression is observed in smooth
muscle cells of the aortic wall of babies and children, however, expression is decreased in
young adults (Miyagawa et al., 1995).
HB-EGF also plays a role in the development of other tissues and organs. HBEGF knock-out mice show enlarged mesenchymal tissue of the lung and heart, and have
immature aveoli (Jackson et al., 2003).
HB-EGF knock-out mice also have delayed
eyelid closure during development, where HB-EGF is expressed solely at the leading
edge of the migrating epithelial sheet (Mine et al., 2005).
Mice expressing an
uncleavable form of HB-EGF display identical defects in wound closure, suggesting that
HB-EGF autocrine signaling is crucial for driving persistent cell migration to close the
wound (Mine et al., 2005). HB-EGF is expressed at higher levels in the embryonic and
neonatal kidney compared to the adult, and may play a role in the development of renal
collecting ducts and promote renal epithelial cell branching (Takemura et al., 2001;
Takemura et al., 2002).
1.1.5.2 Diphtheriatoxin
ProHB-EGF was first studied as the unidentified receptor for diphtheria toxin
(Naglich et al., 1992). Diphtheria was once a serious world health threat that has largely
been eliminated in developed nations after introduction of a vaccine in the early 1900s.
Diphtheria is caused by the protein diphtheria toxin, which is secreted by
Corynebacteriumdiphtheriae. The toxin consists of two fragments (A and B chain), of
which the B chain binds to the EGF-like domain of proHB-EGF and serves as the toxin's
route to enter the cell via receptor-mediated endocytosis (Naglich et al., 1992). Once
internalized, the A chain inhibits protein synthesis by catalyzing ADP-ribosylation of
elongation factor-2, rendering it inactive (reviewed in (Neville and Hudson, 1986)). Just
as association of proHB-EGF with CD9 upregulates juxtacrine activity, it also increases
binding affinity and sensitivity of diphtheria toxin binding to proHB-EGF (Cha et al.,
2000; Iwamoto et al., 1991). Additionally, the association of proHB-EGF with heparin or
HSPGs on the cell surface increases in the affinity of diphtheria toxin for proHB-EGF
(Shishido et al., 1995). The conjugation of bacterial immunotoxins, such as diphtheria
toxin, to cell specific targeting molecules is in investigation for treatment of cancer
(Reviewed in (Kreitman, 2009)). Additionally, proHB-EGF's role as the diphtheria toxin
receptor has been used for targeted cell ablation by expressing the human form of proHBEGF in a targeted cell type within the mouse (Saito et al., 2001). As diphtheria toxin
does not interact with rat or mouse proHB-EGF, transgenic human proHB-EGF
expression under a tissue or cell-specific promoter in mice can lead to conditional and
tissue-specific cell ablation by administering diphtheria toxin. CRM-197 is a non-toxic
mutant of diphtheria toxin, which inhibits HB-EGF by preventing it from binding to EGF
receptors (Mitamura et al., 1995) and is under investigation as a treatment for ovarian
cancer (reviewed in (Miyamoto et al., 2006))
1.1.5.3 HB-EGF in wound healing
HB-EGF autocrine signaling is a crucial regulator of keratinocyte migration and
re-epithelialization in skin wound healing.
HB-EGF leads to an increased rate of
keratinocyte migration (Shirakata et al., 2005; Tokumaru et al., 2000), and topical
application to bums increases wound closure (Cribbs et al., 1998). Additionally, HBEGF is also found in wound fluid (Marikovsky et al., 1993). HB-EGF expression is
upregulated in keratinocytes at the margin of the wound and expression is increased by
disruption of lipid rafts (Mathay et al., 2008; McCarthy et al., 1996). HB-EGF also plays
a role in closing wounds of the eye, as it is an autocrine factor secreted by corneal
epithelial cells in response to wounding, and inhibition of HB-EGF leads to impaired
wound closure (Xu et al., 2004). Experiments suggest that sudden reduction of spatial
constraints is sufficient for HB-EGF release and EGFR activation to stimulate corneal
epithelial cell migration (Block et al., 2004).
1.1.5.4 HB-EGF in cancer
HB-EGF gene expression is increased in many types of cancer, including tumors
of the pancreas, liver, esophagus, skin, colon, stomach, ovary, bladder and brain and is
associated with the acquisition of malignant phenotypes (reviewed in (Miyamoto et al.,
2006)).
Additionally,
HB-EGF
is involved
in
chemotherapy
resistance,
as
chemotherapeutic agents can cause HB-EGF ectodomain shedding and protect tumor
cells from apoptosis (Wang et al., 2007). Much attention has been directed to the role of
HB-EGF in ovarian cancer, as HB-EGF is associated with poor clinical outcome (Tanaka
et al., 2005). High mortality is predominantly caused by spread of the tumor into the
peritoneal cavity (reviewed in (Miyamoto et al., 2008)). HB-EGF appears to be involved
in tumor formation and survival in the peritoneal cavity, as tumor formation by injection
of the human ovarian cancer cell line SKOV3 and RMG1 in nude mice is enhanced by
proHB-EGF expression and blocked completely blocked by inhibition of HB-EGF gene
expression (Miyamoto et al., 2004). Soluble HB-EGF levels are significantly elevated in
the peritoneal cavity at levels sufficient for ovarian cancer cell survival (Miyamoto et al.,
2004).
1.1.5.5 HB-EGF in the cardiovascularsystem
HB-EGF is a strong mitogen and chemoattractant for smooth muscle cells, whose
expression is significantly increased in smooth muscle cells and macrophages of
atherosclerotic plaques (Miyagawa et al., 1995; Nakata et al., 1996). Therefore HB-EGF
has been suggested to play a role in smooth muscle cell proliferation and migration in
atherosclerosis. HB-EGF expression is increased in restricted carotid arteries, and is
associated with increased lumen narrowing, thickening of the artery wall, and increased
circumference that is not observed in HB-EGF knock-out mice (Zhang et al., 2008).
EGFR transactivation via HB-EGF shedding in response to GPCR agonists additionally
plays a role in vascular biology, as HB-EGF induced EGFR activation is necessary for
endothelin-1 induced vasoconstriction response (Chansel et al., 2006), and HB-EGF is
shed by angiotensin II stimulation in smooth muscle cells, causing proliferation and
migration (Yang et al., 2005).
Cardiac hypertrophy occurs when individual cardiomyocytes (the beating heart
cell) expand in size under excess mechanical force to increase output and meet
physiologic demands (Chien, 1999). As a normal adaptive response mechanism, cardiac
hypertrophy is not dangerous until the condition of persistent stress over time evolves
into dysfunction and myocardial failure (Chien, 1999). Cardiac hypertrophy is associated
with an increase in cell size, protein synthesis, and re-expression of fetal genes
(Sadoshima and Izumo,
1997).
Increasing evidence
suggests that stress on
cardiomyocytes leads to production of endothelin-1, which induces metalloprotease
cleavage of pro-HB-EGF (Anderson et al., 2004). The soluble growth factor is then free
to diffuse and activate the EGFR and subsequent intracellular signaling pathways which
lead to cardiac hypertrophy. Evidence which supports this hypothesis includes: strain on
myocytes activates the EGFR and increases the concentration of HB-EGF in cell medium
(Anderson et al., 2004), inhibition of pro-HBEGF cleavage prevents GPCR agonist
induced hypertrophy (Asakura et al., 2002); and inhibition of NAD(P)H oxidase, which
leads to endothelin-1 release, inhibits HB-EGF shedding (Anderson et al., 2004). Studies
suggest that ADAM12 is the metalloprotease responsible for hypertrophic HB-EGF
shedding in the heart (Asakura et al., 2002).
overexpression
Our own studies with adenovirus
of HB-EGF the heart show that HB-EGF expression leads to
cardiomyocyte hypertrophy and degradation of connexin43, a crucial gap junctional
protein in the heart in a localized area around the site of transfection (Yoshioka et al.,
2005). This study also demonstrates that the spatial range of HB-EGF diffusion in the
heart was very restricted, only affecting the cell which produced HB-EGF and its
immediately adjacent neighbor.
1.1.5.6 HB-EGF in pregnancy
HB-EGF expression 6-7 hours prior to attachment of the blastocyst to the luminal
epithelium is the earliest known marker for blastocyst implantation. The blastocyst exists
early in embryogenesis and is covered by an outer layer of cells composing the
trophectoderm. During implantation, the blastocyst adheres to the luminal epithelium of
the endometrial wall, and stromal cells that surround the blastocyst decidualize to embed
the embryo in the stromal bed (reviewed in (Wang and Dey, 2006)).
HB-EGF is
involved in early cross-talk between the luminal epithelium and the blastocyst during
implantation. ErbB4 and EGFR are expressed at the apical surface of trophectoderm
cells (Paria et al., 1999).
Pro-HBEGF is expressed in the luminal epithelium, only
surrounding the blastocyst, and juxtacrine interaction with EGFR and HSPGs on the
blastocyst increases attachment during the implantation process (Das et al., 1994; Raab et
al., 1996). Blastocysts synthesize the HSPG perlecan on the trophectodermal surface,
which is required for attachment, and this synthesis is increased during the periimplantation period (Carson et al., 1993; Farach et al., 1988). Addition of exogenous
heparin or digestion of cell surface heparan sulfate with heparinase considerably reduces
the rate of attachment of embryos to monolayers of uterine epithelial cells (Farach et al.,
1987).
The blastocyst itself also expresses HB-EGF, and the paracrine HB-EGF
produced may trigger HB-EGF gene expression in the luminal epithelium (Hamatani et
al., 2004). HB-EGF null mice show deferred implantation, however, amphiregulin, but
not epiregulin, can partially compensate for the loss of HB-EGF (Xie et al., 2007).
1.1.5.7 HB-EGF in the kidney
In the adult kidney, HB-EGF is primarily localized to tubular epithelial cells of
the S3 segment of the outer stripe in the outer medulla (Nakagawa et al., 1997). HB-EGF
expression is induced in the distal tubules injured by ischemia/reperfusion (Takemura et
al., 1997), in epithelial cells involved in the formation of lesions of focal and segmental
glomerular sclerosis (Paizis et al., 1999), in renal epithelial cells in the obstructed kidney
acting to inhibit apoptosis (Nguyen et al., 2000), in mesangial cells driving proliferation
in glomerulonephritis (Takemura et al., 1999b), and in polycystic kidney disease
(MacRae Dell et al., 2004).
Interestingly, proHB-EGF juxtacrine signaling, but not
autocrine/paracrine signaling, promotes renal epithelial cell survival which is increased
by co-expression of CD9 (Takemura et al., 1999a; Takemura et al., 1997) and increases
transepithelial resistance in polarized Madin-Darby canine kidney (MDCK) cells (Singh
et al., 2007a). CD9 and
p1
integrin expression are upregulated in the medullary thick
ascending limbs (nephron segments that are normally exposed to higher and variable
extracellular osmolality) after dehydration, where HB-EGF is also expressed, suggesting
they may form an osmotically relevant membrane signaling complex (Sheikh-Hamad et
al., 2000). Urea activation of GPCRs in renal medullary cells leads to HB-EGF cleavage
and EGFR activation, which may prevent hypertonic stress induced damage (Zhao et al.,
2003).
19
Chapter Two: Experimental approach for visualization of HBEGF
2.1 Introduction
Traditional methods of visual protein tracking in cell culture is accomplished with
either antibody detection or fusion of a fluorescent protein. Antibody detection, also
known as immunocytochemistry, requires an antibody that recognizes the protein or an
epitope tag which has been fused to the protein of interest. However, antibody detection
for intracellular domains of the protein requires cell fixation and permeabilization, and
therefore rules out any live cell tracking of the protein of interest. Fusion of a fluorescent
protein is accomplished by transfecting circular DNA of the gene of interest with the
DNA of the fluorescent protein, such as green fluorescent protein (GFP), to either the N
or C-terminus of the molecule. This method allows for live cell tracking of the molecule;
however, neither method is ideal for the extracellular domain of the protein of interest in
this study, HB-EGF. Antibody detection is not ideal, as live cell imaging is desirable for
this study, and our previous experience has shown that antibodies to HB-EGF are not
very sensitive and produce high levels of background.
Conjugation of a fluorescent
protein is feasible for HB-EGF, but only for the C-terminal tail as the N-terminus is
cleaved during processing to form mature proHB-EGF. As the extracellular domain can
be detached from the C-terminal intracellular domain via protease cleavage, and both can
serve as signaling molecules, the ability to track both independently is desirable.
In order to visually track the extracellular domain of HB-EGF, we chose the
method of acceptor peptide biotinylation with the enzyme biotin ligase. Biotin ligase is
an E. Coli enzyme that can specifically biotinylate one lysine residue within a fifteen
amino acid acceptor peptide sequence (Figure 2.5. 1a) in the presence of ATP (Beckett et
al., 1999). The acceptor peptide sequence is the minimal substrate required from the
biotin carbonyl carrier protein subunit of acetyl-CoA carboxylase that biotin ligase will
specifically recognize. Once placed within the extracellular region of a protein, biotin
ligase can be used to covalently attach one biotin molecule per protein. Then, as no
extracellular proteins are endogenously biotinylated, these proteins can be specifically
visualized taking advantage of the high binding affinity between streptavidin conjugated
labels and biotin, which can minimize the background signal. Additionally, there is no
need to fix the cells, which gives the flexibility of live-cell imaging. As the C-terminus
of HB-EGF can become separated from the extracellular domain via protease cleavage
and serve as a signaling molecule after translocation to the nucleus, GFP was conjugated
to the C-terminus of HB-EGF to allow for independent tracking of the extracellular
domain versus the C-terminal tail.
The mouse HB-EGF gene was chosen for fusion of the acceptor peptide to keep the
method flexible for future in vivo experiments. The acceptor peptide must be inserted
into an extracellular region of HB-EGF which does not affect the protein's structure or
function. Using domain analysis, four potential insertion sites for the acceptor peptide
were chosen in an effort not affect protein cleavage, glycosylation, or binding to the
receptor and heparan sulfate. Therefore, the following proposed insertion sites are not
within the EGF-like domain or heparin-binding domain.
I. After Asparagine 91: This is two amino acids before the last putative N-terminal
cleavage site predicted from the human gene. The exact site of the mouse gene is
unknown, therefore it was assumed to be similar to the human site. Insertion at
this site would label all size isoforms of HB-EGF produced after N-terminal
cleavage.
II. After Aspartic Acid 106: This site lies between the heparin-binding and EGF-like
domain. However, a very small portion of the heparin-binding domain overlaps
the EGF-like domain, so insertion in this site could alter heparin binding.
III. After Aspartic Acid 63: This site lies just before the last N-terminal cleavage site.
This will label only the largest of the five size isoforms of HB-EGF, as it lies after
the other four protease cleavage sites.
IV. After Threonine 147: This insertion site is two amino acids away from the Cterminal cleavage site for HB-EGF at 149. This region is between the
juxtamembrane stalk and the EGF-like domain, which could affect cleavage of the
protein.
Biotinylation and visualization of acceptor peptide tagged proteins is accomplished
through two different methods depending on the objective of the experiment. The first
method utilizes commercially available biotin ligase. Cells expressing acceptor peptide
tagged proteins are incubated with biotin ligase, ATP, and biotin in a buffer solution,
which biotinylates any acceptor peptide that is expressed extracellularly (Figure 2.5.1 B).
After washing, the cells are incubated with a streptavidin-conjugated fluorophore, and
washed again, then visualized with fluorescence microscopy.
A second, less time
consuming and less expensive method of visualization of the acceptor peptide tag is cotransfection with the DNA for BirA-ER (Figure 2.5.1 C). The BirA-ER plasmid encodes
for biotin ligase fused to an endoplasmic reticulum (ER) localization sequence.
Therefore when a cell expresses both the acceptor peptide tagged protein, and the BirAER protein, the acceptor peptide tag is biotinylated in the endoplasmic reticulum before
presentation on the cell surface utilizing the cells own ATP and biotin supplemented into
the culture media. The cell surface biotinylated acceptor peptide is then visualized after
incubation with a streptavidin-conjugated fluorophore. However, the downside of this
method is a higher degree of cell-cell variability in the amount of acceptor peptide
biotinylated, as it alters with the amount of BirA-ER expressed, which varies depending
on the degree of plasmid uptake and the co-transfection ratio.
The use of streptavidin-conjugated fluorophores to visualize acceptor peptide
biotinylation leads to good labeling with little background.
However, wild-type
streptavidin exists as a tetramer with the ability to bind four biotin molecules. Therefore,
labeling cell surface biotinylated acceptor peptide tagged proteins with wild-type
streptavidin likely leads to crosslinking on the cell surface (Figure 2.5.2 B), which could
affect protein cleavage or endocytosis.
To circumvent this problem, the Ting lab has
engineered a monovalent streptavidin protein that only binds one biotin molecule (Figure
2.5.2 A). This protein consists of one active, wild-type subunit of streptavidin, with three
subunits that are dead in their biotin binding ability. After purification in the Ting lab,
the monovalent streptavidin is labeled with Alexa-Fluor 568 to allow for fluorescent
detection of the marker.
2.2 Materials and Methods
Reagents from Alice Ting's laboratory (MIT)
The BirA-ER plasmid was produced in Alice Ting's laboratory at MIT. The plasmid
encodes biotin ligase (BirA) fused to an endoplasmic reticulum localization sequence.
Monovalent streptavidin fused to Alexa Fluor 568 (mSA-AF568) was produced by Peng
Zou in Alice Ting's laboratory according to the published protocol (Howarth and Ting,
2008). Biotin ligase enzyme was purified in Alice Ting's laboratory at MIT according to
the published protocol (Howarth and Ting, 2008).
Cell culture
COS-7 cells were cultured in DMEM (Gibco 11965) supplemented with 10% FBS and
1%penicillin-streptomycin.
Cell surface acceptor peptide labeling for imaging
COS-7 cells were plated onto 0.1% gelatin coated 35mm Mattek glass bottom dishes at
225,000 cells per dish. After 24 hours, the cells were cotransfected with an acceptor
peptide fusion protein and BirA-ER at 1:1 molar ratios using Fugene6 (Roche) or Mirus
LTI transfection reagent (Mirus) according to the manufacturer's instructions. Media
was supplemented with 10 ptM biotin and incubated for 24 hours. Cells were plated on
ice and washed twice with cold PBS+ (PBS supplemented with calcium and magnesium),
then incubated with 10 tg/mL of monovalent streptavidin-alexa fluor 568 (mSA-AF568)
and 1% pre-dialyzed bovine serum albumin (BSA) for 10 minutes on ice. Cells were
washed twice with PBS+ and imaged in PBS+.
Cell surface acceptor peptide labeling for western blotting
COS-7 cells on tissue culture plastic were washed twice with cold PBS+ (PBS
supplemented with calcium and magnesium)and placed on ice. The cell surface acceptor
peptide was biotinylated with a solution of 0.3 ptM biotin ligase, 1 mM ATP, and 10 pM
biotin in PBS+ for 20 minutes on ice. Cells were washed twice with cold PBS+ and
immediately lysed with RIPA lysis buffer (lx PBS, 1% NP-40, 0.5% sodium
deoxycholate, 0.1% SDS, 1 mM sodium orthovanadate, 0.57 mM PMSF, 1x protease
inhibitor cocktail (Sigma P2714)) on ice for 10 minutes, then scraped to a tube. After gel
electrophoresis and membrane transfer of the lysate, the membrane was blocked for 1
hour in a solution of TBS/T supplemented with 3-5% BSA, then the biotinylated acceptor
peptide was probed with streptavidin-HRP (Molecular Probes S911) at 1:3000 in a
solution of 3-5% BSA for at least 1 hour at room temperature.
Imaging
Phase contrast and fluorescent images were obtained with a digital CCD camera
(CoolSNAP HQ, Roper Scientific) and an inverted microscope (Olympus IX-70). Phase
contrast images were acquired at 30x with a 20x phase-contrast objective (Olympus
LCPlanF NA 0.40) and an additional 1.5x magnification on the microscope. Fluorescent
images were acquired using a 40x water immersion objective (Olympus UApo/340 NA
1.15).
Phase contrast and fluorescent images were aligned manually using Hoescht
stained nuclei (images not shown).
Western blotting
Western blotting was performed with the following antibodies:
anti-phospho-p42/44
MAPK (Cell Signaling Technology 9101S) and goat polyclonal human HB-EGF
antibody (Calbiochem PC319L).
Constructing the mouse HB-EGF acceptor peptide plasmids
The mouse HB-EGF gene was previously inserted into the pShuttle-IRES-hrGFP-1
plasmid (Stratagene, Catalog #240082) by Scott Perkins in Richard Lee's laboratory.
The 15 amino acid acceptor peptide sequence, GLNDIFEAQKIEWHE, represented by
the DNA base pair sequence, GGCCTGAACG ACATCTTCGA AGCCCAGAAA
ATCGAATGGC ACGAA, was inserted into the HB-EGF mouse gene in four different
spots.
The four insertion sites are after amino acids N91, D106, D63, and T147 of
murine HB-EGF. The insertion was done with the Qiagen QuikChange Site-Directed
Mutagenesis Kit (Stratagene, Catalog # 200518), which is designed to change one amino
acid by making a primer complimentary upstream and downstream of the mutation site.
However, it has been reported that the kit can also be used to integrate large fragments at
any site within a plasmid (Geiser et al., 2001). This was accomplished by using primers
with a region of approximately 20 base pairs complimentary to the DNA sequence
upstream of the insertion site, then the 45 base pair acceptor peptide sequence, then
approximately 20 more base pairs complimentary to the plasmid downstream of the
insertion site. The second primer in the mix is the reverse compliment of the above. The
primers used for the N91 insertion site were 5'-GGC CAC CCC AAG CAA AGA AAG
GAA TGG CCT GAA CGA CAT CTT CGA AGC CCA GAA AAT CGA ATG GCA
CGA AGG GAA AAA GAA GAA GAA AGG AAA GGG GTT-3' and its reverse
complement. The primer used for the D105 insertion site were 5'-AGG AAA GGG GTT
AGG GAA GAA GAG AGA CGG CCT GAA CGA CAT CTT CGA AGC CCA GAA
AAT CGA ATG GCA CGA ACC ATG CCT CAG GAA ATA CAA GGA CTA CTG-3'
and its reverse complement. The primers used for the D63 insertion site were 5'-GTG
CTC AGG GGG TCC AGG ACG GCC TGA ACG ACA TCT TCG AAG CCC AGA
AAA TCG AAT GGC ACG AAT TGG AGG GGA CAG ATC TGA ACC TTT TCA-3'
and its reverse complement. The primers used for the T147 insertion site were 5'-GAC
ACA GGT GTC ATG GGC TGA CTG GCC TGA ACG ACA TCT TCG AAG CCC
AGA AAA TCG AAT GGC ACG AAC TAC CAG TGG AGA ATC CCC TAT ACA
CAT ATG A-3' and its reverse complement. All primers were synthesized and PAGE
purified by Integrated DNA Technologies.
The PCR reaction consisted of a 50 IL
volume consisting of 100 ng of the pShuttle-IRES-hrGFP- 1 + HB-EGF (9.4 kb) with 125
ng each of the insertion primers plus 1 piL of PfuUltra Hotstart DNA Polymerase
(Stratagene, Catalog #6003 90), 5 pL of lOx PfuUltra HF Reaction Buffer and 200 piM of
each dNTP from a deoxynucleotide mix (Sigma, Catalog #D-7295). The PCR reaction
consisted of a denaturation step at 95'C for 5 minutes, followed by 18 cycles of 50
seconds at 95"C, 50 seconds at 55'C, and 2 minute/kb template plasmid DNA at 68'C.
The cycle ended with 7 minutes at 68"C. All thermal cycling was performed in capped
PCR tubes in a MJ Research, PTC-200 Peltier Thermal Cycler using the heated lid. The
restriction enzyme Dpn I (New England Biolabs, Catalog #R0176S) was used (1 ptL in 50
ptL PCR sample) to digest methylated template DNA at 374C for >3 hours. The PCR
reaction was transformed into XL10-Gold Ultracompetent Cells using the protocol
present in the Quikchange II XL Site-Directed Mutagenesis Kit (Stratagene, Catalog
#200521), plated onto kanamycin agar dishes and incubated at 370 C overnight.
The
following day colonies were picked and growth in 3 mL of kanamycin supplemented LB
medium with vigorous shaking at 37"C overnight. Media was collected and the plasmid
DNA was purified with the Qiagen QlAprep Spin Miniprep Kit (Qiagen, Catalog
#27104). In order to check for the presence of the acceptor peptide sequence, the purified
DNA was subjected to a PCR reaction with the end primers HB-EGF forward (5'-ATA
TAT ACT AGT ATA TGA AGC TGC TGC CGT C-3') and HB-EGF reverse (5'-ATA
TAT CTC GAG TCA GTG GGA GCT AGC C-3)' with RedTaq DNA Polymerase
(Sigma, Catalog #D-4309).
The PCR thermal cycle consisted of 5 minutes at 94'C,
followed by 30 cycles of 94"C for 30 seconds, 55*C for 30 seconds and 72'C for 1 minute,
with a follow up step of 72*C for 5 minutes.
The product was loaded onto a 1.2%
agarose gel and the plasmids that contained HB-EGF with the acceptor peptide could be
identified by the 45 base pair increase in the DNA fragment size. The positive fragments
were subcloned into the pcDNA3.1N5-His TOPO (Invitrogen, Catalog #K4800-01)
plasmid vector using the TOPO ligation feature according to the manufacturer's
instructions.
The plasmid was then transformed into One Shot TOP10 Chemically
Competent E. Coli cells (Invitrogen, Catalog #C4040-10) and spread onto ampicillin agar
plates and incubated at 370C overnight. The following day colonies were picked and
cultured in 3 mL of LB-medium supplemented with ampicillin and subjected to vigorous
shaking at 37*C overnight. The following day, the plasmid DNA was purified with the
Qiagen QlAprep Spin Miniprep Kit. In order to assay for correct directionality of the
insert, a third PCR reaction was performed with the T7 forward sequencing primer
provided by with the pcDNA3.1 N5-His TOPO plasmid vector and the HB-EGF reverse
primer. Only correct directionality of HB-EGF will result in a PCR product with this
method. The PCR reaction was run on a 1.2% agarose gel to check for bands. Positive
plasmids were sequenced in the forward and reverse direction with the T7 forward primer
and the BGH reverse primer provided with the pcDNA3.1 /V5-His TOPO plasmid vector
by the Brigham and Women's Hospital DNA Core Sequencing Facility (Boston, MA).
Plasmids with the correct sequence were mass produced by culturing transformed
OneShot TOP 10 E. Coli cells in 100 mL ampicillin supplemented LB medium overnight
with vigorous shaking at 37C. The following day, the plasmid DNA was isolated and
purified with the Qiagen Plasmid Maxi Kit (Qiagen, Catalog #12162). The concentration
of the purified DNA was measured with a Smart Spec 3000 Spectrophotometer using
absorbance measurements at 250, 280, and 320 nm.
Constructing the mouse HB-EGF-GFP acceptor peptide plasmid
The AP-HBEGF plasmid at the N91 insertion site was fused to GFP at the C-terminus by
subcloning the AP-HBEGF-AP DNA sequence in pEGFP-N1 (Clontech) using the XhoI
and EcoRI restriction sites.
AP-HBEGF (N91) was amplified from the plasmid in
pcDNA3.1 with primers containing the XhoI and EcoRI restriction sites flanking at the
ends.
The XhoI sense primer contains a two base pair overhang, the 6 base pair XhoI
restriction site, a Kozak sequence, and 23 base pairs complementary to HB-EGF
including the start codon (5'- TAC TCG AGA CCA TGA AGC TGC TGC CGT CGG
TG -3'). The EcoRI reverse primer contains a two base pair overhang, the 6 base pair
EcoRI restriction site, one base pair to put GFP in frame with HB-EGF, and 27 base pairs
complementary to the end of HB-EGF minus the stop codon (5' - ATG AAT TCA GTG
GGA GCT AGC CAC GCC CAA CTT CAC - 3'). The sense and antisense primers
were used at a concentration of 25 pM to amplify the AP-HBEGF(N91) template (10 ng
of template was used in a 25 pL volume). The PCR reaction was run on an agarose gel
and the ~650 base pair AP-HBEGF(N91) product was cut out and purified with a
QAIExII Purification Kit (Qiagen).
The AP-HBEGF(N91) purified fragment and the
pEGFP-N1 vector were double digested with the restriction enzymes XhoI and EcoRI
overnight at 370 C. The restriction enzyme digest reaction was run on an agarose gel and
the cut pEGFP-N 1 vector and AP-HBEGF(N9 1) fragment was gel purified. The cut ends
the pEGFP-N1 vector were dephosphorylated with shrimp alkaline phosphatase for 2
hours are 37C to prevent self-ligation, then the DNA was purified via ethanol
precipitation.
The cut pEGFP-N1 vector and AP-HBEGF(N91) were fused together
using the T4 ligation enzymes for 10 minutes at room temperature, then the reaction was
transformed into GC10 Chemically Competent E. Coli (Genechoice) and plated onto
kanamycin agar dishes for incubation overnight at 370 C. The following day colonies
were picked and tested for correct insertion of the AP-HBEGF(N91) fragment into
pEGFP-N1 by PCR.
Positive colonies were grown overnight in LB Medium with
kanamycin and DNA was purified the following day with the Qiagen Maxiprep Kit.
After further analysis, an error was found in the primer design for this subcloning. The
EcoRI antisense primer encodes a stop codon consisting of the overhang DNA plus the
EcoRI restriction sequence. Therefore, when expressed in cells, AP-HBEGF(N91) was
expressed and correctly biotinylated with biotin ligase, but GFP was not fused to the Cterminal as determined by fluorescent imaging and western blotting.
To correct this
problem, site directed mutagenesis was performed to change a T to an A immediately
after the HB-EGF protein sequence. Two complementary primers were designed that
have 27 base pairs complementary to the end of HB-EGF and 29 base pairs
complementary to the EcoRI restriction site and the following multiple cloning region of
pEGFP-N1 (5'- GTG AAG TTG GGC GTG GCT AGC TCC CAC CGA ATT CTG
CAG TCG ACG GTA CCG CGG GCC -3' and in reverse 5'- GGC CCG CGG TAC CGT
CGA CTG CAG AAT TCG GTG GGA GCT AGC CAC GCC CAA CTT CAC -3'). The
site-directed mutagenesis reaction was completed in the presence of 1 tg of the APHBEGF(N91)-GFP plasmid with a stop codon, 210 ng of each primer, 1 unit of Pfu
HotStart Ultra DNA Polymerase (Stratagene 600390) and 1 ptL of a 10 mM dNTP mix in
a reaction volume of 50 iL. The reaction was denatured at 95'C for 3 minutes, following
by 18 cycles of denaturing at 95*C for 50 seconds, annealing at 60'C for 50 seconds, and
elongation at 68 0 C for 5 minutes and 21 seconds (based on 1 minute/lkb of plasmid
length). The last cycle contained a long elongation step of 7 minutes. The completed
0
PCR reaction was incubated with the restriction enzyme DpnI at 37 C overnight to digest
the original methylated DNA, leaving only the newly formed plasmid generated via PCR
intact, which was used to transform GC10 Competent K Coli cells and plated on
kanamycin agar dishes. Colonies were picked the following day and grown in 3 mL of
LB medium with kanamycin overnight at 37'C with shaking at 225 rpm. Successful
mutagenesis was confirmed with sequencing on DNA purified with the Qiagen Miniprep
Kit. The leftovers from the 3 mL starter culture were used to start a 100 mL culture in
LB medium with kanamycin overnight at 37'C with shaking at 225 rpm, and the APHBEGF(N91)-GFP plasmid was purified from this bacteria with the Qiagen Maxiprep
Kit and the concentration was measured with absorbance.
Purification of HB-EGF
Media was collected from 10 cm dishes of COS-7 cells after 48 hours of transfection with
HB-EGF plasmids.
The media was concentrated via filtration and biotinylated with
biotin ligase in solution. The purified media was subjected to western blotting with
streptavidin-HRP as described above. Cell lysates from AP-CFP-TM transfected COS-7
cells were used as a positive control for streptavidin-HRP binding.
Bands above -32
kDa represent endogenous proteins in the media which bind streptavidin. AP-HBEGF is
represented by a faint band around 28 kDa, with T147 showing the highest level of HBEGF expression in the media.
Note that much of the sample may be lost during
concentration with the centricon filter, as HB-EGF sticks to the filter membrane.
Therefore, western blotting may not accurately represent the amount of HB-EGF present
in the media. However, it is assumed that the relative amounts of HB-EGF between
samples are correct.
Bioactivity assay
COS-7 cells (3.7x106 cells) were cultured in 150 cm2 cell culture flasks in DMEM with
10% fetal bovine serum, and 1%penicillin-streptomycin. The following day, the media
was replaced with DMEM without penicillin-streptomycin and 10 IM biotin, and cells
were co-transfected with AP-HBEGF(N91) and BirA-ER, and BirA-ER alone for the
control.
After twenty-four hours of transfection, the media was supplemented with
sodium chlorate (50 mM) to reduce HSPG sulfation and prevent HB-EGF from binding
to HSPGs rather than being released into the conditioned medium after cleavage. Twenty
four hours later, the media was removed and replaced with 15 mL of PBS with calcium
and magnesium, 200 ig/mL phorbol 12-myristate 13-acetate (PMA) (Sigma P8139) and
0.01% hydrogen peroxide (Sigma 216763), which have been reported to stimulate
cleavage of HB-EGF.
After twenty-four hours incubation with cleavage activators,
conditioned media was collected and combined with DMEM removed the previous day,
then floating cells were removed via centrifugation. A positive control for heparin bead
binding was made, consisting of a solution of 30 mL of PBS+ with 50 mg/mL of
recombinant human HB-EGF (Sigma E4643).
Heparin acrylic beads (H5263) were
washed twice with PBS, and 50 iL of the bead slurry was incubated with the conditioned
medium and positive control on a rotator at 4'C for >8hours. Beads were washed with
PBS+ and HB-EGF was eluted with a 2 M NaCl solution at 37"C for 1 hour. The control
eluant, biotinylated AP-HBEGF and positive control eluant were run on a gel and probed
with streptavidin-HRP to detect the biotinylated form of the protein to confirm the
presence of AP-HBEGF.
To test the activity of the AP-HBEGF heparin bead eluant
versus the control eluant, serum starved confluent monolayers of naYve COS-7 cells were
treated with 0, 0.01, 0.1, 1 and 10 iL of the heparin bead eluant, with and without
pretreatment for 30 minutes with 10 pM of the EGFR tyrosine kinase inhibitor AG1478.
Cells were lysed after 15 minutes of stimulation and lysates were probed via western
blotting for ERK phosphorylation.
2.3 Results
The fifteen amino acid acceptor peptide sequence was successfully inserted into
the mouse HB-EGF gene at four different locations: after asparagine 91 (N91), aspartic
acid 106 (D106), aspartic acid 63 (D63), and threonine 147 (T147). All four AP-HBEGF
constructs, including HB-EGF without the AP tag were successfully expressed in COS-7
cells and recognized by a polyclonal human HB-EGF antibody (Figure 2.5.3 A). The
fifteen amino acid insertion caused a shift in mobility, with the acceptor peptide HB-EGF
samples appearing slightly larger than the wild-type HBEGF. Additionally, HB-EGF
was not endogenously expressed at levels high enough to be detected via western blot in
COS-7 cells, as shown by the GFP transfected and wild-type, non-transfected controls
(Figure 2.5.3 A). The AP-HBEGF construct existed as multiple bands, denoting the
multiple N-terminal cleavage sites which results in five different HB-EGF size isoforms.
The protein ran at an approximate molecular weight of 22 kDa; however, the positive
control, which is recombinant human HB-EGF ran at approximately 9 kDa.
This
observation has been reported in the literature, and is attributed to heavy 0-glycosylation
of the eukaryotic synthesized protein causing a large shift in electrophoretic mobility.
Additionally, the AP-HBEGF is expressed in the pro-form (uncleaved), while the
recombinant protein is only the mature, soluble extracellular domain.
The AP-HBEGF constructs expressed in COS-7 cells were biotinylated on the cell
surface with exogenous biotin ligase, which does not penetrate the plasma membrane.
After biotinylation the cells were lysed and subjected to SDS PAGE gel electrophoresis.
The membrane was then probed for biotinylated proteins with streptavidin-HRP (Figure
2.5.3 B). The upper bands represent endogenously biotinylated proteins that exist inside
the cell, as the sample transfected with HB-EGF without the acceptor peptide has
identical positive bands. Two of the four constructs show successful biotinylation of APHBEGF (N91 and D63) indicating that the construct was correctly incorporated into the
plasma membrane and expressed on the cell surface of COS-7 cells.
The D63 AP-
HBEGF construct is representative of only one isoform of the five HB-EGF different size
isoforms, while the N91 constructs shows all five HB-EGF bands. The positive control
in this experiment is a construct engineered in Alice Ting's lab in the MIT chemistry
department. This construct consists of an extracellular cyan fluorescent protein molecule
and an acceptor peptide tag fused to the transmembrane domain of the PDGF receptor
(AP-CFP-TM) (Chen et al., 2005). The D106 construct had no detectable cell surface
biotinylation. It is speculated that the acceptor peptide inserted after D106 is sterically
unavailable to biotin ligase as it lies immediately after the highly structured EGF-like
domain and before the heparin-binding domain. The T147 construct has the acceptor
peptide inserted after the juxtamembrane stalk of HB-EGF, therefore increasing the
length of the pro-HBEGF stalk. One study reported that increasing the stalk length of
HB-EGF increased the cleavage efficiency by TACE/ADAM17 (Hinkle et al., 2004).
Additionally, a blot of media from COS-7 cells transfected with the various HB-EGF
plasmids had a high level of biotinylated T147 in the media compared to N9 1, D106 and
D63 (Figure 2.5.3 C). Therefore, we speculate that the lengthened stalk of HB-EGF in
the T147 construct leads to increased cleavage so that it is present in undetectable
amounts on the cell surface and accumulates in the media. Interestingly, the different
size isoforms of HB-EGF identified by a HB-EGF polyclonal antibody versus cell surface
biotinylation of the acceptor peptide tag differ. Biotinylated cell surface HB-EGF N91
shows the presence of a strong band at approximately 14 kDa (Figure 2.5.3 B), which is
not observed with the HB-EGF antibody (Figure 2.5.3 A). This suggests that the 14 kDa
band is not recognized by the HB-EGF antibody, perhaps due removal of the recognition
epitope. As D106 did not get biotinylated, T147 caused altered cleavage rates, and D63
did not label all size isoforms of HB-EGF, N91 was chosen as the best construct with
properties closest to wild-type HB-EGF.
As the C-terminus of HB-EGF can be detached from the extracellular domain via
protease cleavage, a method to independently track the C-terminus from the extracellular
domain would allow for independent tracking and assessment of cleavage. The N91 APHBEGF gene was cloned in the plasmid vector pEGFP-N1 to attach GFP to the Cterminal tail of proHB-EGF, which is referred to here as AP-HBEGF-GFP (Figure 2.5.4
A). To assure that the acceptor peptide, and biotinylation of the acceptor peptide does
not alter the protein's activity, the AP-HBEGF extracellular domain was tested for EGFR
bioactivity. Biotinylated AP-HBEGF was precipitated from the conditioned medium of
COS-7 cells with heparin acrylic beads, and a control heparin bead eluant was also
prepared from conditioned medium of COS-7 cells that were not transfected with APHBEGF.
Both heparin bead eluants were added to naYve COS-7 cells in increasing
dosages, lysed and assayed for phospho-ERK.
The biotinylated AP-HBEGF heparin
bead eluant activated phospho-ERK at levels much higher than the control bead eluant
(Figure 2.5.4 B). Additionally, this activation was inhibited by preincubation of the cells
with the EGFR tyrosine kinase inhibitor, AG1478, suggesting that the ERK activation is
indeed mediated by the EGFR. Therefore, biotinylated AP-HBEGF has the ability to
activate phospho-ERK through the EGFR in a paracrine fashion. As the biotinylated APHBEGF protein was purified by heparin beads, this additionally demonstrates that
insertion of the acceptor peptide after N91 does not interfere with HB-EGF's ability to
bind heparin.
Transfection of this construct into COS-7 cells and imaging of the extracellular
acceptor peptide tag shows that the C-terminal tail (GFP, green) and the extracellular
domain (AF568, red) signals overlapped, suggesting HB-EGF was primarily in the proform (Figure 2.5.5). Additionally, epifluorescence imaging shows that HB-EGF was
localized primarily to sites of cell-cell contact in a confluent monolayer of COS-7 cells
(Figure 2.5.5 A).
This fact is highlighted when the cells were sparsely plated, as the
majority of HB-EGF concentrated only at the cell-cell junction rather than at the free
edges (Figure 2.5.5 B).
2.4 Discussion
Biotin ligase and acceptor peptide labeling proved to be a viable technique for
HB-EGF, with little background noise. The technique is powerful for the case of HBEGF to be able to distinguish the cell surface pool of HB-EGF from that in intracellular
compartments.
Additionally, with the dual labeling technique presented here for
independent tracking of the C-terminal tail with EGFP and the extracellular domain with
the acceptor peptide tag, proteolytic cleavage of HB-EGF could be visualized due to
differential localization of the fluorescent signals.
However, HB-EGF shows no
difference in the extracellular and intracellular tags, indicating that the protein is
primarily uncleaved, which is validated via western blotting.
Utilization of the co-
transfection method with BirA-ER makes the protocol cost sensitive, as no purified
recombinant biotin ligase is necessary.
This technique allows for live cell tracking of the HB-EGF ligand, as cell fixation
is not required with the acceptor peptide labeling method.
The utilization of an
engineered monovalent streptavidin prevents streptavidin from cross-linking biotinylated
proteins on the cell surface. However, the large size of monomeric streptavidin tetramers
(53 kDa) may inhibit native processing of the protein, such as endocytosis or cleavage.
Additionally, biotinylation of HB-EGF at the acceptor peptide tag does not affect the
activity of the protein, however, a streptavidin complexed to the biotinylated acceptor
peptide tag could introduce steric effects preventing HB-EGF from activating the
receptor, even though the acceptor peptide tag is sufficiently far away from the EGF-like
domain.
2.5 Figures
AF568
Biotin
G LN DIF EAQ K I EW HE
Acceptor Peptide (AP)
c
b
B
B
4BirAadded
AP
B
BirAER
Figure 2.5.1:
Two methods of BirA (biotin ligase) labeling of the
acceptor peptide protein. (A) The enzyme BirA (biotin ligase) covalently attaches
a biotin to one lysine residue within the fifteen amino acid acceptor peptide sequence.
The acceptor peptide is then visualized by the addition of a streptavidin-conjugated
fluorophore, such as Alexa-Fluor (AF568) (B) The acceptor peptide tagged protein is
produced in the endoplasmic reticulum (ER), and then presented on the cell surface.
Exogenous BirA (biotin ligase) enzyme is added to the cell media along with ATP and
biotin, leading to biotinylation of only the extracellular fraction of the acceptor peptide
tagged protein. The biotinylated acceptor peptide tag can then be visualized with a
streptavidin-fluorophore conjugate. (C) The acceptor peptide tagged protein is coproduced along with the construct BirA-ER, which encodes for biotin ligase with an
endoplasmic reticulum (ER) localization sequence. Acceptor peptide tagged proteins are
biotinylated with BirA-ER in the endoplasmic reticulum (ER) using the cells own
intracellular ATP and biotin supplemented in the culture media. The acceptor peptide
tagged protein is presented on the cell surface already biotinylated, and the biotinylated
acceptor peptide can be visualized with a streptavidin-fluorophore conjugate.
(Figure2.5.1B&C adaptedfrom Howarth et al, 2008 (Howarthand Ting, 2008))
.........................................................................
. ................
. . ........
--
-
.........
...........
. .......
BI jB Engineer_,
Wild-type
streptavidin (A4)
Kd4.4 x 10-14 M
Monovalent
streptavidin
(A1D3)
Kd4.8
Divalent
antibody
Wild-type
streptavidin
x10-14
M
Monovalent
streptavidin
Out
Cell surface protein
Figure 2.5.2.
Design of a monovalent streptavidin.
(A) Wild-type
streptavidin is a tetramer with four biotin [B] binding sites [A4]. The Ting lab has
engineered streptavidin that is dead in the biotin binding pocket [gray, D]. Monovalent
streptavidin is engineered by combining three subunits of mutant streptavidin unable to
bind biotin with one of the wild-type subunits fused to a His-Tag [AlD3], which is able
to retain low Kd values. (B) Divalent antibodies and wild-type streptavidin cross-link
proteins on the cell surface with their ability to bind multiple proteins. Monovalent
streptavidin is designed to reduce protein cross-linking on the cell surface as it can only
bind one biotinylated protein. (Figureadaptedfrom Howarth et al, 2008 (Howarth and
Ting, 2008))
. .
.
..
....
..
. ......
. ........
0
4/
0k
49
-
38
-
28
-
14
-
0
0
(0
39-
4.
C
2814-
28-
Figure 2.5.3 Validation of AP-HBEGF constructs.
(A) Expression of all
four AP-HBEGF plasmids named by their insertion site (N91, D106, D63, T147), and
wild-type HB-EGF in COS-7 cells was assessed by western blotting with a human HBEGF polyclonal antibody. GFP serves as a transfection control, wild-type (WT)
represents non-treated COS-7 cells, and the positive control represents recombinant
human HB-EGF. (B) Cell surface biotinylation of AP-HBEGF was successful only in the
N91 and D63 constructs. The extracellular acceptor peptide was biotinylated in COS-7
cells expressing the AP-HBEGF plasmids, then lysed and detected with streptavidin-HRP
after gel electrophoresis. N91 shows biotinylation of several size isoforms of HB-EGF
between 14 and 34 kDa, and D63 shows biotinylation of the largest size isoform around
34 kDa. No detectable biotinylation is observed for D106 or T147. The positive control
represents AP-CFP-TM showing biotinylation at 42 kDa. The two large molecular
weight bands and the one at 36 kDa represent endogenously biotinylated proteins, as they
are also present in the HB-EGF (no acceptor peptide) control. (C) The T147 protein is
enriched in the cell media in the large molecular weight form at around 28 kDa.
However, little is detected in the media for N9 1, D106 and D63.
..............
.............
AP
a
N:SH
PRO
.......
-- .. ..............
GFP
D
EGC
soluble HB-EGF
pro HB-EGF
Heparin Bead Eluant
Control
HB-EGF-Biotin
0 10-210-1 1 10 0 1 21 11 10
b
Phospho-ERK
Phospho-ERK
+AG1478
Figure 2.5.4 Final gene map of AP-HBEGF-GFP and bioactivity. (A)
Schematic representation of the final mouse HB-EGF construct (AP-HBEGF-GFP) with
the acceptor peptide inserted after N91. The arrows show sites of modification, where
EGFP was inserted at the C-terminus, and the acceptor peptide was inserted after N91.
Triangles represent sites of cleavage at the N and C-terminus of the human protein. The
gene is broken up into the secretory region (SEC), the pro-region (PRO), the heparinbinding domain (HBD), the EGF-like domain (EGF), and a transmembrane domain. (B)
Bioactivity of the soluble, mature form of biotinylated AP-HBEGF-GFP was assessed by
a concentration dependent phosphorylation of ERK after 15 minutes incubation with the
roughly purified protein eluted from heparin beads compared to a control eluant. The
numbers above the blot represent the volume of heparin bead eluant in microliters that
cells were stimulated with, as the concentration of HB-EGF here is unknown. The
biotinylated HB-EGF activates ERK at lower concentrations than the control eluant.
Additionally, most of the ERK activation at lower concentrations is through the EGFR, as
preincubation of naive COS-7 cells for 30 minutes with 10 pM AG1478, an EGFR kinase
inhibitor, blocked activation of ERK with the heparin bead eluant.
b
Figure 2.5.5 AP-HBEGF-GFP is localized to cell-cell contact sites. (A)
After 24 hours of co-transfection of AP-HBEGF-GFP with BirA-ER in COS-7 cells, the
biotinylated, extracellular acceptor peptide in AP-HBEGF-GFP was labeled with
monovalent streptavidin-alexa fluor 568 (red, left) and imaged alongside the cytoplasmic
tail conjugated to EGFP (green, middle), and phase contrast (right) in a confluent
monolayer and (B) in sparsely plated cells. Arrows show localization of HB-EGF to sites
of cell-cell contact. Each row represents the same field. Scale bars are 40 pm.
40
Chapter Three: Polarization of HB-EGF at the wound edge
3.1 Introduction
HB-EGF is a chemotactic factor for many cell types, as activation of the EGFR
often leads to an increase in migration.
However, HB-EGF specifically has been
identified as an autocrine factor released by keratinocytes (Shirakata et al., 2005;
Tokumaru et al., 2000) and comeal epithelial cells (Block et al., 2004; Xu et al., 2004)
upon wounding that is crucial to activate cell migration to drive wound closure. The
direct stimuli that lead to HB-EGF shedding varies depending on the system, however
ATP released after wounding (Yin and Yu, 2009), or simply the increase in available area
for cell migration can lead to cleavage of the pro-form of HB-EGF to produce the soluble
form for autocrine signaling (Block et al., 2004).
It has been hypothesized that the EGF-EGFR system has potential for spatial
localization of the autocrine signaling loop (Maheshwari et al., 2001).
The
Lauffenburger lab has computationally predicted that spatially localized autocrine
signaling is possible (Maly et al., 2004) and has produced experimental data of human
mammary epithelial cell migration which suggests a spatially orientated autocrine loop
may exist and drive persistent cell migration (Maheshwari et al., 2001).
The
computational kinetic model of EGFR autocrine signaling includes ligand shedding,
activation of the EGFR, recruitment of Grb2 and Sos, and phosphorylation of Raf, MEK,
and ERK within one cell. In addition, the model includes positive feed back, where ERK
activates additional ligand shedding as well as negative feedback where ERK
phosphorylates Sos, rendering it unable to participate in the EGFR-Grb2-Sos complex.
This model predicted three stables states of autocrine signaling: no signaling, symmetric
EGFR signaling and polarized EGFR signaling. In the polarized state, EGF release as
well as ERK activation is concentrated to one pole of the cell. The stable state which the
system adopts was dependent on the density of cell surface receptors; densities of
approximately 400 to 800 receptors/ptm 2 resulted in polarized signaling.
Experimental evidence for a spatially polarized autocrine loop was discovered in
pro-EGF expressing human mammary epithelial cells, which had increased directional
persistence (Maheshwari et al., 2001).
In this study, cell lines were created
overexpressing the pro-membrane bound form of EGF (EGF-Ct) and a form of EGF that
contains the soluble portion only without the transmembrane stalk (sEGF).
EGF
expression in both forms (sEGF & EGF-Ct) was shown to increase migration speed in all
samples, however only pro-EGF expression led to an increase in persistence time and
path length. Interestingly, the addition of exogenous HB-EGF to these cells abrogated
the increased directional persistence. Therefore, it was hypothesized that the increased
directional persistence was due to an asymmetrical autocrine signaling loop, leading the
cell to migrate in the direction of EGFR activation.
An additional quantitative experimental study of ligand capture in the EGF-EGFR
system in the Lauffenburger lab outlines conditions that must be met for autocrine, rather
than paracrine signaling (DeWitt et al., 2001). In this study, the rate of EGF ligand
production was altered using a tet-off expression system, metalloprotease inhibitors were
utilized to block the amount of ligand released, and an EGFR blocking antibody
(mAb225) was used to vary the number of receptors accessible on the cell surface. This
study showed that the amount of ligand captured by the cell that released it was
dependent on the ratio of the ligand production rate (VLT) to the receptor production rate
At VLT/VR values of 0.3 or less, the system was dominantly autocrine, and
captured the majority of ligand released. However as the ratio increased, more ligand
(VR)-
was released into the bulk medium. EGF binds to the EGFR only; however HB-EGF
binds not only to the receptors, but to HSPGs, which are plentiful on the cell surface.
Therefore, as the number of potential binding sites on the cell surface for HB-EGF is
much higher than that of EGF, one would predict HB-EGF signaling to be predominantly
autocrine in nature. As HB-EGF serves as an autocrine chemotactic ligand during wound
healing, and the EGFR system may have the ability to produce spatially polarized
autocrine signaling, we sought to test the hypothesis that spatially polarized HB-EGF
autocrine signaling loops exist and drive persistent cell migration to close wounds.
3.2 Materials and Methods
Materials
Batimastat (BB-94) was a generous gift from Steve Wiley's lab at Pacific Northwest
National Laboratory where it was custom synthesized by Kimia Corporation (Santa Clara,
CA). The EGFR blocking antibody (mAb225) was isolated from a hybridoma cell line
obtained from the American Type Culture Collection (Gill et al., 1984). We additionally
used the following: GM6001 (Calbiochem 364206), 2R-[(4-biphenylsulfonyl)amino]-Nhydroxy-3-phenylpropionamide (BiPS) (Calbiochem 444249), PD98059 (Calbiochem
513000), AG1478 (Calbiochem 658552), PP2 (Calbiochem 529576). Adenoviruses for
wild-type dynamin-1, wild-type dynamin-2, dynamin-1 S45N, dynamin-2 1690K, and
tTA were a generous gift from the Schmid laboratory (Soulet et al., 2006).
Wound healing assay
Mattek 35 mm glass bottom dishes were coated with 0.1% gelatin for > 1 hour at 370 C.
COS-7 cells were plated in a confluent monolayer at 225,000 cells per dish. Wounds
were induced by dragging a P200 pipette tip with constant velocity over the cell
monolayer.
Cell surface labeling with mSA-AF568 was performed as described in
section 2.2 with gentle washing, as not to disrupt cells on the edge of the wound. In the
dual-labeling experiment streptavidin-fluorescein was utilized (Vector Laboratories SA5001)
Plasmids
For all experiments in this chapter and on, the HB-EGF construct has the AP insertion
after amino acid N91 in murine HB-EGF, which is referred to as AP-HBEGF-GFP. The
construct AP-CFP-TM consists of an extracellular acceptor peptide, cyan fluorescent
protein, and the transmembrane domain of the PDGFR (Chen et al., 2005).
This
construct was made in Alice Ting's laboratory, MIT Chemistry Department to serve as a
positive control for acceptor peptide biotinylation and visualization.
3.3 Results
3.3.1 ProHB-EGF is missing from the wound edge
In order to track the localization of HB-EGF during cell migration, we chose the
wound healing assay. This assay is a convenient method to measure and stimulate cell
migration with the advantage of a known direction of cell migration toward wound
closure without time lapse imaging, as is required for single migrating cells. A wound is
induced by scratching a confluent monolayer of cells with a P200 pipette tip, then the
remaining cells proliferate and migrate in to close the free area of the wound, typically
over timescales of 12 to 48 hours.
COS-7 cells are rather stationary and migrate
minimally until stimulated by creation of a wound. In order to track HB-EGF during
wound healing, the AP-HBEGF-GFP
construct was transfected into confluent
monolayers of COS-7 cells. After twenty-four hours a wound was induced in the APHBEGF-GFP expressing cells and the wound was allowed to heal for four hours, then the
extracellular acceptor peptide tag was labeled with mSA-AF568. It was observed that
AP-HBEGF-GFP was absent from free or leading edge of cells on the wound edge
(Figure 3.5.1 A).
However, cells within the monolayer maintained a symmetrical
distribution of HB-EGF at cell-cell contact sites (data not shown).
Additionally, the
control construct AP-CFP-TM does not change localization at the wound edge after four
hours of wound healing when compared to cells in the monolayer (Figure 3.5.2 B). As
AP-CFP-TM is still detectable at the leading edge, it is not likely that thinning of the cell,
and therefore less fluorescence at the lamellipod is the cause of the loss of HB-EGF.
We hypothesized that the loss of HB-EGF at the leading edge of the wound was
due to proteolytic cleavage of the extracellular domain from the leading edge and fast
internalization of the cleaved C-terminal domain. Additionally, we hypothesized that this
spatially localized proteolytic cleavage led to EGFR activation at the leading edge which
may lead to chemotactic cell migration in the direction of would closure.
3.3.2 HB-EGF loss from wound edge is not due to proteolytic release
In order to test this hypothesis, we utilized many reagents to block the loss of HBEGF from the leading edge after wounding, internalization of the C-terminal tail, and
other intracellular signaling nodes that may cause positive or negative feedback in the
cascade. First, reagents aimed at blocking protease cleavage of HB-EGF at the leading
edge were utilized, including batimastat (BB-94), GM6001 and BIPS. Batimastat is a
broad spectrum matrix metalloproteinase (MMP) inhibitor that inhibits MMP activity by
binding the zinc ion in the activate site of the MMP and has been shown to inhibit
ovarian and breast cancer tumor growth (Davies et al., 1993; Low et al., 1996).
Batimastat has been demonstrated to block cleavage of EGF ligands, including HB-EGF
(Prenzel et al., 1999; Sahin et al., 2004). GM6001 is a broad spectrum hydroxamic acid
inhibitor of matrix metalloproteases that also interferes with the zinc binding site of
MMPs and has been experimentally shown to inhibit proHB-EGF cleavage (Armant et al.,
2006). BiPS and TAPI-2 are also broad spectrum matrix metalloproteinase inhibitors,
and BiPs has been shown to inhibit HB-EGF cleavage in COS-7 cells specifically
(Mifune et al., 2005). Cells were pretreated with the metalloproteinase inhibitors for one
hour, then a wound was induced and the wound was allowed to close for 4 hours in the
presence of the inhibitors.
No change in HB-EGF localization was observed in the
presence of metalloproteinase inhibitors batimastat (10 pM), GM6001 (10 pM), BiPs (10
pM), or TAPI-2 (20 pM) (Figure 3.5.2 B-E), suggesting that ligand cleavage is not
responsible for the loss of HB-EGF at the leading edge of cells at the edge of the wound.
To test if loss of HB-EGF from the wound edge was due to cleavage induced by
positive feedback through EGFR signaling or downstream mediators, agents aimed at
blocking parts of this pathway were targeted. First, directly blocking the EGFR was
tested with AG1478 and mAb225. AG1478 is a tyrosine kinase inhibitor for the EGFR
that blocks downstream intracellular signaling triggered by the EGFR. mAb225 is a
mouse monoclonal antibody that binds to the EGFR and blocks ligands from binding and
activating the receptor, therefore inhibiting downstream signaling (Gill et al., 1984).
Pretreatment of monolayers of COS-7 cells with 10 piM AG1478 or 10 ptg/mL of
mAb225 for one hour prior to induction of the wound, and continuous incubation during
four hours of wound healing did not reduce the loss of HB-EGF at the wound edge
(Figure 3.5.2 F-G). To investigate other pathways downstream of EGFR, monolayers of
COS-7 cells were pretreated for one hour prior to induction of the wound with 25 piM of
the MEK1 inhibitor PD98059 that blocks activation of ERK, 20 pM of the protein kinase
C (PKC) inhibitor rottlerin, 10 pM of the PKC inhibitor bisindolylmaleimide I (BIM I),
100 nM of the phosphatidylinositol-3 kinase (P13K) inhibitor wortmannin, 10 pM of the
Src inhibitor PP2, and 10 ptg/mL of the P13K inhibitor LY294002. However, none of the
inhibitors stopped loss of HB-EGF from the wound edge (Figure 3.5.2 H-M). BIM I
treatment did lead to poor labeling of the extracellular acceptor peptide tag and spotting
in all cells, even those that were non-transfected. Likely BIM I was cytotoxic and led to
leaky plasma membranes, allowing streptavidin to permeate the cell and bind to
endogenous biotin in the mitochondria; or BIM I caused large amounts of HB-EGF
cleavage and the extracellular domain remained on the cell surface in a spotted pattern.
As the loss of HB-EGF from the leading edge is not likely due to ligand cleavage
or EGFR activation, it was hypothesized that extension of the lamellipodia may push
proHB-EGF to the rear of the cells. In order to inhibit cytoskeletal processes during
wound healing, actin polymerization was blocked with 200 nM cytochalasin D, 10 pM of
the microtubule stabilizer taxol, and 300 nM of the microtubule polymerizing inhibitor
nocodazole. However, none of these agents inhibited the loss of HB-EGF at the edge of
the wound (Figure 3.5.2 N-P).
Therefore, action of the cytoskeleton during wound
healing is unlikely to control the loss of HB-EGF at the wound edge.
3.3.3 Newly synthesized HB-EGF localizes to cell-cell contact sites only
As protease inhibitors did not stop the loss of HB-EGF at the wound edge, it is
likely that the entire pro-form of HB-EGF is removed from the leading edge, with the
extracellular domain still attached to the C-terminal domain. One possible mechanism
for loss of the pro-form of the protein could be internalization from the leading edge via
endocytosis. In order to block internalization, the temperature was lowered to 4"C, which
blocks internalization of cell surface proteins. Preincubation of COS-7 cells at 4"C for
one hour prior to induction of the wound, then continuous incubation of the cells for the
entire four hours of wound healing did inhibit loss of HB-EGF at the wound edge and
extension of lamellipodia into the wound area (Figure 3.5.2 R). However, lowering the
cell temperature to 4'C likely inhibits many processes, including incorporation of new
protein into the plasma membrane. Therefore, it could be that only newly synthesized
protein localizes away from the wound edge, and the protein at the leading edge is turned
over at an average rate. Recycling of all proteins was blocked with the chemical inhibitor
monensin at 10 [M. Monensin did not lead to complete inhibition of the loss of HB-EGF
at the wound edge, however, it was reduced (Figure 3.5.2
Q),
suggesting that newly
synthesized or recycled proHB-EGF is localized preferentially at sites of cell-cell contact
rather than at the leading edge.
The chemical inhibitor of endocytosis, phenyl arsine
oxide (PAO), was utilized, however was toxic to COS-7 cells at concentrations required
for endocytosis inhibition.
To block the clathrin-mediated endocytosis pathway more specifically, dominant
negative mutants in dynamin were employed. The S45N and 1690K mutants in dynamin1 and dynamin-2, respectively, in adenovirus forms were transfected into COS-7 cells
(Soulet et al., 2006). However, inhibition of endocytosis, as assessed by EGF induced
EGFR downregulation, was never achieved with these constructs in confluent
monolayers of COS-7 cells. These constructs were communicated to work inefficiently
in confluent monolayers of cells (unpublished data), however, confluency is required for
wound healing experiments. Therefore, endocytosis inhibition was never achieved here
and the role of clathrin mediated endocytosis at the wound edge could not be tested.
In order to investigate the turnover of cell surface HB-EGF during the wound
healing experiment, a pulse-chase experiment was performed. A confluent monolayer of
COS-7 cells was co-transfected with AP-HBEGF (without the GFP tag on the
cytoplasmic tail) and BirA-ER and cultured for 24 hours with biotin supplemented in the
media. Cells were washed and the extracellular biotinylated acceptor peptide tag on HBEGF was labeled with mSA-AF568 (red), which does not allow protein cross-linking.
Cells were placed back in full media with serum supplemented with biotin, and wounds
were produced. After four hours of wound healing, the cell surface pool of HB-EGF was
labeled with streptavidin-fluorescein (green), and the cells were immediately imaged
(Figure 3.5.3). The cell surface fraction of AP-HBEGF labeled before production of the
wound was primarily localized inside the cell (Figure 3.5.3 A). However, the cell surface
pool of AP-HBEGF detected after four hours showed localization to sites of cell-cell
contact and was missing from the wound edge, as expected (Figure 3.5.3 B). As the bulk
of the mSA-AF568 signal was internal, this suggests that newly synthesized HB-EGF
becomes preferentially localized to sites of cell-cell contact rather than at the wound edge
over the experimental timecourse of four hours.
3.4 Discussion
In summary, the discovery that HB-EGF was missing from the wound edge in
migrating monolayers of COS-7 cells led us to hypothesize that this loss was due to
preferential proteolytic cleavage from the leading edge.
However, this hypothesis is
likely not correct for the COS-7 cell line, as inhibition of several points along this
pathway did not prevent loss of HB-EGF from the wound edge. Rather it appears that
most of the HB-EGF present on the cell surface at the time of induction of the wound is
internalized over the experimental time course of four hours. Rather newly synthesized
or recycled HB-EGF is localized preferentially at sites of cell-cell contact and not at the
edge of the wound. ProHB-EGF could be preferentially deposited at sites of cell-cell
contact, or could be incorporated uniformly in the plasma membrane, then diffuse to and
stay at cell-cell contact sites.
However, this raises the question:
what molecular
interactions govern proHB-EGF localization at sites of cell-cell contact?
The known
protein interactions of HB-EGF include the EGFR, ErbB4, HSPGs, and CD9 on the
extracellular domain, and BAG-1 for the intracellular domain. As BAG-1 does not have
any discovered cytoskeletal binding properties, we hypothesize that an interaction
between the extracellular domain of HB-EGF and one of its binding partners on
neighboring cells holds HB-EGF at cell-cell contact sites. This ensuing hypothesis is
pursued in Chapter 4.
-- -
- - -----
3.5 Figures
(A)
Figure 3.5.1
HB-EGF is absent from the wound edge.
After 24h of co-
transfection of (A) AP-HBEGF-GFP or (B) AP-CFP-TM with BirA-ER in COS-7 cells,
the monolayer was wounded with a P200 pipette tip. After four hours of wound healing,
the biotinylated, extracellular acceptor peptide was labeled with monovalent streptavidinalexa fluor 568 (red, left) and imaged alongside (A) EGFP (green, middle) or (B) CFP
(cyan, middle), and phase contrast (right). Note that the cells were pre-treated with
Hoescht to label nuclei blue to assist with image alignment between the fluorescent
images and the phase contrast images, as they were taken with different objectives, then
compensated for scale during image processing. Therefore the presence of the light cyan
nuclei in (B) is due to Hoescht blue fluorescence overlap into the cyan channel. Only the
large brightly labeled cell in the middle is AP-CFP-TM transfected. Each row represents
the same field. Scale bars are 40 pm.
.......
..
.......................
..........
(A) Control
(B) Batimastat
(C) GM6001
(D) BiPs
(E) TAPI-2
(F) AG1478
(G) mAb225
(H) PD98059
(1 Rottlerm
(J) BIM
(K) Wortmaninin
(L) PP2
(M)LY294002
N) Cytochalasin D
(Q) Monensin
(R) 4C
I
O0 Nocodazolo
P) Taxol
Figure 3.5.2 Effect of inhibitors on polarization of HB-EGF at wound
edge. After four hours of wound healing, the extracellular domain of HB-EGF is
missing from the wound edge in COS-7 cells transfected with AP-HBEGF in the (A)
untreated control. Pretreatment with inhibitors for 1 hour prior to wounding and
continuous incubation with inhibitors after induction of the wound did not prevent the
loss of HB-EGF at the leading edge of cells at the would edge (B-M). Polarization of
HB-EGF at the wound edge was tested after incubation with the protease inhibitors (B)
batimastat (10 iM), (C) GM6001 (10 p.M), (D) BiPs (10 piM), and (E) TAPI-2 (20 pM).
EGFR signaling was inhibited with (F) AG1478 (10 pM) and (G) mAb225 (10 Ig/mL).
Intracellular signaling pathways were inhibited with (H) PD98059 (25 pM), (I) rottlerin
(20 pM), (J) BIM 1 (10 pM), (K) wortmannin (1 iM), (L) PP2 (10 pM), (M) LY294002
(10 ig/mL). Cytoskeletal components were inhibited with (N) cytochalasin D (200 nM),
(0) nocodazole (300 nM), and (P) taxol (10 pM). Protein recycling was blocked with
(Q) monensin (10 ptM) and (R) endocytosis was inhibited by lowering the incubation
temperature to 4C. The dotted line represents the wound edge with empty space on the
right side. Scale bar represents 20 im.
AP Taa Blotinviation
(A
(mSA-AF568)
AP Taa Biotinviation (SA-Fluoresceini
(c)
(B)
Hoescht
Figure 3.5.3 Turnover of cell surface HB-EGF at wound edge. Confluent
monolayers of COS-7 cells were co-transfected with AP-HBEGF (without the GFP tag
on the cytoplasmic tail) and BirA-ER and cultured for 24 hours with biotin supplemented
in the media. Cells were washed and the extracellular biotinylated acceptor peptide tag
on HB-EGF was labeled with monovalent-streptavidin-AF568 (mSA-AF568, red) in (A),
then cells were placed back in full media and a wound was induced with a pipette tip.
After four hours of wound healing, the extracellular biotinylated acceptor peptide tag on
HB-EGF was labeled again, this time with streptavidin-fluorescein (SA-fluorescein,
green) in (B), the nuclei were stained with Hoescht (C), and then cells were immediately
imaged. After four hours of wound healing most of the cell surface HB-EGF was
internalized into the cell (red), and newly synthesized HB-EGF exists at sites of cell-cell
contact (green). The dotted line represents the wound edge. Scale bar represents 40 jim.
52
Chapter Four:
The heparin-binding domain mediates
localization of proHB-EGF to cell-cell contact sites
4.1 Introduction
Heparan sulfate proteoglycans (HSPGs), present on the cell surface and in the
extracellular matrix, are capable of binding many growth factors.
A traditionally
proposed purpose of this interaction is to restrain a soluble ligand to the cell surface and
increase the local concentration to activate a receptor (Schlessinger et al., 1995). Most
ligands reported to interact with HSPGs are soluble secreted factors, such as fibroblast
growth factors (Gospodarowicz et al., 1984; Maciag et al., 1984; Shing et al., 1984),
vascular endothelial growth factor (Ferrara and Henzel, 1989), hepatocyte growth factor
(Nakamura et al., 1984; Zhou et al., 1999) and platelet-derived growth factor (Schilling et
al., 1998). However, HSPGs also interact with a few growth factors that are anchored to
the cell surface via a transmembrane domain, particularly those that belong to the
epidermal
growth
factor receptor
(EGFR)
ligand
family,
including
HB-EGF
(Higashiyama et al., 1991), amphiregulin (Cook et al., 1991), betacellulin (Shing et al.,
1993), and certain isoforms of neuregulin (Holmes et al., 1992; Loeb and Fischbach,
1995). All ligands in the EGFR family have the ability to activate their receptors in the
diffusible
form
produced
after
proteolytic
release
from
the
cell
surface
(autocrine/paracrine signaling). However, of the EGFR family ligands, only HB-EGF
(Higashiyama et al., 1995), amphiregulin (Inui et al., 1997), transforming growth factor-a
(Anklesaria et al., 1990) and betacellulin (Tada et al., 1999) have been reported to
activate their receptors in the pro-form, while anchored to the membrane before cleavage
(juxtacrine signaling). Autocrine/paracrine signaling with these ligands has been studied
much more extensively than juxtacrine signaling; however, the majority of HB-EGF
remains on the cell surface in the pro-form at sites of cell-cell contact (Goishi et al.,
1995). Therefore, here we focus on the role of HSPG binding to the pro-form of HBEGF capable of juxtacrine signaling.
4.2 Materials and Methods
Constructing heparin-binding domain mutants of AP-HBEGF-GFP
Four constructs with various mutations to the heparin-binding domain of AP-HBEGFGFP were produced. The region of the heparin-binding domain which is not included
within the EGF-like domain (#93-105) was deleted (AP-delHBD-HBEGF-GFP), the first
five lysine residues of the heparin-binding domain (#93-97) were mutated to alanine (AP97A-HBEGF-GFP), all lysine and arginine residues in the heparin-binding domain which
lies outside of the EGF-like domain (93-105) were mutated to alanine (AP-105AHBEGF-GFP), and finally all lysine and arginine residues in the entire heparin-binding
domain (93-113), including the portion that is included in the EGF-like domain, were
mutated to alanine (AP-1 13A-HBEGF-GFP).
The AP-delHBD-HBEGF-GFP mutant
was constructed using a primer complementary to AP-HBEGF-GFP for 20 amino acids
before amino acid 93 and after amino acid 105 (5' - AAA TCG AAT GGC ACG AAG
GGG ACC CAT GCC TCA GGA AAT A -3') and its reverse complement. In order to
make AP-97A-HBEGF-GFP, the primer HBD-A(93-97)F (5'- AAA TCG AAT GGC
ACG AAG GGG CGG CAG CCG CTG CGG GAA AGG GGT TAG GGA AGA AGA 3') and its reverse complement were designed to be complementary to AP-HBEGF-GFP
for 20 and 22 amino acids before and after the region of the heparin-binding domain to be
mutated (93-97) with a five residue stretch of alanine to replace the five lysine residues.
The AP-105A-HBEGF-GFP plasmid was made based on the AP-HBEGF-GFP plasmid
and using a primer, ALA-HBD-F, (5'- AAA TCG AAT GGC ACG AAG GGG CGG
CAG CCG CTG CGG GAG CAG GGT TAG GGG CGG CAG CCG ACC CAT GCC
TCA GGA AAT A-3') and its reverse complement that is complementary before amino
acid 93 and after 105 for twenty amino acids.
The amino acid sequence
AAAAAGAGLGAAA, as a replacement for KKKKKGKGLGKKR, was in between the
complementary regions.
The AP-1 13A-HBEGF-GFP plasmid was constructed by
mutating amino acids RKYK in stretch 110-113 in the EGF-like domain of AP- 105AHBEGF-GFP to AAYA.
The primer, RKA1 13 SEM Forward, (5'- CAG CCG ACC
CAT GCC TCG CAG CGT ACG CAG ACT ACT GCA TCC ACG GGG A -3') has 17
base pairs complementary to AP- 105A-HBEGF-GFP before amino acid 110 and 20 base
pairs after 113 with an the sequence for AAYA in between. Using the above primers and
base plasmids, site-directed mutagenesis via whole plasmid PCR was completed with an
annealing temperature of 50'C and an elongation time of 14 minutes. The PCR reaction
was digested with DpnI restriction enzyme for >6 hours, transformed into XL- 10 Gold E.
Coli, and plated onto kanamycin agar dishes. The following day, colonies were picked
and placed into 3 mL of LB Media with kanamycin and cultured on a shaker overnight at
225 rpm and 37*C. The following day, 1.5 mL of the bacteria culture was used to purify
the plasmid with the Qiagen Miniprep Kit. Miniprep samples were sent for sequencing to
screen colonies for the correct mutation. Positive cultures were expanded into 100 mL of
LB Media with kanamycin, and large amounts of the plasmid were purified with the
Qiagen Maxiprep Kit.
Constructing AP-Amphiregulin-GFP plasmid
The human amphiregulin DNA coding sequence in pBM-IRESpuro was a generous gift
from Steve Wiley. Amphiregulin DNA was removed from pBM-IRESpuro and cloned
into pEGFP-N1. First, human amphiregulin was amplified by PCR from pBM-IRESpuro
with the forward primer AR-XhoI-fwd (5'- TAC TCG AGA TGA GAG CCC CGC T -3'),
which contains an overhang with the XhoI restriction enzyme recognition sequence, and
the reverse primer AR-EcoRl-rev (5'- ATG AAT TCT TGC TAT AGC ATG TAC ATT
TCC ATT CTC TTG -3'), which contains an overhang for the EcoRI restriction enzyme.
The PCR fragment was gel purified with the Qiaquick gel purification kit. The PCR
fragment and the pEGFP-N1 plasmid backbone were subjected to restriction enzyme
digestion with XhoI and pEGFP-N1 at 37'C for >1hour. The digested samples were run
on a gel, bands corresponding to the amphiregulin digested PCR product and linearized
pEGFP-N1 were cut out and gel purified. The PCR fragment was ligated into linearized
pEGFP-N1 with T4 DNA ligase. This was transformed via T4 ligation into E. Coli and
plated onto kanamycin agar dishes, and incubated overnight at 370 C. The following day,
>5 colonies were picked into 3 mL of LB media with kanamycin and incubated overnight
at 37'C with shaking at 225 rpm. DNA was purified from 1-2 mL of bacterial culture
with the Qiagen Miniprep Kit. Insertion of amphiregulin was confirmed via sequencing
with above forward primer. Positive colonies were expanded into 10OmL of LB media
with kanamycin overnight at 37"C at 225 rpm shaking. Large amounts of plasmid DNA
were purified with the Qiagen maxiprep kit. The fifteen amino acid acceptor peptide
sequence was then inserted into the AR-GFP plasmid after valine 107 in amphiregulin.
This was achieved by whole plasmid PCR with the forward primer AR-APV107 Fwd (5'TCA GTC AGA GTT GAA CAG GTA GTT GGC CTG AAC GAC ATC TTC GAA
GCC CAG AAA ATC GAA TGG CAC GAA AAG CCC CCC CAA AAC AAG -3') and
its reverse complement, AR-APV107 Rev (5'- CTT GTT TTG GGG GGG CTT TTC
GTG CCA TTC GAT TTT CTG GGC TTC GAA GAT GTC GTT CAG GCC AAC
TAC CTG TTC AAC TCT GAC TGA -3'). The forward primer consists of the first 24
base pairs of the human amphiregulin sequence before the insertion site after V107, the
fifteen amino acid acceptor peptide DNA sequence, then 18 more base pairs analogous to
amphiregulin after V107.
PCR was performed with 18 rounds, and an annealing
temperature of 50'C, and an extension temperature of 68*C for 12 minutes per round.
The reaction was digested with the DpnI restriction enzyme, then transformed into XL 10
Gold Cells (Stratagene) and plated onto kanamycin plates. The following day positive
colonies were selected into 3 mL of LB media with kanamycin and incubated overnight
at 37'C at 225 rpm shaking. The following day, DNA was purified with the Qiagen
Miniprep kit and sent for sequencing to check for correct insertion of the acceptor peptide
with the AP-XhoI-Fwd primer. Colonies positive for the acceptor peptide were grown
into 100 mL of LB media with kanamycin and large amounts of plasmid DNA was
purified with the Qiagen maxiprep kit.
Constructing the heparin-binding domain mutant of amphiregulin (AP-143AAmphiregulin-GFP)
In order to mutate all basic lysine and arginine residues to neutral alanine in the heparinbinding domain of human amphiregulin in AP-AR-GFP (corresponding to residues 123-
143 in the human amphiregulin gene sequence) (KPKRKKKGGKNGKNRRNRKKK
mutated to APAAAAAGGANGANAANAAAA),
AP-AR-GFP was subjected to whole
plasmid PCR with the forward primer (AR-HBD-R/KtoA Fwd: 5'- AAG ACG GAA
AGT GAA AAT ACT TCA GAT GCT CCC GCT GCT GCT GCT GCT GGA GGC
GCT AAT GGA GCT AAT GCT GCT AAC GCT GCT GCT GCT AAT CCA TGT
AAT GCA GAA TTT CAA AAT TTC T -3') and its reverse complement (AR-HBDR/KtoA Rev: 5'- AGA AAT TTT GAA ATT CTG CAT TAC ATG GAT TAG CAG
CAG CAG CGT TAG CAG CAT TAG CTC CAT TAG CGC CTC CAG CAG CAG
CAG CAG CGG GAG CAT CTG AAG TAT TTT CAC TTT CCG TCT T -3'). The
mutagenesis primer consists of 27 base pairs of the human amphiregulin sequence before
the heparin-binding domain, then the DNA sequence for the mutated heparin-binding
domain, and then 31 base pairs of amphiregulin after the heparin-binding domain. PCR
with these primers replaced the wild-type heparin-binding domain with the mutated
sequence. PCR was performed with 18 cycles of PCR at an annealing temperature of
504C and an extension temperature of 68'C for 12 minutes per round. The reaction was
digested with the DpnI restriction enzyme, then transformed into XL10 Gold Cells
(Stratagene) and plated onto kanamycin plates. The following day positive colonies were
selected into 3 mL of LB media with kanamycin and incubated overnight at 370 C at 225
rpm shaking. The following day, DNA was purified with the Qiagen Miniprep kit and
sent for sequencing to check for mutation of the heparin-binding domain with the APXhoI-Fwd primer. Positive colonies were expanded into 100 mL of LB media with
kanamycin and large amounts of plasmid DNA were purified with the Qiagen maxiprep
kit.
Addition of the heparin-binding domain to AP-CFP-TM
The heparin-binding domain of mouse HB-EGF, consisting of the portion that lies
outside the EGF-like domain (#93-105), was added to the AP-CFP-TM control protein
between the acceptor peptide and the CFP molecule. The heparin-binding domain was
inserted after a three amino acid linker before CFP, then another three amino acid linker
was inserted after the heparin-binding domain. Therefore the sequence consists of the
acceptor peptide, the linker Gly-Ala-Pro, the heparin-binding domain, the linker Ala-Gly-
Gly, then CFP. This was accomplished with the primer CFPAP-HBD-F (5'- AGT GGC
ACG AGG GCG CGC CGA AAA AGA AGA AGA AAG GAA AGG GGT TAG GGA
AGA AGA GAG CGG GCG GCA TGG TGA GCA AGG GCG AGG A -3') and its
reverse complement, whose first and last twenty amino acids are complementary to APCFP-TM. The PCR reaction was run under the same conditions as the AP-HBEGF-GFP
plasmid, digested with the DpnL restriction enzyme, transformed into XL-10 Gold Cells
(Stratagene) and plated on ampicillin agar dishes. Colonies were picked 24 hours after
plating and grown in 3 mL of LB media with ampicillin overnight with shaking at 225
rpm at 37"C. The following day, 1 mL of the culture was harvested and DNA was
purified using the Qiagen Miniprep kit. The purified DNA was sent for sequencing with
the T7 forward sequencing primer to verify correct insertion of the heparin-binding
domain. After identifying a positive colony, this colony was allowed to grow overnight
in 100 mL of LB Media and ampicillin shaking at 225 rpm. The following day, the
bacteria was harvested and purified with the Qiagen Maxiprep Kit. After further analysis
of the structure of cyan fluorescent protein, I noticed that the N-terminus of the protein
actually ends at the same side of the protein as the C-terminus. This means that the
protein is likely oriented so that the acceptor peptide and the heparin-binding domain are
pointing down toward the membrane rather than out into the media. In order to correct
this, I chose to insert a flexible protease-resistant linker of 20 amino acids. The linker
sequence was designed based on a study of linker sequences and protease resistance
(Robinson and Sauer, 1998). The length was set at 20 residues, resulting in a linker of
approximately 80 angstroms to reach around the CFP molecule (-50 angstroms). I chose
to build off of the AGG sequence already present in the 3 amino acid linker and follow it
with SEGGGSEGGTSGATG.
The insert was made with a PCR reaction on the AP-
HBD-CFP-TM plasmid with the following forward CFPAPHBD Linker Insert Forward
Primer (5'- GAA GAA GAG AGC GGG CGG CTC TGA AGG CGG CGG CAG CGA
AGG CGG CAC CAG CGG CGC GAC CGG AAT GGT GAG CAA GGG CGA GGA
-
3') and its reverse complement, which contains 19 and 20 amino acids complementary to
the AP-HBD-CFP-TM before and after the linker insertion site, respectively.
The
methods for inserting the linker are identical to those presented in this section above for
insertion of the heparin-binding domain.
4.3 Results
4.3.1 Heparin and heparan sulfate alter the localization of proHB-EGF
As proHB-EGF was observed primarily at sites of cell-cell contact, the question
arose of what molecular interactions between cells may lead to proHB-EGF
concentration in this area. As the extracellular domain of proHB-EGF has the ability to
interact with HSPGs, which are present on the cell surface, we hypothesized that this
interaction may control localization of HB-EGF to cell-cell contact sites. To test the
hypothesis, we sought to compete for HSPG binding to proHB-EGF with exogenous
heparin and heparan sulfate. Heparin and heparan sulfate (100 pig/mL) dramatically
changed the localization of AP-HBEGF-GFP (Figure 4.5.1 A&B).
The extracellular
(AF568) and intracellular (GFP) domain of AP-HBEGF-GFP changed from localization
primarily at cell-cell contact sites to a homogenous distribution over the entire cell
surface.
However, the addition of the glycosaminoglycan chondroitin sulfate (100
ig/mL) did not affect the localization of AP-HBEGF-GFP (Figure 4.5.1 C). Images
shown are after four hours of treatment; however changes in proHB-EGF localization
were observed as soon as five minutes after the addition of heparin (Figure 4.5.2). The
timecourse depicted in figure 4.5.2 illustrates a highlighted area of cell-cell contact where
proHB-EGF is localized to cell-cell contact sites. Heparin (100 ig/mL) was added after
the first frame (0 seconds), and after only 20 seconds the intensity of HB-EGF was
reduced at cell-cell contact sites. The intensity of HB-EGF here continued to decrease,
and was observed to be primarily diffusely localized throughout the membrane of the
transfected cell after 300 seconds of heparin treatment. To analyze this in more detail,
the fluorescence intensity per pixel along the vertical line graphed in figure 4.5.2 B is
mapped over time after the addition of heparin. The fluorescence intensity at the cell-cell
contact site was at maximum before the addition of heparin.
This peak intensity
decreased over time, and reached its half-maximal value at approximately five minutes
post heparin addition.
After the removal of heparin for 24 hours, AP-HBEGF-GFP
localized back to sites of cell-cell contact as observed before the addition of heparin (data
not shown).
4.3.2 ProHB-EGF in microdomains at cell-cell contact sites is inaccessible to BirA
and streptavidin.
The amount of HB-EGF on the cell surface can be distinguished from the total
pool of HB-EGF by biotinylating the cell surface fraction with exogenously added biotin
ligase. After cell lysis and gel electrophoresis, the biotinylated protein can be detected
with streptavidin-HRP. In this experiment, it was observed that pretreatment of the APHBEGF-GFP expressing COS-7 cells with 100 ptg/mL of heparin in the media led to a
larger amount of biotinylated AP-HBEGF on the cell surface (Figure 4.5.3 A). It was
therefore hypothesized that heparin increased the cell surface fraction of HB-EGF by
increasing the translocation of HB-EGF from the cytoplasm to the plasma membrane, or
stabilized HB-EGF on the cell surface, slowing the protein's cell surface half life.
However, further study showed that the heparin-induced increase in the biotinylated cell
surface fraction was not time dependent, and even at times as little as 1 minute showed
no change in the amount of HB-EGF on the cell surface than at 60 minutes (data not
shown). Additionally, preincubation for 30 minutes and continued treatment throughout
the experiment with 10 pM monensin or at 4'C had no effect on the amount of
biotinylated HB-EGF on the cell surface after the addition of heparin. Monensin is a
carboxylic acid ionophore that disrupts the transport of membrane vesicles from the golgi
complex to the plasma membrane (Stein et al., 1984) and lowering the temperature to 4'C
inhibits endocytosis and protein trafficking. As neither of these pretreatments stopped
the heparin-induced increase of HB-EGF on the cell surface, it is unlikely that heparin is
affecting the translocation of HB-EGF to the cell surface. It was additionally observed in
fluorescent microscope imaging of biotinylated AP-HBEGF that although the majority of
the GFP signal overlapped the extracellular mSA-AF568 signal, there were some areas
that were unlabeled (Figure 4.5.3 B). These areas are typically flat planar-like structures
where two cells overlap. The initial hypothesis was that HB-EGF was cleaved in these
areas and only the C-terminal tail was left, however, western blotting with a GFP
antibody for total HB-EGF shows no evidence of a C-terminal tail fragment (data not
shown). Additionally, these patch-like structures were highly mobile upon the addition
of heparin (data not shown), suggesting that HB-EGF is in the pro-form and interacting
with HSPGs. Therefore, it is assumed that these patch-like areas are tightly closed and
sterically unavailable to biotin ligase for biotinylation or mSA-AF568 to label the prebiotinylated protein. However, upon the addition of heparin, HB-EGF is released from
HSPGs and cell-cell contact sites, and accessibility to the acceptor peptide is unhindered.
4.3.3 ProHB-EGF interaction with HSPGs controls cell-cell contact site localization
The change in HB-EGF localization after the addition of heparin away from cellcell contact sites suggests that interaction with HSPGs was responsible for localizing HBEGF to this area. To further test this hypothesis, we aimed to diminish HSPG-proHBEGF interactions by two alternative means: (a) removing HSPGs, and (b) altering the
proHB-EGF sequence.
Removal of HSPGs by two different methods changed the
localization of AP-HBEGF-GFP from cell-cell contact sites to a homogenous distribution
over the cell surface, as seen with the addition of heparin. Sodium chlorate is an inhibitor
of protein sulfation that blocks the enzyme ATP-sulfurylase (Baeuerle & Huttner, 1986).
Culture of COS-7 cells expressing AP-HBEGF-GFP with 50 mM sodium chlorate for 24
hours led to a homogeneous distribution of AP-HBEGF-GFP over the cell surface (Figure
4.5.4 A). Additionally, digestion of cell surface heparan sulfate with heparinase III (1.6
mU/mL) for four hours similarly changed the localization of AP-HBEGF-GFP to a
homogenous distribution over the cell surface, although some AP-HBEGF-GFP remained
at cell-cell contact sites (Figure 4.5.4 B). Thus, the reduction of all sulfation with sodium
chlorate, or the removal of cell surface heparan sulfate with heparinase III reduced the
amount of AP-HBEGF-GFP at cell-cell contact sites, leading to a more homogenous
distribution over the cell surface. It may be that heparan sulfate chains bound to HB-EGF
are relatively protected from degradation by heparinase, as this is the case with bFGF and
mammalian heparanase (Tumova & Bame, 1997).
For a converse experiment, we made four different mutants of AP-HBEGF-GFP
to inhibit the ability of HB-EGF to interact with heparan sulfate (Table 1). The heparinbinding domain of HB-EGF consists of amino acids 93-113 of the mouse protein, which
contains a combination of 12 basic lysine and arginine residues, which have been shown
to be responsible for the HB-EGF interaction with heparin (Thompson et al., 1994).
However, residues 108-113 lie within the EGF-like domain and contain three of the basic
amino acids. Deletion of the portion of the heparin-binding domain that lies outside the
EGF-like
domain (93-105)
(AP-delHBD-HBEGF-GFP)
led to
a homogeneous
distribution of HB-EGF over the cell surface (Figure 4.5.5 A), similar to that seen with
the addition of heparin. In contrast, mutation of the first five positive lysine residues (9397) in the heparin-binding domain to non-polar alanine (AP-97A-HBEGF-GFP) had no
effect on the localization of HB-EGF (Figure 4.5.5 B). Mutation of an additional four
lysine and arginine
residues to alanine (93-105)
(AP-105A-HBEGF-GFP)
was
nonetheless sufficient to change the localization of AP-HBEGF-GFP to be homogenously
spread over the cell surface (Figure 4.5.5 C). The same result was observed with the
mutation of all 12 basic amino acids in the heparin-binding domain to alanine, including
those within the EGF-like domain (AP-1 13A-HBEGF-GFP) (Figure 4.5.5 D). These data
suggest that binding of HB-EGF via the heparin-binding domain to HSPGs on the cell
surface is required for HB-EGF localization to cell-cell contact sites.
Name
AP-HBEGF-GFP
AP-delHBD-HBEGF-GFP
AP-97A-HBEGF-GFP
AP-105A-HBEGF-GFP
AP-113A-HBEGF-GFP
Heparin-Binding Domain Sequence
KKKKKGKGLGKKRDPCLRKYK
--------------------------- DPCLRKYK
AAAAAGKGLGKKRDPCLRKYK
AAAAAGAGLGAAADPCLRKYK
AAAAAGAGLGAAADPCLAAYA
# Basic AAs
12
3
7
3
0
Table 1. Mutations in the heparin-binding domain of HB-EGF. Mutations of basic lysine and arginine
residues to alanine in the twenty-one amino acid heparin-binding domain of murine HB-EGF, with the
unaltered heparin-binding domain listed in AP-HBEGF-GFP with the number of basic amino acids left
after deletion. A '-' represents amino acid deletion, rather than mutation.
4.3.4 ProHB-EGF interacts with HSPGs in trans
To further investigate the role of HSPGs on the localization of HB-EGF to cellcell contact sites, wild-type CHO-K1 cells and mutant CHOpgsD-677 cells, which are
deficient in heparan sulfate production were utilized (Lidholt et al., 1992). Confluent
monolayers of each cell type were transfected for 24 hours with AP-HBEGF-GFP and the
localization of HB-EGF was evaluated with the GFP-tagged cytoplasmic tail. The wildtype CHO-K1 cells, which have functional HSPGs, show localization of HB-EGF to sites
of cell-cell contact (Figure 4.5.6 A), as observed with COS-7 cells.
However,
transfection of AP-HBEGF-GFP into the CHOpgsD-677 cells that lack heparan sulfate
showed no HB-EGF localization to cell-cell contact sites (Figure 4.5.6 C). As observed
in the COS-7 cells, AP-1 13A-HBEGF-GFP was diffusely localized over the cell, and not
concentrated at sites of cell-cell contact in both CHO-Ki (Figure 4.5.6 B) and CHOpgsD677 cells (Figure 4.5.6 D). Despite localization of AP-HBEGF-GFP to cell-cell contact
sites in CHO-Ki cells, large flattened areas of the membrane with strong planes of
proHB-EGF concentration (as shown in Figure 4.5.3 B) was not observed in this cell line.
Therefore, this type of localization may be controlled by a second interaction partner in
addition to HSPGs that is present in the COS-7 cells and not the CHO-Ki cells.
Demonstrating that HSPGs were required for HB-EGF localization to cell-cell
contact sites raised the question of the source of HSPGs for this interaction. HB-EGF
could interact in cis with HSPGs on the same cell that expresses HB-EGF, holding it at
cell-cell contact sites. Alternatively, HB-EGF may interact with HSPGs in trans, and
therefore HSPGs would be required on the neighboring cell for localization to cell-cell
contact sites. To distinguish between these two possibilities, CHO-Ki (wild-type) and
CHOpgsD-677 cells were transfected with AP-HBEGF-GFP (green) or mCherry (red)
and co-cultured together in different combinations. The positive control (Figure 4.5.7 A)
showed AP-HBEGF-GFP at sites of cell-cell contact between mCherry and AP-HBEGFGFP transfected wild-type CHO-Ki cells.
Additionally, the negative control (Figure
4.5.7 D) showed no cell-cell contact localization of AP-HBEGF-GFP between mCherry
and AP-HBEGF-GFP transfected CHOpgsD-677 cells. However, CHOpgsD-677 cells
transfected with AP-HBEGF-GFP showed localization of HB-EGF to the cell-cell contact
sites with a wild-type CHO-Ki mCherry transfected neighbor (Figure 4.5.7 B). Wildtype CHO-Ki cells transfected with AP-HBEGF-GFP showed no HB-EGF localization
to sites of cell-cell contact with mCherry transfected CHOpgsD-677 neighbors (Figure
4.5.7 C). These data demonstrate that the localization of proHB-EGF to sites of cell-cell
contact is dependent on interaction with HSPGs on a neighboring cell only; therefore proHBEGF interacts with HSPGs in trans.
As transient transfection is utilized to express AP-HBEGF-GFP in COS-7 cells,
not all of the cells in the monolayer take in the plasmid and express the construct.
Therefore, there is a heterogeneous mix of AP-HBEGF-GFP transfected and non-
transfected cells on the dish in these experiments.
It was observed that HB-EGF
localization to cell-cell contact sites it strongest between a transfected and a nontransfected cell (Figure 4.5.8). The GFP-tagged cytoplasmic tail of HB-EGF is observed
to be missing, or weakly localized to cell-cell contact sites between two transfected cells,
as highlighted by the white arrows. It is hypothesized here that proHB-EGF competition
with HSPGs on neighboring cells leads to less HB-EGF at cell-cell contact sites when
two neighboring cells both overexpress AP-HBEGF-GFP.
However, it is unclear
whether this is due to overexpression of the protein, or if this could happen at physiologic
expression levels. However, if this is a natural phenomenon, it may suggest that strong
localization of HB-EGF to sites of cell-cell contact is reserved for non-homogenous
tissues where HB-EGF expression is not equal between two different cells types.
Another hypothesis here is that HB-EGF is excluded from sites of cell-cell contact during
mitosis and continues to be excluded after return to GO/G1. As shown in Figure 4.5.8 C,
HB-EGF is missing from the area that will become the site of cell-cell contact between
the cells in the late phase of mitosis, with the nuclei almost completely separated.
It
should be noted here that many studies on the localization of HB-EGF are done in stably
transfected cell lines, where all cells are expressing HB-EGF, and the localization of HBEGF to cell-cell contact sites in these experiments is likely diminished compared to the
studies here with heterogeneous HB-EGF expression.
4.3.5 Heparin-binding controls amphiregulin localization, but engineered heparinbinding is insufficient for cell-cell contact localization
HB-EGF is not the only ligand in the EGFR family capable of heparin-binding, as
amphiregulin, betacellulin and some isoforms of neuregulin also interact with HSPGs.
HB-EGF and amphiregulin have similar heparin-binding domains located at the Nterminus of the protein after the EGF-like domain. These domains are both twenty-one
amino acids in length, with over half of the residues represented by basic lysine or
arginine, which allow the domain to interact with negatively charged heparin and heparan
sulfate (Thompson et al., 1994).
We sought to test whether heparin changed the
localization of amphiregulin to see if this mechanism was common for other heparinbinding EGFR ligands.
Acceptor peptide labeling of the extracellular portion of
amphiregulin, and imaging of the C-terminal GFP tag, showed that amphiregulin is
similar to HB-EGF in that the extracellular and intracellular fluorophores were colocalized, suggesting that amphiregulin is primarily present in the pro-form (Figure 4.5.9
A). Additionally, amphiregulin was concentrated around the perimeter of the cell at sites
of cell-cell contact.
The addition of heparin (100
tg/mL) for four hours led to a
homogenous distribution of amphiregulin over the cell surface, as seen with HB-EGF
(Figure 4.5.9 B). Therefore, it is likely that the heparin-binding domain of amphiregulin
is also responsible for its localization to sites of cell-cell contact. The heparin-binding
domain of amphiregulin is very similar to HB-EGF, consisting of a 21 amino acid region
of
which
14
of
the
amino
(KPKRKKKGGKNGKNRRNRKKK).
acids
are
basic
lysine
or
arginine
In order to stop the ability of amphiregulin to
interact with HSPGs, all of the lysine and arginine residues were mutated to neutral
alanine (APAAAAAGGANGANAANAAAA).
The heparin-binding domain mutant of
amphiregulin similarly led to less amphiregulin at sites of cell-cell contact (Figure 4.5.9
C) as observed after the addition of heparin. This suggests that the same interaction of
HSPGs in trans with proHB-EGF to localize the protein to sites of cell-cell contact
applies for pro-amphiregulin.
In order to test whether the heparin-binding domain of HB-EGF was sufficient to
localize any transmembrane protein to cell-cell contact sites, an engineered construct was
made consisting of an extracellular acceptor peptide sequence, followed by the portion of
the heparin-binding domain of HB-EGF which lies outside the EGF-like domain
(residues 93-105), cyan fluorescent protein, then the transmembrane domain of the PDGF
receptor.
This construct (AP-HBD-CFP-TM) (Figure 4.5.10 B) showed the same
localization as the control construct (Figure 4.5.10 A) without a heparin-binding domain
or flexible linker (AP-CFP-TM), with a homogenous localization over the cell surface.
Therefore, the heparin-binding domain alone was insufficient to localize any protein to
cell-cell contact sites. This suggests that an additional co-factor which interacts with HBEGF and amphiregulin may be required for HSPG-mediated localization to cell-cell
contact sites, or removal of proteins unassociated with HSPGs, or a particular
conformation of the heparin-binding domain is required that is not present in AP-HBDCFP-TM.
4.4 Discussion
The key finding of Chapter 4 is that a trans interaction between HSPG and the
HB-EGF heparin-binding domain is responsible for localizing proHB-EGF to sites of
cell-cell contact. HSPGs change the localization of pro-amphiregulin in a similar manner,
suggesting that this may be a common mechanism for transmembrane heparin-binding
ligands of the EGFR family. As HB-EGF (Higashiyama et al., 1995), amphiregulin (Inui
et al., 1997), and betacellulin (Tada et al., 1999) are heparin-binding and capable of
signaling in juxtacrine mode, and the heparin-binding domain of HB-EGF and
amphiregulin inhibits binding to the receptor until in complex with heparin or heparan
sulfate (Higashiyama et al., 1993; Johnson and Wong, 1994; Piepkorn et al., 1994;
Takazaki et al., 2004), the role of this interaction may be to assist in holding prospective
juxtacrine ligands at sites of cell-cell contact, likely in concert with CD9, to bind a
receptor on a neighboring cell.
HB-EGF is localized to sites of cell-cell contact in the two cell lines explored in
this chapter which express HSPGs, COS-7 and CHO-Ki cells. However, COS-7 cells, in
addition to localizing HB-EGF at sites of cell-cell contact, have proHB-EGF localized in
large planar areas (Figure 4.5.3 B) that are not accessible to streptavidin or BirA,
suggesting that they are tightly sealed junctions, sterically inaccessible to exogenously
added proteins.
These structures were quickly dissolved by the addition of heparin,
however, they were not observed in the CHO-Ki cell line. These structures are very
similar to tetraspanin enhanced microdomains observed by Singethan and colleagues
(Singethan et al., 2008) after treatment with a CD9 clustering antibody. Therefore, we
hypothesize that proHB-EGF transfection may have the ability to cluster CD9 into
tetraspanin microdomains at cell-cell contact sites along with proHB-EGF-HSPGs, as
proHB-EGF interacts with CD9 via its heparin-binding domain to the second
extracellular loop of CD9 (Sakuma et al., 1997). Additionally, heparin may have the
ability to dissipate CD9 from cell-cell contact sites, as it does for proHB-EGF. Evidence
to support this hypothesis is the fact that the CD9 clustering antibody (K41) binds the
same region of CD9 that HB-EGF interacts with (Singethan et al., 2008), and these
structures are not present in CHO-Ki cells that have been demonstrated to have very little
to no CD9 expression (Jennings et al., 1994).
The interaction of HSPGs with pro-HB-EGF in trans was demonstrated with wildtype and mutant CHO cells which lack heparan sulfate (Figure 4.5.7). The localization of
HB-EGF to cell-cell contact sites is unlikely to depend on interaction with EGFR, as
CHO-Ki cells lack endogenous EGFR, yet still show strong localization of HB-EGF to
cell-cell contact sites. This experiment also suggests that HSPGs are not required as an
intracellular chaperone for proHB-EGF during transport to the cell surface, as cells
lacking heparan sulfate were still able to localize HB-EGF to sites of cell-cell contact
when neighbored with a HSPG producing cell. HSPGs have previously been shown to
interact in trans with VEGFR-2 in VEGFR-mediated angiogenesis (Jakobsson et al.,
2006) and the Xenopus receptor caALK4 via the co-factor Vgl during mesoderm
migration in early left-right development (Kramer and Yost, 2002). Additionally, this
interaction likely plays a role in blastocyst implantation during pregnancy, as the
interaction between HB-EGF on the luminal epithelium with EGFR and HSPGs on the
adjacent blastocyst are required for successful attachment, which is reduced by
exogenous heparin or blastocyst heparinase treatment (Farach et al., 1987; Farach et al.,
1988; Raab et al., 1996). The trans interaction also likely plays a role in diphtheria toxin
infection, as the presence of heparan sulfate is required for diphtheria toxin binding to
proHB-EGF (Shishido et al., 1995).
The localization of proHB-EGF to sites of cell-cell contact sites between two APHBEGF-GFP transfected cells is much weaker than observed between a transfected and a
non-transfected cell (Figure 4.5.8). We speculate that this is due to competition between
the heparin-binding domain of proHB-EGF and HSPGs at the cell-cell junction between
two transfected cells. This would suggest that strong localization of HB-EGF to cell-cell
contact sites may be reserved for junctions between two different cell populations, with
only one expressing proHB-EGF. Given HB-EGF's role in embryonic development and
that proHB-EGF juxtacrine EGFR activation can be growth inhibitory, this may serve as
a signal for tissue polarization and boundary formation.
In the kidney, HB-EGF is localized to the basolateral surface of collecting tubule
cells, and is absent from the apical surface of collecting tubules, where cell-cell contact is
missing from the basolateral surface in contact with urine (MacRae Dell et al., 2004).
This localization is similar to what we observed in wounded COS-7 cells, which are also
kidney cells from the African green monkey (Figure 3.5.1A). However, in mouse models
of autosomal-recessive polycystic kidney disease, HB-EGF is mis-localized in the
collecting tubules, with increased expression and localization to the apical surface in
addition to the basolateral surface (MacRae Dell et al., 2004). Additionally, EGFRs are
mis-localized to the apical surface of the related autosomal dominant autosomal-recessive
kidney disease (Du and Wilson, 1995). Polycystic kidney disease is not well understood,
but is a genetic disease leading to renal failure associated with over-proliferation of
epithelial cells and is linked to a defect in cilia and polarization of cells in the collecting
tubules. Mis-localized proHB-EGF may contribute to or be a consequence of loss of cell
polarity in polycystic kidney disease, or contribute to epithelial cell proliferation. The
change in localization of HB-EGF in the collecting tubules from healthy mice compared
to mouse models of autosomal-recessive polycystic kidney disease is reminiscent of the
change in localization observed with the addition of heparin in COS-7 cells (Figure 4.5.1
and Figure 4.5.2).
4.5 Figures
Extracellular HB-EGF
AP Tag Blotinylation (mSA-AF568)
C-Terminal HB-EGF Tail
EGFP
Phase Contrast
(A)
Heparin
(B)
Heparan
Sulfate
(C)
Chondroitin
Sulfate
Figure 4.5.1. Heparin and heparan sulfate changed the localization of
HB-EGF from cell-cell contact sites to a homogenous distribution over
the cell surface. After 24h of transfection in COS-7 cells with AP-HBEGF-GFP, the
biotinylated, extracellular acceptor peptide in AP-HBEGF-GFP was labeled with
monovalent streptavidin-alexa fluor 568 (red, left) and imaged alongside the cytoplasmic
tail conjugated to EGFP (green, middle), and phase contrast (right). Addition of (A)
heparin (100 gg/mL) or (B) heparan sulfate (100 ptg/mL) for four hours changed the
localization of both the extracellular and intracellular domains of AP-HBEGF-GFP to a
diffuse distribution over the cell surface, rather than at cell-cell contact sites. (C)
Addition of chondroitin sulfate (100 pg/mL) for four hours had no effect on localization
of AP-HBEGF-GFP. Each row represents the same field. Scale bars are 40 pim.
..........
:m::::::r
.............
:...........
...
-
._ _ '&A
-
-
-
__
-
__=W
(A)
AP-HBEGF-GFP Fluorescence intensity at Cell-Cell Junction
After Heparin Addition
-Om
(B)
-Im
2m
-3m
-4m
-5m
-7mSm
140
120-
-aSm
--
100 -
9m
10m
80
11M
12m
14m
-
60
40
60
-14m
17m
U.-
-18
20 00
10
20
30
40
50
60
70
Pixels
Figure 4.5.2 Localization change of HB-EGF after addition of heparin
over time. (A) Confluent monolayers of COS-7 cells were transfected with APHBEGF-GFP for 24 hours. Cells were placed in PBS+, and heparin (100 pg/mL) was
added after time zero. Images were collected every 20 seconds and AP-HBEGF-GFP
localization was monitored via the GFP-tagged cytoplasmic tail. Individual frame width
is 65.5 pm. (B) The fluorescence intensity of the GFP signal is plotted versus pixel
position along the yellow line drawn on the inset fluorescence image, with the top of the
line representing pixel zero. Fluorescence intensity at the cell-cell contact site peaks at
time zero and is reduced over time, reaching the half maximal value at approximately
five minutes.
0
1
2
3
4
hours heparin
51kDa
51kDa-
51kDa-
4C
s
Monensin
Figure 4.5.3 Heparin increases accessibility of cell surface AP-HBEGFGFP. (A) Confluent monolayers of COS-7 cells expressing AP-HBEGF-GFP had
heparin (100p g/mL) added to the cell culture medium. At the indicated times, cells were
washed and the cell surface acceptor peptide was biotinylated with exogenously added
BirA on ice, then cells were immediately lysed. Lysate was probed after gel
electrophoresis with streptavidin-HRP for biotinylated proteins. After the addition of
heparin, an increase in the biotinylated cell surface form of pro-HBEGF is observed, and
this pattern is not changed with preincubation for 30 minutes and incubation during the
experiment length at 4"C or with monensin. (B) In confluent monolayers of COS-7 cells
expressing AP-HBEGF-GFP and BirA-ER, cell surface biotinylated AP-HBEGF-GFP
was labeled with mSA-AF568 (red, left) and imaged alongside the C-terminal tail with
the EGFP label (green, middle). The merged image (right) combines the red extracellular
domain with the green intracellular domain, where yellow represents co-localization of
the two signals. Flat areas of the cell with concentrated GFP signal are not labeled with
the extracellular mSA-AF568 red signal. Scale bar represents 10 im.
Extracellular HB-EGF
AP Taa Biotinviation (mSA-AF568)
C-Terminal HB-EGF Tail
EGFP
Phase Contrast
(A)
Sodium
Chlorate
(B)
Heparanase IlIl
Figure 4.5.4 HSPGs target pro-HB-EGF to cell-cell contact sites. (A)
After 24h of transfection of COS-7 cells with AP-HBEGF-GFP, cells were treated with
50 mM sodium chlorate in media without penicillin-streptomycin. After 24h of sodium
chlorate treatment, the biotinylated, extracellular acceptor peptide in AP-HBEGF-GFP
was labeled with monovalent streptavidin-alexa fluor 568 (red, left) and imaged
alongside the cytoplasmic tail conjugated to EGFP (green, middle), and phase contrast
(right). Sodium chlorate treatment led to a homogenous distribution of AP-HBEGF-GFP
over the cell surface, rather than at cell-cell contact sites (B) After 24 hours of
transfection with AP-HBEGF-GFP, cells were treated with 1.6 mU/mL heparinase III for
4 hours, then labeled with monovalent streptavidin-alexa fluor 568 (red, left), alongside
the cytoplasmic tail (EGFP) and phase contrast (right). Heparinase III treatment also led
to a homogenous distribution of AP-HBEGF-GFP over the cell surface. Each row
represents the same field. Scale bars are 40 pm.
Extracellular HB-EGF
AP Tag Biotinylation (mSA-AF568)
C-Terminal HB-EGF Tail
EGFP
Phase Contrast
(A)
AP-delHBD
HBEGF-GFP
(B)
AP-97A
HBEGF-GFP
(C)
AP-105A
HBEGF-GFP
(D)
AP-113A
HBEGF-GFP
Figure 4.5.5. The heparin-binding domain targets pro-HB-EGF to cellcell contact sites.
After 24 hours of plasmid transfection, the biotinylated,
extracellular acceptor peptide in the HB-EGF mutants was labeled with monovalent
streptavidin-alexa fluor 568 (red, left), and imaged along side the cytoplasmic tail of HBEGF conjugated to EGFP (green, middle), and phase contrast (right). (A) Deletion of the
portion of the heparin-binding domain of AP-HBEGF-GFP which lies outside of the
EGF-like domain (AP-delHBD-HBEGF-GFP), led to a homogenous distribution of HBEGF over the cell surface. (B) Mutation of the first five positively charged lysine
residues in the heparin-binding domain of HB-EGF to alanine (AP-97A-HBEGF-GFP)
had no effect on the localization of the protein. (C) Mutation of all nine positive lysine
and arginine residues of the heparin-binding domain, which lie outside the EGF-like
domain (AP- 105A-HBEGF-GFP) led to a more diffuse distribution of HB-EGF over the
cell surface, however some remains localized at cell-cell contact sites. (D) Mutation of
all twelve positive lysine and arginine residues in the heparin-binding domain of HBEGF to alanine (AP-113A-HBEGF-GFP), both those outside and inside the EGF-like
domain, led to a homogenous distribution of HB-EGF over the cell surface. Each row
represents the same field. Scale bars are 40 ptm.
AP-HBEGF-GFP
AP-113A-HBEGF-GFP
CHO-KI
(WT)
CHOpgsD-677
(-HS)
Figure 4.5.6 HSPGs and the heparin-binding domain of HB-EGF are
required for localization to cell-cell contact sites. Wild-type CHO-K1 cells
(A) & (B) and CHOpgsD-677 mutant cells (C) & (D), which do not synthesized heparan
sulfate, were transfected with either AP-HBEGF-GFP (A) & (C) or AP-l 13A-HBEGFGFP (B) & (D) and HB-EGF localization was determined based on the GFP-tagged
cytoplasmic tail. AP-HBEGF-GFP was localized to cell-cell contact sites only in CHOK1 cells (A), and was localized homogenously over the cell surface in CHOpgsD-677
cells (C). AP- 113A-HBEGF-GFP was not localized to cell-cell contact sites in either cell
line (B) & (D). Arrows highlight HB-EGF localization to cell-cell contact sites. Scale
bar represents 40 tm.
CHO-K1 (WT)
mCherry transfected
-a
A
CH~pgsD-677 (-HS)
mCherry transfected
B
~LL
0 LL
Positive control
Ci s
C
D
Tranis
Negative conitrol
~0
~U)
0 W
13L
Figure 4.5.7.
.....
HSPGs interact in trans with pro-HB-EGF.
.
Confocal
imaging of the localization of HB-EGF (green) at the junction of an mCherry (red)
transfected cell and an AP-HBEGF-GFP transfected cell. (A) Positive control sample
with AP-HBEGF-GFP and mCherry both in CHO-Ki (wild-type) cells had HB-EGF at
cell-cell contact sites (white arrows). (B) Cis binding with AP-HBEGF-GFP in a CHOKl (wild-type) cells and mCherry in CHOpgsD-677 cells (- HS) showed little HB-EGF at
cell-cell contact sites. (C) Trans binding with AP-HBEGF-GFP in CHOpgsD-677 (-HS)
cells and mCherry in CHO-Kl (wild-type) cells showed HB-EGF present at the cell-cell
junction (white arrows). (D) Negative control sample with AP-HBEGF-GFP and
mCherry in CHOpgsD-677 (-HS) cells showed no concentration of HB-EGF at the cellcell junction. Scale bars are 20 pm.
............
..
II-
-
-
-
,
:
- -
-
- -
- -
- -
-
-
I
(A)
(B)
(C)
Figure 4.5.8
ProHB-EGF is preferentially localized to cell-cell contact
sites when neighbored by a non-expressing cell. Monolayers of COS-7 cells
transfected with AP-HBEGF-GFP for 24 hours were Hoescht stained (blue), and HBEGF localization was assessed by imaging of the GFP-tagged cytoplasmic tail. (A) &
(B) HB-EGF is strongly localized to sites of cell-cell contact when neighbored with a
non-transfected cell. However, HB-EGF localization to cell-cell contact sites between
two AP-HBEGF-GFP cells is much weaker. This is also true for cells during division, as
(C) shows two AP-HBEGF-GFP cells in the late stages of mitosis. White arrows
highlight cell-cell contact sites that are missing strong AP-HBEGF-GFP localization
between two transfected cells. Images are 225 pLm wide.
Extracellular AP Tag Biotinylation
C-terminal Amphiregulin Tail
mRA.AF rAR
(:|I:P
Phann Contrant
(A)
AP-AR-GFP
(B)
AP-AR-GFP
+ heparin
(C)
AP-HBDmut
AR-GFP
Figure 4.5.9
The heparin-binding domain of amphiregulin controls
localization to cell-cell contact sites. After 24 hours of plasmid co-transfection
with BirA-ER, the biotinylated, extracellular acceptor peptide in amphiregulin was
labeled with monovalent streptavidin-alexa fluor 568 (red, left), and imaged along side
the EGFP-tagged cytoplasmic tail of amphiregulin (green, middle), and phase contrast
(right). (A) The extracellular and intracellular domains of AP-AR-GFP were localized to
sites of cell-cell contact. (B) Addition of heparin (100ptg/mL) for four hours changed the
localization of both the extracellular and intracellular domains of AP-AR-GFP to a
diffuse distribution over the cell surface, rather than at cell-cell contact sites. (C)
Mutation of all basic lysine and arginine residues in the heparin-binding domain of
amphiregulin to neutral alanine led to a diffuse distribution of amphiregulin over the cell
surface, not concentrated to sites of cell-cell contact. Each row represents the same field.
Scale bars are 40 pm.
............
Extracellular AP Tag Biotinylation
mSA-AF568
*vt'raiI'*r
r-PP
Phan& Contrat
(A)
AP-CFP-TM
(B)
AP-HBD
CFP-TM
Figure 4.5.10 Addition of a heparin-binding domain to an engineered
transmembrane protein was insufficient for localization to cell-cell
contact sites. After 24 hours of plasmid transfection, the biotinylated, extracellular
acceptor peptide was labeled with monovalent streptavidin-alexa fluor 568 (red, left), and
imaged alongside the extracellular CFP-tagged control construct (cyan, middle), and
phase contrast (right). (A) The control construct AP-CFP-TM had a diffuse localization
of the protein over the cell surface, and (B) the addition of a heparin-binding domain to
the control construct (AP-HBD-CFP-TM) did not alter this localization. Each row
represents the same field. Scale bars are 40 im.
79
Chapter Five: Role of the heparin-binding domain
5.1 Introduction
Heparan sulfate proteoglycans (HSPGs), present on the cell surface and in the
extracellular matrix, are capable of binding many growth factors.
A traditionally
proposed purpose of this interaction is to restrain a soluble ligand to the cell surface and
increase the local concentration to activate a receptor (Schlessinger et al., 1995).
Additionally, heparan sulfate binding can modulate the activity of signaling molecules or
protect them from proteolytic degradation (Conrad, 1998). Binding to heparin or HSPGs
is required for both amphiregulin and HB-EGF to activate EGFR in an
autocrine/paracrine manner (Higashiyama et al., 1993; Johnson and Wong, 1994;
Piepkom et al., 1994; Takazaki et al., 2004).
Takazaki and colleagues have suggested
that the three basic residues of the heparin-binding domain of HB-EGF that lie within a
cysteine di-sulfide loop of the EGF-like domain repels the rest of the basic residues on
the heparin-binding domain, which alters the conformation of the EGF-like region,
making it unable to activate the receptor (Takazaki et al., 2004). However, when the
heparin-binding domain is bound to heparan sulfate, the basic amino acids are neutralized,
changing the conformation of the EGF-like domain and increasing its affinity for the
EGFR.
5.2 Materials and Methods
Western blotting
Western blotting was with the following antibodies: anti-phospho-p42/44 MAPK (Cell
Signaling Technology 9101S), anti-pY 148 Phospho-EGFR Antibody (Cell Signaling
Technology 4404S), anti-EGFR (C74B9) (Cell Signaling Technology 2646S), anti-GFP
(Abcam Ab 6556), anti-Actin (Sigma A5060), anti-GAPDH (Sigma G8795) and
appropriate secondaries conjugated to horse radish peroxidase. Detection of biotinylated
proteins was accomplished with streptavidin-horse radish peroxidase (Molecular Probes
S911) with 3-5% BSA as the blocking and incubation buffer.
Confocal imaging
Confocal images were acquired with an Olympus FB1000 confocal microscope with a
slice height of 0.3 pm using a 60x oil objective.
Constructing AlkPhos-AP-HBEGF-GFP and AlkPhos-AP-113A-AlkPhos-HBEGFGFP
A plasmid for human HB-EGF conjugated to human placental alkaline phosphatase
(Raab et al., 1996) was obtained from Roselyn Adam at Children's Hospital that was
originally produced in Michael Klagsbrun's lab at Children's Hospital, Boston.
fusion protein is in the pRc/CMV vector (Invitrogen).
The
Human placental alkaline
phosphatase was amplified from this vector with the primers Fwd Alk PhosF2 (5'-CCT
GGC CAC CCC AAG CAA AGA AAG GAA T ATC ATC CCA GTT GAG GAG GAG
AAC CCG GAC TTC TGG AAC CGC-3') and reverse primer Rev Alk Phos R2 (5'TTT CTG GGC TTC GAA GAT GTC GTT CAG GCC GTC GGT GGT GCC GGC
GGG G -3'). The primers are designed to PCR human placental alkaline phosphatase out
of the plasmid with overhangs that are complementary to AP-HBEGF-GFP before and
after the N91 insertion site. Adding 10% DMSO and 3 mM total MgCl 2 to the reaction
mixture increased the yield of the appropriate PCR product. The reaction was run on an
agarose gel, and the band corresponding to alkaline phosphatase was cut out and purified
with the Qiaquick gel purification kit (Qiagen). This PCR product then served as the
primer for a quickchange mutagenesis reaction modified for large inserts on the APHBEGF-GFP and AP-1 13A-HBEGF-GFP plasmids. This purified primer was in the
PCR reaction at a concentration of 130 ng/pL with 20 ng of the base plasmid, 3 mM total
MgCl 2 , Pfu Ultra DNA polymerase, Pfu Ultra buffer, and dNTPs at standard
concentrations recommended by the manufacturer. Eighteen cycles of PCR were
performed at 95'C for 50 seconds, 60"C for 50 seconds, then 72'C for 14 minutes and
then digested with DpnI for 2 hours and transformed into GC10 E. Coli (GeneChoice).
Alkaline phosphatase release experiment
COS-7 cells were plated on 96-well dishes (6,500 cells per well) for one day, then
transfected with AlkPhos-AP-HBEGF-GFP or AlkPhos-AP-113A-HBEGF-GFP. The
following day, samples were pretreated with batimastat (10 pM) in PBS+ for 1 hour, then
stimulated with a 100 ptL solution of 100 pg/mL heparin in PBS+ supplemented with 1%
BSA. Supernatants were collected at various time points, then washed for one minute
with 100 ptL of 1.5 M NaCl in PBS+ supplemented with 1% BSA to remove any soluble
HB-EGF bound to HSPGs. The salt wash was combined with the supernatant and 40 ptL
was removed to a new plate and combined with 100 jiL of p-Nitrophenyl Phosphate
(Millipore ES009), incubated for 2 hours at 37'C, then the optical density was read at 410
nm.
Wound healing
COS-7 cells were plated in 96-well dishes at 6,500 cells/well. The following day, the
cells were serum starved and transfected with either AP-HBEGF-GFP, AP-1 13AHBEGF-GFP or GFP. Inhibitors were placed in serum free media and pre-incubated with
the monolayer for one hour. The inhibitor solution was removed, wounds were
immediately produced with a P200 pipette tip in PBS+, the PBS+ was removed and the
inhibitor solution in serum free media was replaced. Images were taken immediately at
4x. Wounds were allowed to close for 24 hours at 374C, 5% CO 2 , and wound area
images were taken again at this time. Wound area was measured in Image Pro (Version
4.5.0.29, Media Cybernetics) by highlighting the open wound area. Wound area closed
was calculated by subtracting the wound area at the beginning of the experiment from the
wound area measured at 24 hours.
Monolayer migration speed measurements
Epithelial and mesenchymal human mammary epithelial cells were each incubated with 8
ptM green-fluorescent CMFDA (Invitrogen) for 20 minutes and washed twice with PBS
before trypsinizing. Labeled and unlabeled cells were mixed at a 1:20 ratio and seeded at
60,000 cells/cm 2 for epithelial cells and 30,000 cells/cm 2 for mesenchymal cells in 24well tissue culture plates. This procedure creates a homogeneous monolayer with only a
fraction of the cells labeled to facilitate tracking of individual cells within the monolayer.
Cells were incubated overnight (16-18 hours) and serum-starved for 24 hours before
growth factor stimulation.
Samples were imaged with a Nikon TE2000 microscope
(Nikon Instruments; Melville, NY) equipped with a Solent environmental chamber
(Solent Scientific; Segensworth, United Kingdom) at 37'C and 5% CO 2 . After 1 hour of
stimulation with added factors, cells were imaged with a 4x DIC objective via brightfield
and 488 nm mercury excitation over 18 hours with 30 minute time intervals.
(Experimentsperformed by Hyung-Do Kim, Lauffenburger lab)
(Method adaptedfrom Hyung-Do Kim's Ph.D. thesis, Quantitative Analysis of 2D and
3D Models for Epidermal Growth Factor Receptor-Dependent Cell Migration in the
Context of the ExtracellularMicroenvironment, December 2008, MIT, Chapter3.2)
5.3 Results
5.3.1 The heparin-binding domain controls cell surface localization
In Chapter 4, it was demonstrated that proHB-EGF interacts with HSPGs on
neighboring cells. We hypothesized that this interaction may stabilize proHB-EGF on
the cell surface and protect it from constitutive turnover, therefore increasing the cell
surface fraction of the protein.
To test this hypothesis, the amount of proHB-EGF
expressed on the cell surface was evaluated in the different heparin-binding domain
mutants. Cell surface HB-EGF was tagged by biotinylation of the extracellular fraction
with exogenously added biotin ligase after heparin pretreatment for five minutes to make
all cell surface HB-EGF equally accessible to biotin ligase. The cell surface fraction of
HB-EGF was detected via western blotting with streptavidin-HRP and total HB-EGF was
detected with a GFP antibody to the C-terminus. Mutation of basic arginine and lysine
residues to neutral alanine in the heparin binding domain led to less cell surface proHBEGF (Figure 5.5.1 A). The amount of HB-EGF on the cell surface appears to scale with
the number of basic residues mutated. The mutants with fewer lysines and arginines
mutated had less on the surface, which suggests that heparin-binding affects cell surface
levels of HB-EGF.
In Chapter 4, it was also demonstrated that the addition of the heparin-binding
domain of HB-EGF to the control construct AP-CFP-TM was not sufficient to localize it
to sites of cell-cell contact. However, the addition of the heparin-binding domain did
increase the cell surface fraction of the protein dramatically (Figure 5.5.1 B). COS-7
cells transfected with either AP-CFP-TM or AP-HBD-CFP-TM for 24 hours were
biotinylated with biotin ligase, then lysed. The cell surface fraction of the protein is
shown with streptavidin-HRP blotting and the total protein with an anti-GFP antibody,
which recognizes CFP. The cell surface fraction of the protein is represented by one
band, and the addition of the heparin-binding domain and linker led to an appropriate
increased in size. The GFP blot for total AP-CFP-TM shows multiple size isoforms of
both AP-CFP-TM and AP-HBD-CFP-TM, which may be degradation products. This
experiment suggests that the interaction of the heparin-binding domain with HSPGs
increases the cell surface fraction of the protein.
To further investigate this hypothesis, the cell surface fraction of AP-HBEGFGFP and the heparin-binding domain mutant (AP- 113A-HBEGF-GFP) was evaluated in
CHO-KI versus CHOpgdD-677 cells, which lack heparan sulfate.
Surprisingly, the
amount of cell surface HB-EGF was equal in CHO-Ki and CHO-pgsD677 cells, and a
reduced amount of AP-1 13A-HBEGF-GFP mutant was observed on the cell surface of
both cells lines (Figure 5.5.1 C). This finding is not supportive of the hypothesis that the
interaction of the heparin-binding domain with HSPGs increases the cell surface fraction
of transmembrane proteins.
This experiment suggests that the larger amount of AP-
HBEGF-GFP on the cell surface compared to AP-l13A-HBEGF-GFP is independent of
heparan sulfate, as both CHO-KI and CHOpgsD-677 cells without heparan sulfate have
identical cell surface expression.
In both COS-7, CHO-Ki and CHOpgsD-677 cells, the distribution of HB-EGF
between the different size isoforms is altered when comparing the AP-HBEGF-GFP and
the AP- 113A-HBEGF-GFP mutant, as demonstrated with the GFP blot for total HB-EGF
(Figure 5.5.1 A,C).
Wild-type AP-HBEGF-GFP is primarily expressed at the 39 kDa
form in all three cells types. However, the heparin-binding domain mutant AP-1 13AHBEGF-GFP is expressed in the larger 51 kDa form. In the CHO cells, the larger 51 kDa
is not presented on the cell surface. Therefore, this larger form which accumulates in the
AP-1 13A-HBEGF-GFP mutant may represent altered N-terminal processing.
As less of the AP-l 13A-HBEGF-GFP mutant is presented on the cell surface
compared to wild-type AP-HBEGF-GFP, the localization of the remaining AP- 13AHBEGF-GFP was investigated. In order to address this, confocal imaging was employed
to determine the intracellular localization of the protein by imaging one optical slice
through a cell. COS-7 cells were cultured in confluent monolayers and transfected with
either AP-HBEGF-GFP or AP-1 13A-HBEGF-GFP with BirA-ER for one day with biotin
supplemented in the media.
Cells were washed, and the extracellular biotinylated
acceptor peptide was labeled with mSA-AF568. The cells were then fixed and samples
were stained with Hoescht to determine localization of the nucleus. Confocal imaging
(Figure 5.5.2) shows that wild-type AP-HBEGF-GFP had little intracellular signal, with
the majority of the extracellular and intracellular C-terminal tail co-localized in the
plasma membrane (Figure 5.5.2 A). However, the AP-1 13A-HBEGF-GFP mutant had a
dramatically different intracellular distribution of the GFP C-terminal tail (Figure 5.5.2
B). The majority of the C-terminal tail was in intracellular structures that resemble the
rough endoplasmic reticulum. The GFP C-terminal tail was additionally localized to the
nuclear envelope. Only a fraction of the total AP-1 13A-HBEGF-GFP was in the plasma
membrane, labeled by mSA-AF568 (red). However, less AF568 is also expected in the
plasma membrane as it is known to be distributed over the entire cell surface, while the
AP-HBEGF-GFP construct is concentrated at cell-cell contact sites.
The heparin-binding domain controls the amount of HB-EGF and AP-CFP-TM
on the cell surface, however it is not due to an interaction with HSPGs. Therefore, it is
possible that the heparin-binding domain interacts with a second partner, or that the
intracellular accumulation and/or differential N-terminal processing of AP-l 13AHBEGF-GFP alters the steady state cell surface expression. The second extracellular
loop of CD9 has been shown to interact with a peptide corresponding to the heparinbinding domain of HB-EGF (Sakuma et al., 1997), which may control cell surface
expression.
5.3.2 ProHB-EGF does not cluster EGFR
As a trans interaction between HSPGs and proHB-EGF clusters HB-EGF to cellcell contact sites, the question arose of whether a trans reaction between proHB-EGF and
EGFR could cluster EGFR at cell-cell contact sites. Two populations of COS-7 cells
were co-cultured together after transfection with AP-HBEGF-GFP alone or a construct
for acceptor peptide tagged EGFR (AP-EGFR) and BirA-ER. Without co-transfection of
BirA-ER with AP-HBEGF-GFP, only AP-EGFR is biotinylated and visualized with
mSA-AF568, and AP-HBEGF-GFP is visualized with the C-terminal GFP tag only. APEGFR was localized homogenously over the cell surface, with no preferential localization
to sites of cell-cell contact. AP-EGFR cells in contact with AP-HBEGF-GFP expressing
cells showed no change in localization of either construct or any significant enrichment
of EGFRs at sites of cell-cell contact (Figure 5.5.3 A). This suggests that proHB-EGF
does not cluster EGFRs at cell-cell contact sites.
As HSPGs cluster HB-EGF at cell-cell contact sites, the role of this cluster may
be to activate the EGFR in a trans manner. HB-EGF is known to signal with EGFR and
ErbB4 in a juxtacrine manner. As the addition of heparin or mutation of the heparinbinding domain diminishes the clustering of HB-EGF at cell-cell contact sites, this
localization change may decrease juxtacrine activation of the EGFR as less is available at
cell-cell contact sites for activation of the receptor. Therefore, we hypothesized that the
localization change induced by mutation of the heparin-binding domain may decrease
juxtacrine signaling and lead to a drop in EGFR phosphorylation. However, no change in
EGFR phosphorylation was observed after transfection of AP-HBEGF-GFP or AP-l 13A-
HBEGF-GFP compared to GFP transfected COS-7 cells (Figure 5.5.3 B). To reduce
noise from autocrine signaling, samples were treated with batimastat (10 [M) for five
hours before lysis. Transfection with AP-HBEGF-GFP or AP-113A-HBEGF-GFP did
not affect ERK or EGFR activation, nor led to downregulation of the EGFR. Reducing
autocrine signaling with batimastat reduced phospho-EGFR and phospho-ERK, however
there was still no change across different transfection conditions. As an endogenous
EGFR mediated autocrine signaling loop is present in COS-7 cells (Kain and Klemke,
2001), we hypothesize that transfection with HB-EGF does not additionally contribute to
EGFR signaling.
In order to test the role of heparin on HB-EGF signaling, confluent monolayers of
COS-7 cells expressing AP-HBEGF-GFP, AP-113A-HBEGF-GFP or GFP and serum
starved for 24 hours were treated with heparin (100 tg/mL) for various amounts of time,
then lysed. The addition of heparin led to a dramatic decrease in phospho-ERK levels
after only five minutes (Figure 5.5.4). It was originally hypothesized that this drop in
ERK signaling was due to an immediate decrease in juxtacrine signaling after the
addition of heparin as proHB-EGF is moved away from sites of cell-cell contact.
However, pre-treatment of the cells with AG1478, GM6001, or batimastat (data not
shown) led to a dramatic reduction in phospho-ERK levels at all time points, suggesting
that baseline ERK activation and the heparin-induced drop was due to EGFR autocrine
signaling rather than juxtacrine activation. Additionally, the drop in ERK activation was
observed in GFP transfected cells (data not shown), and AP-1 13A-HBEGF-GFP
transfected cells; however, the heparin-induced decrease for AP- 113A-HBEGFP-GFP
was time delayed compared to AP-HBEGF-GFP (Figure 5.5.4). This suggests that the
drop in phosho-ERK levels was due to an endogenous ligand or signal, rather than the
transfected HB-EGF construct. A similar heparin-induced drop in phospho-ERK was
observed in transfected and non-transfected HeLa cells (data not shown). Total ERK
levels remained constant after heparin addition in COS-7 and HeLa cells (data not
shown).
5.3.3 The heparin-binding domain controls ectodomain shedding
As heparin dramatically changed the localization of proHB-EGF from sites of
cell-cell contact to a homogeneous distribution over the cell surface, we hypothesized
that this localization change may increase access to proteases and affect ligand cleavage.
To assess release of HB-EGF into the media, human placental alkaline phosphatase was
inserted into the extracellular domain of AP-HBEGF-GFP and AP-1 13A-HBEGF-GFP
near the N-terminus. This allows for sensitive detection of HB-EGF release by assaying
for alkaline phosphatase activity in the media of transfected cells. As HB-EGF release is
typically low in non-stimulated conditions, and the basic residues in the heparin-binding
domain likely make the protein non-specifically sticky, ELISA measurements of HBEGF concentration in the media is challenging due to the need for sample concentration
and significant sample loss.
The addition of heparin (100 [ig/mL) to confluent
monolayers of COS-7 cells transfected with wild-type HB-EGF (AlkPhos-AP-HBEGFGFP) increased alkaline phosphatase activity in the media (Figure 5.5.5 A), suggesting
that the heparin-induced localization change of proHB-EGF away from cell-cell contact
sites upregulates ligand cleavage.
Treatment with the protease inhibitor batimastat
(lOpM) inhibited heparin-induced cleavage of both wild-type and mutant HB-EGF.
Interestingly, the heparin-binding domain mutant alkaline phosphatase fusion (AlkPhosAP- 113A-HBEGF-GFP) had higher levels of cleavage compared to wild-type HB-EGF
and was unaffected further by the addition of heparin. These data suggest that the trans
interaction of pro-HB-EGF with HSPGs at cell-cell contact sites prevents proteolytic
release of the ligand.
However, the interaction of HB-EGF with CD9, which also
involves the heparin-binding domain (Sakuma et al., 1997), may serve to inhibit
proteolytic ligand release, as we found a similar increase in alkaline phosphatase activity
in the medium for AlkPhos-AP- 113A-HBEGF-GFP when using the HSPG-lacking
CHOpgsD-677 cells (Figure 5.5.5 B). Since the CD9 interaction appears to operate in cis
(Sakuma et al., 1997), and therefore ought not to depend on HB-EGF localization to cellcell contact regions, the two heparin-binding domain interactions may work in series to
provide a multi-layer control on ligand release.
5.3.4 Wound healing and migration
As HB-EGF is released upon the addition of heparin, we sought to explore
activation of EGFR pathways and its role on migration. As heparin increases proHBEGF cleavage, we hypothesized that HB-EGF transfection and heparin stimulation may
increase migration and wound healing by inducing autocrine HB-EGF signaling. Wound
healing assays in COS-7 cells transiently expressing AP-HBEGF-GFP and AP-1 13AHBEGF-GFP showed no change in wound healing compared to the GFP transfected
control (Figure 5.5.6 A). However, treatment of the samples with batimastat reduced
wound closure, suggesting that cleavage of an endogenous cell surface ligand is driving
wound closure. In order to investigate the driving forces of endogenous wound closure in
COS-7 cells in more detail, an investigation of the EGFR pathway in COS-7 cells shows
that preincubation with the tyrosine kinase inhibitor AG1478 for one hour, and continued
inhibition during the wound healing experiment significantly reduced wound closure
(Figure 5.5.6 B).
Additionally, inhibition of MEKI with the inhibitor PD98059 or
protease cleavage with the inhibitor GM6001 decreased wound closure rates, but not to
the same degree as AG1478. It should be noted, however, that the addition of rhHBEGF
(100 ng/mL) did not increase the rate of wound closure, nor did an HB-EGF blocking
antibody significantly decrease wound closure rates. These data suggest that wound
healing is mediated through the EGFR system in COS-7 cells, however, endogenous
signaling in this system is already saturated. Therefore the addition of rhHB-EGF, or
transfection with HB-EGF has no additive response. This hypothesis is supported by the
fact that AP-HBEGF-GFP transfection shows no increase in phospho-EGFR levels, the
heparin induced drop in phospho-ERK is not affected by transfection either, and a
background signaling loop has been previously observed by others in COS-7 cells (Kain
and Klemke, 2001).
Heparin may increase rates of proHB-EGF cleavage, however,
heparin has an inhibitory effect on wound closure in a dose dependent manner, with
inhibition observed at heparin concentrations of 10-100 ptg/mL (Figure 5.5.6 C).
To investigate the role of heparin on migration in further detail, studies were
completed in human mammary epithelial cells (HMECs). Two different HMEC cell lines
were used. The first (epithelial) were derived from a reduction mammoplasty tissue
sample (Elenbaas et al., 2001). The second cell line (mesenchymal) was derived from the
first, as transformation with three genes (SV40 large-T antigen, the telomerase catalytic
subunit, and an H-Ras oncoprotein) caused the cells to form tumors when injected into
mice (Elenbaas et al., 2001). The tumors formed were poorly differentiated carcinomas
that lacked the estrogen receptor, and therefore may serve as a model for estrogen
receptor-negative breast cancers (Elenbaas et al., 2001). Individual COS-7 cells are not
migratory when cultured in confluent monolayers with cell-cell contact, however wound
healing does stimulate COS-7 migration. However, wild-types HMECs are very dynamic
and migrate even in a confluent monolayer. This model system allows for analyzing
single cell migration behavior, with the role of juxtacrine signaling, as the cells can
remain in contact. Individual cell migration of HMECs in a monolayer is measured by
labeling a subpopulation of the cells (10%) with a cell tracker fluorophore, then the
coordinates of the labeled cells are tracked over time. The migration speed of epithelial,
but not mesenchymal HMECs was increased significantly by the addition of 100 ng/mL
of EGF or HB-EGF, indicating EGFR signaling drives migration in the non-transformed
epithelial cell line (Figure 5.5.7). HMECs have been shown to produce amphiregulin at
high levels, as well as transforming growth factor-a, epiregulin, and HB-EGF (Dong et
al., 1999; Rodland et al., 2008). As amphiregulin and HB-EGF are both heparin-binding
proteins, whose localization is changed upon the addition of heparin, we sought to test the
effect of heparin (100 ptg/mL), heparan sulfate (100 pg/mL) and heparinase III (1.6
mU/mL) on migration. Similar to COS-7 cell wound healing, heparin inhibited the
migration speed of epithelial-like HMECs, but not mesenchymal-like (Figure 5.5.7).
Heparinase III treatment was also inhibitory, as heparan sulfate chains produced by
heparinase likely act in the same manner as endogenously added heparan sulfate,
however the change was not as strong. Together, the COS-7 cell wound healing results
taken with the HMEC migration speed results indicate the heparin inhibits cell migration.
In summary, heparin may increase rates of cleavage, however, it does not lead to
phenotypic changes associated with increased autocrine signaling, such as enhanced
migration or EGFR activation. We hypothesize that heparin may disrupt extracellular
localized autocrine signaling loops of heparin-binding growth factors. This disruption
may lead to an overall decrease in receptor activation in various pathways which affect
migration or disrupt the chemotactic gradient produced by ligand cleavage, binding and
diffusion.
5.4 Discussion
Here we show that mutation of the heparin-binding domain or the addition of
heparin upregulates proteolysis of HB-EGF (Figure 5.5.5 A).
It is conceivable that
heparin and heparan sulfate also dissociate proHB-EGF from HSPGs and thus cell-cell
contacts sites and cause ligand cleavage in vivo. Heparin and low-molecular weight
heparin are administered via intravenous infusion for their anti-coagulant properties in
the clinic for prevention and treatment of thromboembolic disorders.
Intravenous
injection of heparin likely does not affect HB-EGF, as it stays within the bloodstream,
and HB-EGF does not appear to be expressed by endothelial cells (Nakata et al., 1996).
However, heparin is cleared by the kidney's renal tubular cells (Young et al., 2004),
which may interact with basolateral HB-EGF expressed in collecting tubules and change
the basolateral localization of the growth factor to a homogenous distribution over the
cell surface.
Additionally, free heparin is secreted by mast cells upon degranulation.
Mast cells and HB-EGF signaling are both implicated in the biological processes of
wound healing (reviewed in (Noli and Miolo, 2001) (Tokumaru et al., 2000),
angiogenesis (reviewed in (Galinsky and Nechushtan, 2008) (Ongusaha et al., 2004), and
the pathogenesis of atherosclerosis (reviewed in (Kalesnikoff and Galli, 2008) (Nakata et
al., 1996). Additionally, dermal mast cells themselves express HB-EGF mRNA (Artuc et
al., 2002). Aside from the pro-angiogenic effects of mast cells in tumor angiogenesis,
separate studies show mast cell heparin to inhibit tumor growth (reviewed in (Galinsky
and Nechushtan, 2008)). Extracellular free heparan sulfate is generated by degradation of
cell surface HSPGs with heparanase, whose expression is upregulated in all analyzed
human cancers (Vlodavsky et al., 2007). It is possible that heparanase leads to a local
concentration of free heparan sulfate high enough in the interstitial space to dissociate
proHB-EGF from cell-cell contact sites and stimulate cleavage of proHB-EGF.
Heparin and mutation of the heparin-binding domain of HB-EGF led to increased
rates of proHB-EGF proteolysis and accumulation in the media. We hypothesize that
clustering of HB-EGF at cell-cell contact sites via a trans interaction with HSPGs may
prevent proteolysis of the ligand by restricting access to the transmembrane ADAMs.
Therefore, upon the addition of heparin or mutation of the heparin-binding domain, HBEGF is dissociated from cell-cell contact sites and access to ADAMs is increased,
resulting in increased rates of cleavage. Interestingly, it has recently been reported that
heparan sulfate interacts with ADAM12, which controls ADAM12 sheddase activity by
serving as a molecular switch (Sorensen et al., 2008).
Additionally, removal of cell
surface heparan sulfate has been reported to increase TACE activity and ErbB4 cleavage
(Maatta et al., 2009). Therefore, in our experiments, stimulation with heparin may also
increase ADAM activity; however, this is not expected from mutation of the heparinbinding domain.
Therefore, despite the role heparan sulfate may play on ADAM
activation, the heparin-binding domain of HB-EGF also controls proteolysis.
To our surprise, despite the upregulation in release of HB-EGF upon the addition
of heparin, heparin had an inhibitory effect on cell migration in COS-7 wound healing
(Figure 5.5.6 B) and individual cell migration of confluent monolayers of epithelial
HMECs (Figure 5.5.7).
Heparin also dramatically reduced ERK phosphorylation
immediately after its addition (Figure 5.5.4), which may play a role in inhibiting
migration, however ERK levels return to baseline after a few hours and cell migration
continued to be inhibited for experimental lengths of 12-24 hours.
Interestingly, the
mesenchymal transformed HMEC cell line, which is more tumorigenic than the epithelial
HMECs, migrated at lower speeds and did not respond to heparin addition, or even
stimulation with EGF or HB-EGF (Figure 5.5.7).
Therefore, the addition of heparin,
heparan sulfate, or degradation of HSPGs with heparinase made the epithelial HMECs
act more like the tumorigenic mesenchymal HMECs. We hypothesize that HB-EGF may
play a crucial role in maintaining cell polarity, as it is primarily localized at cell-cell
contact sites between non-homogenous cell-cell contact sites (Figure 4.5.8) and proHBEGF juxtacrine signaling can be growth inhibitory (Iwamoto et al., 1999). The loss of
cell polarity of luminal HMECs has been suggested to play a key role in the
morphogenesis to a cancerous phenotype (Zhan et al., 2008). Therefore, we hypothesize
that a localization change of proHB-EGF and/or pro-amphiregulin due to degradation of
HSPGs by heparanase may contribute to a loss of polarization in luminal HMECs and
contribute to cancer morphogenesis.
5.5 Figures
4:
4', /
*
Ilk
PC
4',
(A)
$
Cell Surface HB-EGF
-
51kDa
-
39kDa
Streptavidin-HRP
Total HB-EGF
Anti-GFP
-
'fi
Cell Surface Fraction
39kDa
I CHO-KI
St eptavidin-HRP
39kDa-
Cell Surface HB-EGF
Streptavidin-HRP
-
CHO-677 I
-
SikD*
-
51kDa
otal Construct
Anti-GFP
Total HB-EGF
Anti-GFP
Anti-Actin
Figure 5.5.1 Mutation on the heparin-binding domain decreases the cell
surface fraction of HB-EGF. (A) Cell surface expression of AP-HBEGF-GFP and
three of the heparin-binding domain mutants in order of increasing number of lysine and
arginine mutations show decreasing cell surface expression with increasing mutation of
lysine and arginine. Confluent monolayers of COS-7 cells transfected for 24 hours were
washed, heparin treated (100 ptg/mL, 5 minutes), then cell surface biotinylated with biotin
ligase on ice. Cells were washed, then lysed and run on SDS PAGE. Blotting with
streptavidin-HRP shows the cell surface expression of each protein, and the total protein
is shown with blotting to the GFP C-terminal tail. (B) Addition of the heparin-binding
domain (AP-HBD-CFP-TM) to the control construct AP-CFP-TM increased the fraction
of the protein on the cell surface. GFP shows the total amount of the construct, as it also
recognizes CFP. Actin served as a loading control. (C) Confluent monolayers of CHOK1 and CHOpgsD-677 (CHO-677) cells transfected for 24 hours with AP-HBEGF-GFP
or the heparin-binding domain mutant AP-113A-HBEGF-GFP show decreased
expression of the mutant on the cell surface of both the CHO-Ki and CHOpgsD-677 cells.
Total HB-EGF is probed with an antibody to GFP.
..........
.- ........................
.......................
....
..
....................
..
....
....
....
.............
........
..........
Extracellular HB-EGF
AP Tag Blotinylation (mSA-AF568)
C-Terminal HB-EGF Tail
EGFP
Merge
(A)
AP
HBEGF-GFP
(B)
AP-113A
HBEGF-GFP
Figure 5.5.2 The heparin-binding domain mutant is primarily localized
in the intracellular space. Confluent monolayers of COS-7 cells were transfected
with AP-HBEGF-GFP (A), or the heparin-binding domain mutant AP-1 13A-HBEGFGFP (B) along with BirA-ER and biotin supplemented in the media. After 24 hours, cells
were washed and the extracellular biotinylated acceptor peptide was labeled with mSAAF468 (red, left). Cells were washed, then fixed with 4% paraformaldehyde and nuclei
were stained with Hoescht (blue). The AP-1 13A-HBEGF-GFP has increased GFP Cterminal tail (green, middle) localization in the intracellular space compared to APHBEGF-GFP. The merged imaged shows co-localization of the extracellular biotinylated
AP tag and the GFP C-terminal tail (yellow, merged, right) in the plasma membrane.
Scale bars are 20 pm.
C-Terminal HB-EGF Tail
EGFP
Extracellular EGFR
AP Tag Biotinylation (mSA-AF568)
Overlay
Batimastat
Pre-treatd
HB 113A GFP HB 113A GFP
Total EGFR
~m
7
pY1148 Phospho-EGFR
-
Phospho-ERK
Figure 5.5.3 EGFR localization and activation. (A) COS-7 cells transfected
with AP-EGFR/BirA-ER and AP-HBEGF-GFP were co-cultured in confluent
monolayers. After 24 hours, the biotinylated AP-EGFR was visualized with mSA-AF568
and the non-biotinylated AP-HBEGF-GFP was visualized via the GFP-tagged
cytoplasmic tail. Junctions between a HB-EGF expressing cell and an EGFR expressing
cell show no concentration of EGFR at cell-cell contact sites, as is seen around the rest of
the cell periphery. Scale bars are 20 ptm. (B) COS-7 cells transfected with AP-HBEGFGFP, AP-l 13A-HBEGF-GFP, or GFP and serum starved for 24 hours were western
blotted for total EGFR, phospho-tyrosine 1148 EGFR, and phospho-ERK. Batimastat
(10 pM) treatment was for five hours before lysis.
0 15 30 45 60 90 120 180 240
Time (minutes)
AP-HBEGF-GFP
Phospho-ERK
AP-113A-HBEGF-GFP
Phospho-ERK
AP-HBEGF-GFP
GAPDH
AP-113A-HBEGF-GFP
GAPDH
Figure 5.5.4 Heparin reduces ERK activation. Confluent monolayers of COS7 cells were transfected with AP-HBEGF-GFP or AP- 13A-HBEGF-GFP and serum
starved for 24 hours. Cells were stimulated with heparin (100 pg/mL) for various times
and lysed. Heparin led to a drop in phospho-ERK activation at 30 minutes, which rose
back to baseline levels by 3-4 hours in AP-HBEGF-GFP transfected cells. However,
cells transfected with AP- 113A-HBEGF-GFP showed a similar, but time delayed
decrease in phosho-ERK. GAPDH is a loading control.
Alkaline phosphatase-AP-HBEGF-GFP release
0.5-
0.4.
E
c 0.3-
--
WT HB-EGF
-a-
HBD mutant HB-EGF
-
e--- WT
-
0
0.2-
HB-EGF (+heparin)
HBD mutant HB-EGF (+heparin)
a WT HB-EGF (+BATI +heparin)
o HBD mutant HB-EGF (+BATI +heparin)
0.1!
00
30
60
90
120 150 180 210 240
Time (minutes)
Alkaline Phosphatase-AP-HBEGF-GFP Release in CHOpgsD-677 cells
-.- WT HB-EGF
-o- HBD Mutant HB-EGF
Time (minutes)
Figure 5.5.5
Interaction with HSPGs reduces proHB-EGF cleavage.
COS-7 cells (A) or CHO-pgsDO677 cells (B) transfected with wild-type HB-EGF
(AlkPhos-AP-HBEGF-GFP) (e) or the heparin-binding domain mutant (AlkPhos-AP113A-HBEGF-GFP) (o) were either pre-treated with 10 pM batimastat (BATI) or PBS+
alone for 1 hour. Cells were stimulated with either heparin (100 tg/mL) alone (black
dashed line), or heparin and batimastat (gray dashed line), or unstimulated (black solid
line) and media was collected at various time points, then cells were incubated with a 1.5
M salt solution for 60 seconds to release any soluble HB-EGF bound to HSPGs.
AlkPhos-AP-HBEGF-GFP release is increased by treatment with heparin, and cleavage is
further increased by mutating the heparin-binding domain (AlkPhos-AP-l 13A-HBEGFGFP). Pretreatment with batimastat before the addition of heparin blocks both AlkPhosAP-HBEGF-GFP and AlkPhos-AP- 13A-HBEGF-GFP cleavage. Data shown is average
and standard deviation of three biological replicates from one of three independent
experiments.
(A)
1000000 ~
MAP-HBEGF-GFP
900000 -
60000
MAP-113A-HBEGF-GFP
GFP
t
B
800000 -
0i41
Wound Closure After Twenty-Four Hours
Wound Closure After Twenty-Four Hours
E2 700000
40000
3
600000
MGFP
20000 0 -
500000
400000
S20000(0 -
300000-
C
10000 0 -
200000 100000 0-
Control
Batimastat
Wound Area Closed After Twenty-Four Hours
700000600000 500000-
* AP-HBEGF-GFP
400000-
* AP-1 13A-HBEGF-GFP
300000-
* GFP
2000001000000-
,,
100
I
10
I
1
0.1
0.01
1
0.001
j
0
Heparin (pg/mL)
Figure 5.5.6 Wound healing in COS-7 cells. Confluent monolayers of COS-7
cells were transfected with the plasmids indicated for 24 hours and serum starved.
Wounds were made with pipette tip in 96 well dishes (n=8), wound area was measured,
then the wounds were allowed to close for 24 hours, and wound area was measured again.
Wound area closed was calculated by subtracting the wound area produced at the
beginning of the experiment, and subtracting the wound area measured at 24 hours. (A)
AP-HBEGF-GFP transfection does not change the wound closure rate compared to the
GFP transfected control. Batimastat (10 pM) pre-treatment for one hour prior to
induction of the wound and treatment throughout the experiment decreased wound
closure rates independent of the transfection condition. (B) GFP transfected COS-7
cells were pre-treated with AG1478 (10 pM), PD98059 (25 pM), HB-EGF blocking
antibody (10 pg/mL), GM6001 (10 pM), and recombinant human HB-EGF (rhHBEGF)
(100 ng/mL) for one hour prior to induction of the wound and continuous treatment
throughout the experiment. AG1478 drastically reduced wound closure rates; however,
stimulation of wound healing with rhHBEGF had no effect. (C) Treatment of
transfected COS-7 cells with heparin throughout the wound healing experiment reduced
wound closure at high heparin concentrations (100 pg/mL) independent of the
transfection condition. Data shown is average and standard deviation of eight biological
replicates.
Human Mammary Epithelial Cell Migration Speed
30
0 Control (Serum Free)
U EGF
N HB-EGF
25
20
Heparinase III
0 Heparin
_
E
D:Heparan Sulfate
15
a,
U)
10
0~
0
Epithelial
Mesenchymal
Figure 5.5.7 Migration speed of individual HMECs within a confluent
monolayer. Confluent monolayers of epithelial and mesenchymal human mammary
epithelial cells were seeded with 5% of the cells labeled with green fluorescent CMFDA.
The coordinates of fluorescently labeled cells were tracked with timelapse microscopy
over 18 hours with data points acquired every 30 minutes and migration speed was
calculated based on the distance each individual cell traveled per time. Stimulation with
recombinant EGF (100 ng/mL) and HB-EGF (100 ng/mL) led to a large increase in
migration speed in epithelial, but not mesenchymal HMECs. However, the addition of
heparin (100 [tg/mL), heparan sulfate (100 [tg/mL), or heparinase III digestion (1.6
mU/mL) of cell surface HSPGs led to a reduction in migration speed in epithelial, but not
mesenchymal HMECs. This experiment was executed and analyzed by Hyung-Do Kim
(LauffenburgerLab, MIT)
100
Chapter Six: Conclusions and future directions
In summary, we have developed a novel strategy for two-color labeling and
tracking of murine heparin-binding epidermal growth factor in living cells, with the
ability to distinguish the C-terminal tail from the extracellular EGF-like domain. Using
this construct, we discovered that a trans interaction between proHB-EGF and HSPGs is
responsible for localization of proHB-EGF to sites of cell-cell contact. Additionally, the
heparin-binding domain increases the cell surface fraction of transmembrane proteins and
prevents proteolytic processing of proHB-EGF. As proHB-EGF signaling in juxtacrine
mode can cause growth inhibition, and autocrine signaling leads to cell proliferation and
migration, the role of HSPGs in regulating this balance likely plays a crucial role in the
resulting cell fate. This balance may be upset in tumors, as enzymes that alter the number
of extracellular heparan sulfate chains on proteoglycans have been associated with cancer,
such as heparanase and HSulfl.
6.1 Future directions: Use of AP-tagged EGFR ligands
Acceptor peptide tagging and biotinylation with biotin ligase proved to be very
specific labeling method for ligands in the EGFR family. Additionally, this technique
allowed us to distinguish the cell surface fraction from the total pool of HB-EGF. One
powerful use for the AP-HBEGF-GFP construct that was not adequately explored in this
thesis would be for ratiometric studies of fluorescence of the extracellular domain
compared to the C-terminal tail of the ligand in individual cells, allowing one to study
101
ligand cleavage.
The cell surface levels of proHB-EGF could be detected with the
acceptor peptide tag, and compared to the fluorescence intensity of the GFP C-terminal
tail via flow cytometry to allow for individual cell measurements of proteolysis.
In
addition to HB-EGF, we have also produced identical constructs for amphiregulin and
transforming growth factor-a.
Another possible avenue of investigation with AP-tagged EGFR ligand is
measurements of internalization and recycling.
This avenue has been explored in depth
with the EGF receptor, as receptor downregulation and recycling has important
consequences on EGFR signaling, however very little information exists on this dynamic
for the ligand. The steady state internalization and recycling of the pro-form of EGFR
ligands likely plays a role in juxtacrine EGFR activation and the conversion to the soluble
form for autocrine signaling.
Endocytosis of wild-type versus the heparin-binding
domain mutant would be interesting to compare. The acceptor peptide tag inserted into
the extracellular domain of EGFR ligands could be utilized for these measurements. To
accomplish this, the cell surface pool of the protein is biotinylated, then at different times
monovalent streptavidin is added to quench the cell surface pool and the cells are
immediately lysed. As the biotin-streptavidin bond is extremely strong, under the right
lysis and gel electrophoresis conditions, the bond can remain intact. Therefore, after gel
electrophoresis, the biotinylated protein that was internalized can be detected by western
blotting and probing the gel with streptavidin-HRP.
A similar strategy could be used to measure recycling of the protein back to the
cell surface. The cell surface pool of the AP-tagged EGFR ligand would be biotinylated
with exogenously added biotin ligase and allowed to internalize for a sufficient amount of
time. Then all samples in the timecourse would have the cell surface pool of biotinylated
protein quenched with monovalent streptavidin (un-labeled).
After this, at various
timepoints, the internalized biotinylated non-quenched ligand that returned to the cell
surface would be detected with monovalent-streptavidin-AF568 (or perhaps a fusion of
monovalent streptavidin with alkaline phosphatase or horseradish peroxidase) over time.
These methods could allow for quantification and comparison of endocytosis and
recycling of EGFR ligands.
102
6.2 Future directions: Signaling with heparin-binding domain mutant HB-EGF
COS-7 cells were an excellent experimental system for evaluating the localization
of HB-EGF to cell-cell contact sites as they are easily transfected, large cells that localize
HB-EGF to large planes of cell-cell contact. However, unfortunately, COS-7 cells were
not properly suited for the study of HB-EGF signaling via the EGFR or subsequent
downstream effects, such as cell migration, proliferation, or growth inhibition. Therefore,
studies on the effect of mutation of the heparin-binding domain of proHB-EGF would be
an interesting avenue of pursuit in a proper model system that responds to HB-EGF
autocrine signaling through growth and migration, and responds to HB-EGF juxtacrine
signaling via growth inhibition or apoptosis. Using this new model system, one could
determine whether the localization change induced by heparin, heparan sulfate, or
heparinase digestion of HSPGs could lead to less juxtacrine signaling, and higher levels
of autocrine signaling.
Additionally, we hypothesize that proHB-EGF juxtacrine
signaling may serve as a contact inhibition signal between two different tissue types and
provide a barrier for cell growth of one tissue into another. As the heparin-binding
domain mutant leads to loss of localization at cell-cell contact sites, and therefore would
likely lead to loss of tissue polarization, it would be interesting to evaluate the outcome of
expression of the heparin-binding domain mutant in vivo. This could be achieved by
making a knock-in of the heparin-binding domain mutant HB-EGF in mice. Additionally,
injection of tumor cells expressing wild-type versus the heparin-binding domain mutant
and evaluation of tumor formation would be an interesting study.
6.3 Future directions: HB-EGF localization change in vivo.
As discussed in section 5.4, proHB-EGF may undergo a change in localization as
seen with the addition of heparin in vivo.
This could be stimulated by heparin
administered intravenously as an anti-coagulant, heparin release by degranulating mast
cells, loss of heparan sulfate from cell surface proteoglycans by heparanase or Hsulfl
action, or by heparan sulfate chains generated by degradation of HSPGs with these
enzymes. Therefore it would be interesting to determine if any of these mechanisms do
indeed change the localization HB-EGF in vivo, and if the localization changed led to
any differences in tissue phenotype.
103
6.4 Future directions: Structure of the HSPG-proHBEGF-CD9 complex
ProHB-EGF interacts with both HSPGs and CD9. A peptide corresponding to the
second extracellular loop of CD9 binds to a peptide consisting of the heparin-binding
domain of HB-EGF with a Kd of 37 pM (Sakuma et al., 1997). However, the Kd for
binding of heparin to the heparin-binding domain of HB-EGF is much lower (~28 nM)
(Sakuma et al., 1997). Interestingly, ideas in the literature have been presented the large
second extracellular loop of CD9 might have a structure, including disulfide bridges, and
that this region may control tetraspanin-tetraspanin homo and heterodimerization (Zoller,
2009). Therefore, despite the high Kd of a CD9 peptide binding to the heparin-binding
domain, the Kd of the fully formed, folded protein may be lower. The EGF-like domain
has also been demonstrated to interact with CD9 and play a crucial role in upregulation of
juxtacrine activity (Nakamura et al., 2000) and the membrane-anchoring domain of
proHB-EGF is also required for juxtacrine signaling (Dong et al., 2005). Therefore, the
proHB-EGF-CD9 interaction may span many domains of proHB-EGF.
As HB-EGF,
CD9, and HSPGs all cluster at cell-cell contact sites (Nakamura et al., 2001), we suggest
that multi-protein complex may be required to regulate juxtacrine activity.
Ternary
complexes with HSPGs have been discovered before, as fibroblast growth factor (FGF)
requires HSPG interaction along with the FGF receptor to form an active signaling
complex (reviewed in (Harmer, 2006)).
6.5 Future Directions: Computational analysis
To better understand the role of HSPGs in juxtacrine and autocrine/paracrine
signaling via the EGFR and ErbB4, computational modeling could provide useful insight
in this network. We hypothesize here that proHB-EGF is primarily localized to cell-cell
contact sites between an HB-EGF expressing cell and a non-producing cell because
competition for binding between two cells expressing proHB-EGF reduces the amount
accumulated at cell-cell contact sites. Computational modeling could be employed to
determine if this is possible under physiological conditions of HB-EGF and HSPG
expression. Additionally, as the interaction between proHB-EGF and HSPGs inhibits
104
cleavage of the ligand, computational modeling of the balance between juxtacrine and
autocrine/paracrine signaling of HB-EGF could employed to better understand the
balance between the two. Additionally, this model could shed insight onto the role of
HSPG altering enzymes, such as HSulfI and heparanase, on the balance between HBEGF juxtacrine vs. autocrine/paracrine signaling in cancer.
105
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