ROLE OF THE DISCOIDIN DOMAIN RECEPTOR PROTEINS IN
ARTHEROSCLEROSIS: INTERACTION WITH LIPIDS AND COLLAGEN
DISSERTATION
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy
in the Graduate School of The Ohio State University
By
MICHELLE JON NAUERTH, M.A.
Graduate Program in Biochemistry
The Ohio State University
2011
Dissertation Committee:
Charles L. Brooks, Advisor
Andre Francis Palmer
Dennis Bong
Copyright by
Michelle Jon Nauerth
2011
ABSTRACT
Discoidin domain receptors (DDR1 and DDR2) are unique tyrosine kinase
receptors (RTKs) in that they bind to and become phosphorylated by collagens,
particularly collagen type I; the most abundant protein in the extracellular matrix (ECM).
It is currently not known if collagen is the only ligand for these proteins, since some
members of the discoidin family also bind phospholipids via their discoidin domain. In
either case, the mechanisms for binding any ligand by these two proteins are not
completely understood. It is generally known that receptors from the RTK family bind a
ligand and induce down-stream phosphorylation of the intracellular tyrosine kinase, thus
controlling several cellular processes. The goal of this study is to demonstrate a
possibility for alternative ligands, other than collagen, for the discoidin domain receptor
proteins, and elucidate how a mechanism of binding may occur.
The data presented here qualitatively addresses the possibility that phospholipids
may be a ligand for the DDRs. Several proteins in the blood coagulation pathway bind
phospholipids via their discoidin-like domains. In addition, the three-dimensional crystal
structures of these proteins have been resolved. Utilizing their amino acid sequence, as
well as their three-dimensional structure as a template, a series of experiments was
designed to elucidate a possible DDRs:phospholipid interaction. There is a strong
indication, as my results demonstrate that DDRs bind to phospholipids. Threedimensional molecular models of the DDRs were proposed and used to understand how
ii
these proteins might bind to a plasma membrane, which is comprised of phospholipids.
The finished structure of the DDRs mimics those proteins in the blood coagulation
pathway which bind phospholipids and support my hypothesis that DDRs may favor
phospholipids as a ligand.
Based on the results of the molecular modeling study, the extracellular domain
(ECD), which is the proposed ligand binding region of the DDRs, was employed in a
series of experiments involving platelet aggregation. It is known that the preferred ligand
for the DDRs is fibrillar collagen. The data presented in this study is novel, and shows
that DDRs inhibit platelets from aggregating via an indirect interaction with collagen.
These results may elucidate a possible role for the DDRs in arteriole plaque formation
and rupture.
The last aim of this research involves the role of DDR1 in the remodeling of the
extracellular matrix (ECM). The ECM contains three morphological forms of collagen;
monomeric (M), semi-polymeric (SP), and fibrillar (F). As cells secrete M collagen, it
forms a polymeric intermediate SP form, and eventually composes the fibrillar F form.
The results of the study indicate the preferred ligand for DDR1 is the intermediate SP
form of collagen type I in addition to the known F form. These results provide a better
understanding of morphological collagen usage-initially during collagen processing in the
ECM in response to vascular damage, and their interactions with DDRs.
iii
Dedication
Pan jest moim pasterzem, nie brak mi niczego.
iv
Acknowledgments
I would like to express my gratitude to Dr. Chuck Brooks for his guidance and
kind discussions. I honestly could have never imagined someone could support and listen
to me as he has. I would also like to thank the members of my graduate committee for
serving-Dr. Dennis Bong and Dr. Andre Palmer.
Many people have assisted me through this journey. Most notably those
individuals in the graduate school: Dr. Elliot Slotnick, Shari Breckenridge, and Tim
Watson. There were also many professors and instructors who were willing to assist with
my teaching, research discussions, and anything else. Being too many to mention here, if
you are reading this, know that I am thankful to you all. In addition, I cannot thank those
individuals that I have met along the way for general support and mostly prayers. Please
know that you will always be thought of for your generosity.
I would like to thank Peter and Kathie for continually providing for free-loading
relatives.
Thank you, Jim, for making sure that you always had time for me and
appreciating my favorite word. As I always say, I will miss you most when I leave
Columbus, but leaving is never good-bye. You always say you are a phone call away. I
know you will call me back when I need you.
v
Finally, there were many things that I lost in the fire. Masich, you were one of
them. I knew this journey would be difficult and it just got too hard, too krazy, too
everything. For that I will always be very, very sad and very, very sorry. Please know that
I will eternally be grateful for your thoughts, your prayers, and your assistance. The
completion of this dissertation is the end of a long chapter. I am sorry that you were not
here for the end of it. Thank you for giving me the support to finish. I know you are
happy for me, and proud of my accomplishment.
I would like to thank The Ohio State University, the Ohio State Biochemistry
Program, the National Institute of Health, and the National Science Foundation for
fellowship funding in obtaining this degree.
vi
Vita
July 9, 1966 ....................................................Born-Lakefield, MN
1988................................................................B.S. Physiology, South Dakota State University
1991................................................................M.A. Physiology, Adelphi University
1991-1992 .....................................................Research Assistant, University of Toledo
1992-1994 .....................................................Research Assistant, University of Illinois,
Chicago
1994-1996 ......................................................Research Associate, University of South
Carolina
1996-1998 ......................................................Organic Chemist, Fluorochem, USA
1998-2004 ......................................................Graduate Research Associate, The Ohio State
University
2004-2006 ............................................................ Research Associate, The Ohio State University
2006-Present ........................................................ Graduate Research Fellow, The Ohio State
University
Parts of the work presented in this dissertation were performed by others as follows:
Chapter II
Portions of the data from Figures 2.4 and 2.5 were designed, collected, and evaluated by
Dr. Gunjan Agarwal and Dr. Cosmin Mihai.
vii
Fields of Study
Major Field: Biochemistry
viii
Table of Contents
Abstract ............................................................................................................................... ii Dedication .......................................................................................................................... iv Acknowledgments............................................................................................................... v Vita.................................................................................................................................... vii List of Tables ................................................................................................................... xiii List of Figures ................................................................................................................... xv
CHAPTER 1:
Discoidin Domain Receptor Proteins, Ligands Overview
Overview ..............................................................................................................................
Introduction ..................................................................................................................... 1
Receptor Tyrosine Kinases.............................................................................................. 1 RTK Families and Structures .......................................................................... 2
Discoidin Domain Receptor Proteins (DDRs) ................................................................ 4 Expression of DDRs ........................................................................................ 5 Isoforms of DDRs ........................................................................................... 5 Structure of DDRs ........................................................................................... 8
Ligands for Phosphorylation/Activation of DDRs ........................................................ 12
DDRs bind to Collagen ................................................................................. 12
ix
DDRs bind to phospholipids ......................................................................... 14
Downstream signal cascade........................................................................... 15
Collagen chemistry ........................................................................................ 17
Phospholipid chemistry ................................................................................. 22
The Role of DDRs in Atherosclerosis and Arteriol Wound Healing ............................ 25
Thesis Aims ................................................................................................................... 28
CHAPTER 2:
Functional Analysis of Discoidin Domain Receptors Binding to
Lipids and collagen
Introduction ................................................................................................................... 31
Materials and Methods ......................................................................................................
Materials ........................................................................................................ 34
Sequence Alignment...................................................................................... 34
Preperation of Phospholipid Vesicles (Liposomes) ...................................... 35
Flow Cytometry Binding Assay .................................................................... 37
Surface Plasmon Resonance .......................................................................... 37
DDR1 Phosphorylation Studies..................................................................... 38
Vesicle Binding Assay .................................................................................. 39
SDS PAGE and Western Blotting ................................................................. 39
Results
..........................................................................................................................
Evaluation of Amino Acid Sequences........................................................... 40
Phase Contrast and Fluorescent Microscopy................................................. 40
Flow Cytometry ............................................................................................. 43
Surface Plasmon Resonance Evaluation ....................................................... 46
x
DDR1 Phosphorylation ................................................................................. 48
Vesicle Binding Assay .................................................................................. 49
Discussion ..................................................................................................................... 50
CHAPTER 3:
Structural Prediction of Discoidin Domain receptors (DDR1 and
DDR2): A Potential Lipid Binding Protein
Introduction ................................................................................................................... 56
Methods ...........................................................................................................................
Sequence Alignment...................................................................................... 58
Model Construction ....................................................................................... 59
Simulation Setup ........................................................................................... 60
Results
........................................................................................................................
Choice of Tenplate Structures and Alignment .............................................. 61
Evaluation of Amino Acid Sequences........................................................... 63
Global Aspects of the Simulation .................................................................. 66
The Models .................................................................................................... 78
Discussion ..................................................................................................................... 79
CHAPTER 4:
DDR1 and DDR2 Affect Platelet-Collagen Interaction
Introduction ................................................................................................................... 82
Materials and Methods ......................................................................................................
Reagents/Materials ........................................................................................ 87
Platelet Draw ................................................................................................. 88
SDS PAGE and Western Blotting ................................................................. 89
Collagen Preperation ..................................................................................... 89
xi
Platelet Aggregation ...................................................................................... 90
Statistical analysis ......................................................................................... 90
Results
.........................................................................................................................
Platelets do not Express DDRs ...................................................................... 92
Determining Platelet Aggregation ................................................................. 94
DDR1 and DDR2 Delay Time to Aggregation ............................................. 96
Platelets do not Aggregate in the Presence of DDRs .................................. 100
Discussion ................................................................................................................... 103
CHAPTER 5:
Discoidin Domain Receptor 1 recognizes a Morphological Form
of Collagen Type I
Introduction ................................................................................................................. 107
Materials and Methods ......................................................................................................
Ragents and Material ................................................................................... 109
Collagen Preperation ................................................................................... 109
Cell Culture ................................................................................................. 110
SDS PAGE and Western Blotting ............................................................... 110
Results
.........................................................................................................................
DDR1 Becomes Phopshorylated with Morphological Forms of Collagen . 111
Time to Phosphorylation ............................................................................. 111
Phosphorylation is not Concentration Dependent ....................................... 113
DDR1 Gene Silencing ................................................................................. 114
Discussion
............................................................................................................. 116
xii
CHAPTER 6:
Discussion
Conclusions and Discussions
............................................................................................................. 120
xiii
List of Tables
Table
Page
3.1. The PDB Identification numbers ............................................................................... 65
3.2. The % Identity of DDR1 ............................................................................................ 68 3.3. The % Identity of DDR2 ............................................................................................ 68 xiv
List of Figures
Figure
Page
1.1. Schematic Representation of Discoidin Domains in RTKs ......................................... 3
1.2. Slice Variants of DDR1 ............................................................................................... 7 1.3. The Family of Proteins which contain Discoidin Domains ......................................... 9 1.4. Ribbon Diagram of the DDR2 Discoidin Domain ..................................................... 11 1.5. DDR Proposed Signaling Network ............................................................................ 16 1.6. The Two Steps of Collagen Synthesis ....................................................................... 20 1.7. Schematic Representation of Collagen Molecule Assembly ..................................... 21 1.8. Self Assembly of Phospholipids ................................................................................ 23 1.9. Synthesis of Phospholipids ........................................................................................ 24 1.10. Formation of a Thrombus ........................................................................................ 27 2.1. Amino Acid Alignment.............................................................................................. 42 2.2. Cell Microscopy ......................................................................................................... 44 2.3. Labeled Phospholipids bound to Cells ....................................................................... 45 2.4. Flow Cytometry ......................................................................................................... 47 2.5. Surface Plasmon Resonance ...................................................................................... 49 2.6. SDS PAGE and Western Blotting ............................................................................. 51 2.7. Western Blotting with anti-phosphotyrosine ............................................................. 53 xv
3.1. Amino Acid Alignment of DDR1 and DDR2............................................................ 67 3.2. Root-Mean Square Deviations (RMSDs) .................................................................. 70 3.3. Calculated B-factors................................................................................................... 72 3.4. Cα Distance ................................................................................................................ 73 3.5. Conformational Differences in modeled DDR1 and DDR2 ...................................... 74 3.6 Solvated images of DDR1 .......................................................................................... 76 3.7. Solvated images of DDR2 ......................................................................................... 78 3.8. Molecular Model of DDR1 ........................................................................................ 80 3.9. Molecular Model of DDR2 ........................................................................................ 81 4.1. Blood Coagulation Pathway ...................................................................................... 88 4.2. Western Blots of Washed Platelets ............................................................................ 96 4.3. Platelet aggregationwith DDRs and morphological collagen .................................... 97 4.4. Platelet aggregation in response to collagen .............................................................. 99 4.5. DDR1 aggregation lag time ..................................................................................... 102 4.6. DDR2 aggregation lag time ..................................................................................... 103 4.7. Comparison of lag times .......................................................................................... 104 4.8. Aggregation with collagens incubated with DDRs .................................................. 105 4.9. Aggregation with washed collagens ........................................................................ 106 5.1. Phosphorylation of morphological forms of collagen ............................................. 116 5.2. Time to phosphorylation .......................................................................................... 117 5.3. Phosphorylation of DDR1 dependent on collagen concentration ............................ 118 5.4. DDR1 siRNA ........................................................................................................... 119 xvi
CHAPTER 1
DISCOIDIN DOMAIN RECEPTORS, LIGANDS OVERVIEW
1.1
INTRODUCTION
Living cells must incorporate multiple extracellular signals into purposeful
responses. To arrange these responses in a coordinated manner, a special class of cell
surface receptors exists that bind extracellular ligands such as growth factors,
phospholipids, and collagens; following binding, a signal is transmitted through the cell
membrane into the cytoplasm. Many of these receptors belong to the receptor tyrosine
kinase (RTK) family. RTKs are characterized by an extracellular ligand binding domain,
a single transmembrane domain, a catalytic tyrosine kinase domain, and additional
regulatory sequences also subject to phosphorylation. Approximately 60 RTKs have been
identified to date, and can be divided into 20 subfamilies defined by the structure of the
extracellular domain, which determines ligand specificity (5,6).
1.2
RECEPTOR TYROSINE KINASES
Most RTKs show a high affinity for polypeptide growth factors, cytokines, and
hormones. RTKs have also been shown to be key regulators of normal cellular processes,
including the development and progression of many forms of cancer (7). The basic
1
scheme of RTK activation and function occurs when a growth factor binds to the RTK
extracellular domain (ECD) and induces a reorientation of the receptor in the plasma
membrane. This reorientation leads to autophosphorylation within not only the catalytic
domain, leading to activation and potentiation of kinase activity, but also phosphorylation
of noncatalytic regions of the cytoplasmic domain, creating docking sites for downstream
cytoplasmic targets that are biologically diverse-such as SH2 and SH3 domain proteins.
1.2.1
RTK Families and Structures
Several RTK family members share various common features, but the detailed
mechanisms of family members are unique. Most RTK receptors are found as single,
ligand-free dimers. In some cases, RTKs can dimerize through ligand binding.(8). Each
monomer has an extracellular N-terminal ligand binding region (ECD), a 25-40 residue
single-hydrophobic trans-membrane spanning domain, and an intracellular C-terminal
catalytic region. Figure 1.1 represents several of the RTK families.
The Epidermal Growth Factor Receptor (EGFR) family is a family of four
structurally related RTKs, all of which exist on the cell surface and bind epidermal
growth factors, which in turn, cause EGFR activation. EGFR exists as an inactive
monomer, turning into an active homodimer upon ligand binding. Dimerization leads to
stimulation of the intrinsic protein kinase. As a result, autophosphorylation of several
tyrosine (Y) residues in the C-terminal occurs. This autophosphorylation elicits
downstream activation and signaling of several proteins associated with the
phosphorylated tyrosine(s). These downstream signaling proteins initiate signal
transduction cascades which control cell migration, proliferation, and adhesion (9,10).
2
Figure 1.1. Schematic representation of domain components of several RTK subfamilies.
Abbreviations are as follows: EGFR, epidermal growth factor; FGFR, fibroblast growth
factor receptor; TrkB, tyrosine kinase receptor B (4).
Mutations on EGFRs lead to several mutations which are linked to lung cancers, anal
cancers, and glioblastomas (11).
The Fibroblast Growth Factor Receptor (FGFR) family, as the name implies,
binds to fibroblast growth factors. FGFRs consist of an extracellular ligand-binding
region, composed of three immunoglobulin-like domains, a single transmembrane helix
domain, and a cytoplasmic domain with tyrosine kinase activity (12). When the
extracellular portion of the protein interacts with the fibroblast growth factor ligand, a
3
cascade of down-stream events occur which controls cell mitogenesis and cell migration.
The FGFR family binds both acidic and basic growth factors which are involved in limb
induction. Mutations in this family of genes leads to cleft lip and palate (13). Mutations
also lead to squamous cell lung cancer, and stem cell leukemia lymphoma syndrome (14).
Vascular Endothelial Growth Factor Receptors (VEGFR) are a family of RTKs
which bind vascular endothelial growth factors-often involved in vasculogenesis
(formation of the circulatory system), and angiogenesis (growth of blood vessels and/or
new vasculature) (15). VEGFRs have an N-terminal extracellular portion consisting of 7
immunoglobulin-like domains, a single transmembrane spanning domain, and a Cterminal intracellular portion containing a split tyrosine-kinase domain. (16). Upon
binding a VEGF, the receptor dimerizes, leading to a signal transduction cascade.
The TrkB tyrosine kinase receptor, also known as the neurotrophic tyrosine
kinase, posses a high affinity for neurotrophins, which are small growth factors that
induce the survival and differentiation of neuronal cells. TrkB is characterized by having
2 immunoglobulin-like domains, as well as a leucine-, and cysteine-rich binding region in
its C-terminal. Upon binding of a ligand, dimerization takes place and induces
autophosphrylation. Mutations in this gene have been associated with associated mental
health issues, mental retardation, and some cancers (17,18). In addition, this RTK is
commonly used as a negative control, as it posses an intercellular kinase region, but no
discoidin domain (4).
4
1.3
DISCOIDIN DOMAIN RECEPTOR PROTEINS (DDRs)
Discoidin domain receptors are members of the RTK superfamily. During the
search for tyrosine kinase proteins expressed in human cancers, a novel subfamily of
RTKs was discovered. This subfamily is distinct from other members of the RTK group
due to a homology domain to discoidin, a lectin (sugar-binding protein) found in the
slime mold Dictyostelium discoideum (19,20). Two members of this subfamily are
present in the human genome, DDR1 and DDR2. Unlike most other RTKs, DDRs are not
activated by soluble growth factors; instead, various types of collagen act as ligands for
DDRs (21). This implies that the DDRs may respond to molecules in the local
environment rather than soluble signaling molecules.
1.3.1 Expression of DDRs
DDRs are found in multiple human and mouse cell types (1). DDR1 is widely
expressed during development and in adult tissues such as the kidney, gut, brain, cornea
and skin epithelium; and in corneal (keratinocytes) and dermal fibroblasts (22). In the
pancreas, DDR1 expression is limited to the Islets of Langerhans (23). The expression of
discoidin receptors is regulated. DDR1 mRNA expression in monocytes is upregulated by
tumor necrosis factor α, interleukin-1 β, granulocyte macrophage-colony stimulating
factor (GM-CSF), and lipopolysaccharides (LPS) (22). Very little is known about
regulation in other cell types or tissues. DDR2 expression is more limited, but expression
is found in skeletal muscle, heart, blood vessels, and connective tissues (1).
DDR1 is overexpressed in human tumors, in particular, breast cancer (24),
ovarian cancer (25), and pediatric brain cancer (26). DDR1 is also found overexpressed
5
in highly invasive tumor cells, whereas DDR2 is detected only in the surrounding stromal
cells. This would suggest an involvement of DDR1 and DDR2 in tumor progression.
1.3.2 Isoforms of DDRs
Human DDR1 is located on chromosome 6p21.3, in close proximity to HLA
(human leukocyte antigen) genes, belonging to the telomeric region (class 1) of the major
histocompatibility complex (27). The juxtamembrane regions in DDR1 and DDR2 are
much longer than in other RTKs (176 and 147 amino acids, respectively). In addition, the
ECD of DDR1 is shed which results in a 63-kDa membrane anchored β-subunit and a 54kDa soluble α-unit (23). To date, five isoforms of DDR1 have been cloned as a result of
alternative splicing, designated with the suffixes “a” to “e” (28). The longest transcript
codes for DDR1c and translates to a 919 amino acid residue protein. The b isoform lacks
6 amino acid residues inserted into the kinase domain between exons 13 and 14 (23). The
a, d, and e isoforms are formed through alternative splicing in the juxtamembrane region.
The deletion of exon 11 coding for 37 amino acids generates DDR1a, and the deletion of
exons 11 and 12 results in DDR1d. In DDR1e, the first half of exon 10 and all of exons
11 and 12 are missing (28). DDR1d and DDR1e are truncated variants that lack either the
entire kinase region, or parts of the juxtamembrane region and the ATP binding site.
DDR1a retains the reading frame and is therefore an active kinase. The coding sequence
of DDR1d and DDR1e goes out of frame and because of this, makes both isoforms kinase
dead (Fig. 1.2). DDR1b is the predominate form found during embryogenesis. DDR1a is
more common in human mammary carcinoma cell lines (29).
6
Figure 1.2. Schematic representation of the different forms of DDR1. Upper
panel demonstrates the organization of the genomic locus of DDR1 between
exons 9 and 14. The various splicing events which result in previously known
isoforms DDR1a, DDR1b, and DDR1c, and novel isoforms DDR1d and DDR1e,
are depicted under the genomic locus. Lower panel depicts length, molecular
weight, and overall domain structure of the different isoforms (24).
Although the splice variants for DDR2 are not yet characterized, there is limited evidence
to suggest that they do exist. Using human smooth muscle cells, several protein isoforms
of DDR2 have been detected at different molecular weights-130, 90, 50, and 45 kDa (23).
In addition, two transcripts at 9.5 and 4.5 kb have been discovered (30). There have also
7
been multiple transcripts for DDR2 identified in human cancerous cell lines, as well as
normal human cells (31,32).
1.3.3 Structure of DDRs
Both DDRs are composed of an N-terminal, approximately 160 amino acid
residue long discoidin domain (DD), followed by a “stalk” region; a sequence of
approximately 220 amino acids unique to the DDRs. This entire region is referred to as
the extracellular domain (ECD). Following the single-span transmembrane (TM) region
is a large cytosolic juxtamembrane domain, followed by a C-terminal catalytic tyrosine
kinase domain (33). DDR1 and DDR2 show 58% homology to one another in the DD,
and only 44% homology in the “stalk” region (33). Database searches have identified
three sequences with homology to DDR1 and DDR2 in the genomes of the nematodes
Caenorhabditis elegans, and Caenorhabditis briggsae (34). All three proteins show
common structural features to the DDRs. However, the ligands of these proteins have not
been determined, and it has not been determined if the RTK is an active kinase upon
ligand binding (1). Two other mammalian, non-RTK proteins are the neuropilins and the
neurexins (Figure 1.3). These proteins bind VEGF and are involved in the development
of the nervous system. Both proteins are transmembrane spanning, but do not contain an
RTK domain. DDs are found in a number of secreted proteins, such as the blood
coagulation factors V and VIII. Factor V and VIII bind to phospholipids at the surface of
platelets (35). More than a dozen other mammalian transmembrane and secreted proteins
are known to incorporate DDs into their sequences.
8
Figure 1.3. The family of proteins which contain DDs. Abbreviations: F-A, A-domain
in blood coagulation factor V and viii; CBP, carboxypeptidase; EGF, epidermal
growth factor, TK, tyrosine kinase; A5, homology to A5 antigen; Ig, immunoglobulin;
LamG; laminin-G; FIB, fibronectin-like; CUB, complement binding; and MAM,
meprin/A5/PTPmu. Tyrosines in the N-P-X-Y motives of GCTK and DDR1b are
highlighted. MMP processing of DDR1 is indicated (1).
The Three-dimensional solution structure of the DDR2 DD domain was resolved
using standard heteronuclear multidimensional NMR techniques (36). The overall fold
consisted of 8 β-strands, β1-β8, which are arranged into two antiparallel sheets of five
(β1-β2-β7-β4-β5) and three (β8-β3-β6) strands packed against each other (Fig. 1.4). This
noted β-barrel structure is common amongst other DD proteins: blood coagulation factors
V and VIII C2 domains, neuropilin-1 (Npn-1) b1 domain, and lactadherin (MFG-E8) (379
39). The compact β-barrel possess four flexible loops (“spikes”), forming a ligand
binding surface (40). L1, L2, and L4 were found to be different from those of other
resolved DD domain structures in their length and conformation. The “bottom” of the βbarrel core is closed by three interconnecting segments. “Bottom” is referred to region
against the plasma membrane-the non-binding area of the protein. The N- and C-termini
are connected by a disulfide bond between Cys30 and Cys185 residues (Fig. 1.4).
The collagen binding site of DDR2 is contained solely in the DD region (amino
acids 30-182) (3,41). Using residues 26-186, as well as transferred cross-saturation
measurements (TCS) in the DD of DDR2 purified from Pichia pastoris, loops L1, L3,
L4, and L6 on the ‘top’ of the DDR2-DD domain (Fig. 1.4) are in the proximity of
fibrillar collagen type II (36). These loops form a trench to accommodate the triple helix
of the collagen fiber. The structure of the DDR1 DD domain has not yet been resolved.
Models have been constructed utilizing the known crystal structures of blood coagulation
factors V and VIII (3,33). Loops L1 and L3 were found to be essential for fibrillar
collagen type II binding in these models.
To date, a number of crystal structures of DD domains do exist in various
proteins, but none of those are from the RTK family. Those structures include sugar
recognizing lectins (sugar binding proteins) (42-44), as previously mentioned, the C2
domains of blood coagulation factors V and VIII ((37,38), and the b1 domain of the
VEGF binding protein, Npn-1 (45,46). Although these DD domains show a high
similarity in their overall fold, the differences in their loop regions (L1, L2, L3, L4, and
L6), lead to the recognition of a wide variety of ligands, as noted in the literature (47).
10
The DD domains of coagulation factors V and VIII bind to membranes containing
phosphatidyl-L-serine
Figure 1.4. Ribbon diagram of the DDR2 DD domain. L1-L6 refers to loops 1-6.
Yellow bands demonstrate connecting disulfide bonds (27).
11
(PLS). Both proteins posses long hydrophobic loops which might be suitable for insertion
into the lipid bilayer. The hydrophilic and charged residues in the deep pockets formed
by the loops may contribute to the recognition of the hydrophilic form of PLS (37,48).
1. 4
LIGANDS FOR PHOSPHORYLATION/ACTIVATION OF DDRs
Unlike most other RTKs, the binding of soluble growth factors to the DDRs does
not induce phsphorylation. Since no studies exist characterizing the activation of the
tyrosine kinase region, it is assumed that a lack of phosphorylation does not lead to the
activation of the kinase region. As mentioned, RTKs bind to and are activated by growth
factors. Instead, various types of collagens act as ligands for the DDRs. The idea that
collagens function as ligands for the DDRs suggests that some of the other mammalian
DD domains may interact with matrix proteins as well. DDRs are phosphorylated only
when collagen is in its native, triple-helical form (21). It has been found that heatdenatured collagen (gelatin), which lacks triple helical structure, fails to induce
phosphorylation of the DDRs. Currently; collagens are the only known ligands for the
DDRs.
1.4.1
DDRs Bind to Collagen
DDR1, as well as DDR2, becomes phosphorylated by all collagens which have
been tested so far (type I – type V) (4,21). DDR1 binds to non-fibrillar collagen type IV,
while DDR2 binds to non-fibrillar type X (49). The phosphorylation process is slow,
requiring collagen treatment for 18 hours to reach maximal phsophorylation.
Phosphorylation of the RTK is maintained with no significant down-regulation by
12
endocytosis or receptor degradation for up to 4 days. The native, triple helical
confirmation (monomeric form) of collagen is essential for DDR1 and DDR2
phosphorylation. Collagen must be glycosylated for DDR2 phosphorylation (21).
The mechanism by which collagen binds to DDRs has not been completely
characterized. It has been established that the DD of both proteins is essential for ligand
binding (3). It is reported that DDR2 binds to three sites on a collagen type I molecule
(50). Little is known in regards to the DDR1 interaction sites on collagen (33). However,
it has been determined that the DDR2 binding site is confined to the D2 domain of
collagen type II. Amino acids 394-405 on the collagen type II D2 domain have the
highest binding affinity for DDR2 (41,51).
The significant points about the interaction of DDRs with collagen are: (a) The
entire ECD of DDR1 is essential for collagen binding. The DD of DDR2 is acceptable for
collagen binding; (b) DDRs require dimerization for interacting with collagen; and (c)
phosphorylation of DDRs by collagen is slow (hours), compared to phosphorylation of
other RTKs by their ligands (4).
Little is known in regards to the specifics of DDRs phosphorylation, and the
interactions involved in the downstream signaling pathway. Mutational analysis of DDR1
binding to collagen shows that while collagen may bind to the protein, binding may not
lead to phosphorylation of the tyrosine kinase region (52). And phosphorylation of the
kinase region may not lead to activation of the protein.
13
1.4.2 DDRs Bind to Phospholipids
A small portion of research has been devoted to the lipid binding properties of
DDRs. The binding of DDR1 to negatively charged phospholipids exhibits properties
similar to those observed for binding to the ionic sites of fixed D. discoideum cells (53).
The DDR1-vesicle interaction results from direct electrostatic interaction of the
negatively charged phospholipid vesicle surface and the positively charged domain of the
DDR1 protein (54). Lowering the temperature to 4°C results in a reduction of DDR1
proteins binding to phospholipid vesicles. This would suggest there is a requirement for
vesicle bilayer fluidity that needs to occur for maximal binding of the protein to the
vesicle (55). Binding of the protein to the lipid vesicle also occurs in a cooperative
manner. It was determined that increased concentrations of DDR1 allowed for the
clumping (agglutination-the coalescing of small particles suspended in solution) of
negatively charged phospholipids in solution. The concentration of DDR1 protein
required for phospholipid vesicle agglutination is less than the estimated cytosolic
concentration required for D. discoideum cells to adhere to one another in a natural
environment (53).
The DD domain of the DDRs demonstrates a high level of functional diversity
and similarity. The DD region is homologous to the C2 DD domains found in blood
coagulation factors V and VIII, the milk protein lactadherin (MFG-E8), and the VEGF
protein, neuropilin-1 (Npn-1). It is known that these soluble proteins (factors V, VIII, and
lactadherin) bind phospholipids on the surface of cells, particularly platelets, and it is
concluded that they may posses cell adhesion properties, or control aggregation events
14
such as vasculogenesis and/or angiogenesis (35). There is a high sequence similarity
between the DD (discoidin domains) of factors V, VIII, lactadherin, and the DD domains
of the DDRs. It is interesting to examine the aligned sequences to gain clues as to which
residues, if any, are involved in the binding of ligands-specifically phospholipids.
1.4.3
Downstream Signal Cascades for DDR-induced Signaling
DDR1 and DDR2 have a total of 15 and 13 tyrosine residues in their cytoplasmic
regions, respectively. These residues serve as potential tyrosine kinase phosphorylation
sites upon receptor activation by collagen. Using a technique such as phosphopeptide
mapping, 3 major and 5 minor phosphorylation sites were recently identified in DDR1
(56). Phosphorylation of these tyrosine residues could possibly lead to the binding of a
number of different Src-homology-2 (SH2)- and phsophotyrosine binding (PTB) domaincontaining proteins (Fig. 1.5). Tyrosine residue 513, found in the alternative splice
product DDR1b, associates with the PTB domain of ShcA upon phosphorylation (21). In
macrophages, DDR1 induced ShcA (Src homology 2/α collagen) phosphorylation leads
to activation of the TRAF6 (TNF receptor associated factor 6) complex, which triggers
p38 mitogen-activated protein kinase and NFκβ pathways ((57). DDR1b contains the
amino acid motif LLXNPXY that associates with the phosphotyrosine binding domain of
the ShcA adapter protein upon collagen induced tyrosine phosphorylation (21). The
DDR1a juxtamembrane region binds to fibroblast-growth factor receptor 2 and triggers
the migration and psuedopod extension of leukocytes (22,58). Several other proteins were
also found to be affected by DDR1 phosphorylation: Shp-2, an SH2 domain-containing
phosphotyrosine phosphatase, and Nck2, an SH2- and SH3 domain containing adaptor
15
protein (56,59). The binding site for Shp-2 is mapped to tyrosine 740 of the DDR1 RTK
region. Subunit p85 of phosphatidyl-inositol-3 kinase is mediated by tyrosine 880 (60).
Figure 1.5. DDR proposed signaling network. Schematic representation of signaling
molecules downstream of DDR1 and DDR2. Red arrows indicate a direct association of
tyrosine residue and protein molecule. Gray arrows represent and indirect interaction
(18).
The signaling of DDR2 by fibrillar collagen requires the presence of ShcA and
the Src-like tyrosine kinase (61). Src is hypothesized to be a partner which allows full
16
phosphorylation of DDR2. Similar to DDR1, ShcA binds to the juxtamembrane region of
DDR2, which contains tyrosine 471. This interaction is mediated by the SH2 domain of
the ShcA (62). Unlike DDR1, the SH2 domain, rather than the PTB domain of ShcA, is
mediating the protein interaction. Little is known in regards to specific transcriptional
targets of active DDRs. There is evidence that would suggest DDR signaling increases
the expression of P-selectin glycoprotein ligands and decreases the levels of matrix
proteins such as agrin or syndecan-1 (63). One gap of information absent in this field is
the data necessary to connect the signaling pathways of the DDRs to specific
transcriptional responses.
Prolonged
activation
of
both
DDR1
and
DDR2
upregulates
matrix
metalloproteinases (MMPs 1, 2, and 9). MMPs cleave the extracellular matrix (30,64).
Phosphorylation of DDR1 is known to increase the expression of collagen types I and III
α-chains and of integrin α2. DDR1 also mediates the p53 induced Ras/Raf/MAPK
signaling cascade through interaction with the Ras effector protein (65). Phosphorylated
DDR1 initiates recruitment and activation of plasma associated PI3 kinase. This is
believed to increase breast cancer tumor cells adhesion to one another, as well as
minimize tumor proliferation (66).
1.4.4 Collagen Chemistry
To date, 19 different collagens have been described. Collagen type I, II, III, V,
and XI are widespread and tend to form fibers or sheet-like structures. An increase in
fibrillar collagens leads to a variety of human conditions such as fibrosis, vascular
disease, and tumor angiogenesis (67). Degradation of collagen fibrils possibly leads to
17
metastasis of cancerous cells. Collagen fibers play a role in the extracellular matrix
mechanical properties, which parallel conditions in the arterial vasculature, elastic
extension in skin, and elastic compression in articular cartilage (68). Collagen fibers are
subjected to mechanical stresses on the cells by cell surface proteins (69). Cellular
response are also mediated by the “stiffness” of the cellular matrix in which they move
(64). The most abundant type of collagen is collagen type I. Type I is found throughout
the body, with the exception of cartilaginous tissues. It is the most widely studied
collagen and serves as a model to understand interactions and properties of other types of
collagens. Other forms of collagen appear to regulate the size of fibers or have a more
restricted expression pattern in specialized cells and tissues (70).
Collagen type I is distinct from other proteins in that the molecule is comprised of
three polypeptide α-chains which form a unique triple helical structure. For collagen type
I, two α1(I) chains and one α2(I) comprise a fibril . For these chains to wind into a triple
helix, they must have a glycine (G) at every third residue, since G is the smallest amino
acid (71). Each of the three chains has, therefore, a repeating structure G-X-Y, where X
and Y can be any amino acid, but frequently is the immno acids proline and
hydroxyproline.
Type I collagen {[α1(I)]2α2(I)} is found throughout the body and is synthesized in
response to injury and production of fibrotic nodules found in fibrotic disease. The
synthesis of this collagen occurs in two steps. The first step produces a folded, triplehelical, soluble procollagen molecule that is secreted from the cell. This process occurs
by the α-chains being synthesized as they concurrently undergo cotranslational
18
hydroxylation of the proline and lysine residues. The nascent α-chains are subjected to
glycosylation at the hydroxylysyl residues. Subsequently, the hydroxylation of peptidyl
proline induces conformational changes in the peptide backbone that promotes formation
of the triple helix. The assembly of the pro-α-chains into procollagen starts after the
association of the C-pro-peptides from the three α-chains and the triple helix folding
begins to propagate into in a C to N direction via a zipper-like action. Formation of the
triple helix prevents further hydroxylation of the procollagen (Fig. 1.6) (72). The
condensed structure of this molecule confers a high stability and makes the molecule
resistant to cleavage by proteases such as pepsin, trypsin, and chymotrypsin, but not
collagenase.
The second stage of collagen synthesis includes the extracellular steps of
proteolytic conversion of procollagen to collagen and its polymerization into fibrils, a
process known as fibrillogenesis (Fig. 1.6). The newly formed procollagen molecules are
excreted from the cell into the extracellular matrix (ECM), and undergo cleavage of the N
and C pro-peptides by the procollagen N and C proteinases. The resulting collagen is rodlike (300 nm x 1.5 nm) and spontaneously self assembles to form fibrils. The assembly of
collagen molecules into fibrils is an entropy-driven process, similar to that occur in other
protein self-assembly systems, such as microtubules, actin filaments, and flagella (73).
The major driving force behind collagen fibrillogenesis is the hydrophobic interaction
between the collagen molecules leading to a minimization of the surface exposed to
water. Collagen fibers (fibrils which are longer and thicker) that are formed either in vitro
or in vivo are characterized by a specific organization of collagen molecules. These
19
molecules are staggered by integrals of D with respect to one another; where D is
approximately 67 nm (74). The length of collagen monomers is approximately 300 nm,
and can stagger in five possible configurations starting from full overlap (0 D stagger) to
Figure 1.6. The two steps of collagen synthesis. The first step produces a folded, triplehelical, soluble procollagen molecule that is secreted from the cell. The second stage of
collagen synthesis includes the extracellular steps of proteolytic conversion of
procollagen to collagen and its polymerization into fibrils, a process known as
fibrillogenesis (72).
only one overlap (4 D stagger). The collagen stagger combinations are all found during
fibrillogenesis. The 0 and 1 D stagger is predominately found during the latter stages of
20
fiber development. The higher stagger patterns 2, 3, and 4 D occur in the early stages of
fiber formation (Fig. 1.7). Fiber growth is limited to the interaction between the surface
of the outer layer of the fiber, and the incoming monomer (75).
New collagen monomers prefer to accumulate to the monomers at the fiber tip,
resulting in an abundance of 4 D staggers. Due to the entropic nature of this process, the
Figure 1.7. Schematic representation of collagen molecule assembly. The collagen
molecule is 4.5 D-periods in length. Collagen molecules are staggered by integrals of
D with respect to each other, as determined by scanning electron microscopy. There
are five specific ways in which two molecules of collagen can assemble with respect to
each other. These are defined as 0 D, 1 D, 2 D, 3 D, and 4 D staggers, dependent on the
length of the collagen overhang (92).
21
collagen molecules further minimize their surface area by contact with other collagen
molecules. This induces rearrangements in the fiber organization with the end effect that
0 and 1 D staggers will predominate and give the observed D periodicity of the mature
fiber. The fibers are further stabilized in vivo by lysyl oxidase cross-linking between the
collagen molecules.
1.4.5
Phospholipid Chemistry
The nature of lipid vesicles (also known as liposomes), as well as their relevance
in biological environments, has been extensively studied and applications have been
extended into many fields. Lipid vesicles are commonly used as precursors for the
fabrication of suspended layers (76), mimicking a plasma membrane.
Lipid vesicles are spherical soft-matter particles consisting of one or more bilayer
membranes, mostly comprised of phospholipids which spontaneously form a capsule
surrounding an aqueous fluid. The aqueous fluid is usually that in which the vesicle is
suspended-cytoplasm, blood plasma, buffer solution, etc. Lipid vesicles can be readily
prepared in the lab (77). In vesicle formation, dissolved phospholipids, consisting of a
hydrophilic head group and a hydrophobic tail, self-assemble into lipid droplets when
exposed to a polar environment. The water solubility of the lipids is very low, making the
free-energy decrease, which forces the lipid’s head groups to face the aqueous medium
and the hydrophilic hydrocarbon portion of the head group are forced to face each other
in the bilayer (76). Lipid vesicles are analogs of natural membranes and assemble
spontaneously from pure lipids or lipid mixtures. Figure 1.8 depicts the assembly process,
which leads to a unilameller vesicle (78).
22
The most commonly utilized lipids are the phospholipids, particularly chargeneutral phosphatidylcholine (L-α-PC) and negatively charged phosphatidylserine (PLS). Lα-PC is the most abundant phospholipid in animals and plants, accounting for 50% of
total lipid, and is a key building block of membrane bilayers. In particular, it makes up a
high proportion of the outer leaflet of the plasma membrane. L-α-PC is also the principal
Figure 1.8. Self assembly of phospholipids in aqueous environment. Blue regions are
hydrophilic head groups, with hydrophobic tail groups. Unilamellar vesicle formation
occurs spontaneously. Vesicles are assumed to mimic true plasma membranes.
phospholipid circulating in blood plasma, where it is an integral component of the
lipoproteins. There are several mechanisms for the biosynthesis of L-α-PC. Choline itself
23
is not synthesized in vivo and must be obtained by dietary sources. Once taken into cells,
choline is immediately phosphorylated by a choline kinase in the cytoplasm of the cell to
phosphocholine, which is reacted with cytidine triphosphate (CTP) to form cytidine
diphosphocholine.
The
membrane-bound
enzyme
CDP-choline:1,2-diacylglycerol
cholinephosphotransferase in the endoplasmic reticulum catalyses the reaction of the last
compound with sn-1,2-diacylglycerols to form L-α-PC (79) (Fig. 1.9).
Figure 1.9. Synthesis of phospholipids. Phosphatidylcholine is produced by choline
obtained in the diet. Choline is taken up by cells, where it is phosphorylated by choline
kinase. Final product synthesis occurs in the endoplasmic reticulum. Bottom portion of
scheme demonstrates the formation of phosphatidylerine (PLS) via
phosphatidylcholine
24
PLS, on the other hand, is less than 10% of total phospholipid, and between 10 –
20 mol% of the total phospholipid in the plasma membrane and endoplasmic reticulum of
the cell. PLS is an acidic (anionic) phospholipid with three ionizable groups, i.e. the
phosphate moiety, the amino group and the carboxyl function. PLS is located entirely on
the inner monolayer surface of the plasma membrane (and of other cellular membranes)
and it is the most abundant anionic phospholipid. This normal distribution is disturbed
during platelet activation and cellular apoptosis (80). In animal tissues, there are two
routes to PLS involving distinct enzymes (PS synthase I and II, and PS decarboxylase)
with different substrates and cellular locations. PLS is synthesized in the endoplasmic
reticulum of the cell, by an exchange reaction of L-serine with
L-α-PC
or
phosphatidylethanolamine, catalyzed by PS synthase I. This reaction is strictly dependent
on calcium ions and requires no further source of energy. The new lipid is then
transported to the mitochondria, where it is decarboxylated to phosphatidylethanolamine,
which returns to the endoplasmic reticulum and is converted back to PLS by the action of
PS synthase II (Fig. 1.9). PLS is involved in the blood coagulation process in platelets,
where it is transported to the plasma-oriented surface of membrane vesicles that are
derived from activated platelets. PLS is known to have an important role in the regulation
of apoptosis (programmed cell death). The normal distribution of this lipid on the inner
leaflet of the membrane bilayer is disrupted because of stimulation of the enzyme
scramblase, which can move phosphatidylserine in both directions across the membrane,
and inhibition of aminophospholipid translocases, which returns the lipid to the inner side
of the membrane (81).
25
1.5
The Role of DDRs in Atherosclerosis and Arteriole Wound Healing
Atherosclerosis is characterized by an increase in arteriole wall thickness as the
result of a build-up of fatty materials. It is a syndrome which affects the arterial blood
vessels, and a chronic inflammatory response in the walls of the arteries as a reaction to
the accumulation of macrophages. Atherosclerosis is caused by oxidized low-density
lipoproteins (plasma proteins that carry cholesterol and triglycerides) without adequate
removal of fats and cholesterol from the macrophages by functional high density
lipoproteins (HDL) (82). It is caused by the formation of multiple plaques within the
arteries. Atherosclerosis can remain asymptomatic for decades. Figure 1.10 is a depiction
of the progression of atherosclerosis. Atherosclerotic lesions, or atherosclerotic plaques,
are separated into two broad categories: Stable and unstable (also called vulnerable) (83).
The pathophysiology of atherosclerotic lesions is complicated, however, stable
atherosclerotic plaques, which tend to be asymptomatic, are rich in extracellular matrix
(collagen type I) and smooth muscle cells (84). Unstable plaques are rich in macrophages
and foam cells. The extracellular matrix separating the lesion from the arterial lumen of
the arterial (also known as the fibrous cap) is usually weak and prone to rupture. Ruptures
of the fibrous cap, expose material leading to thrombosis, such as collagen (85) to the
circulation and eventually induce thrombus formation (86) in the lumen. Upon formation,
intraluminal thrombi can occlude arteries completely (i.e. coronary occlusion), but more
often they detach, move into the circulation and eventually occlude smaller downstream
branches
causing
a
thromboembolism (i.e.
stroke).
Apart
thromboembolism, chronically expanding atherosclerotic lesions can
26
from
a
possible
Figure 1.10. Formation of a thrombus (blood clot). Plaque ruptures, exposing
smooth muscle cells and the extracellular matrix, which is rich in fibrillar
collagen. Exposed collagen activates plates via GPVI collagen receptor.
Platelet aggregates block flow of red blood cells.
cause complete closure of the lumen. Chronically expanding lesions are often
asymptomatic until stenosis of the lumen is so severe that blood supply to downstream
tissue(s) is insufficient, resulting in ischemia (87). These complications of advanced
atherosclerosis are chronic, slowly progressive, and cumulative. Most commonly, the soft
plaque suddenly ruptures, causing the formation of a thrombus (Fig. 1.10) that will
rapidly slow or stop blood flow, possibly leading to death of the tissues fed by the artery
in approximately 5 minutes. This catastrophic event is called an infarction. One of the
most common recognized scenarios is called a coronary thrombosis of a coronary artery,
27
causing a myocardial infarction (heart attack). The same process in an artery to the brain
is commonly called a stroke.
Very little is known about the function of DDRs in the cardiovascular system.
However,
in
atherosclerotic
tissue
from
non-human
primates
fed
with
a
hypercholesterolemic diet, both DDR1 and DDR2 were found to be highly expressed by
smooth muscle cells within the fibrous cap (30). The accumulation of collagen, which is a
marker of atherosclerotic plaque formation, was much less severe in DDR1-null versus
wild type mice following copper wire injury in mouse aorta. In addition, stretching
vascular smooth muscle cells in vitro resulted in upregulation of DDR2 expression, a
process dependent on TGF-β and angiotensin II signaling (88). It will be of great interest
to study the precise involvement of DDRs during the progression of atherosclerosis and
vascular injury.
1.6
THESIS AIMS
The focus of this work is to achieve a better understanding of DDR-ligand,
particularly phospholipid as a ligand, interaction and its physiological significance. To
obtain this understanding, we propose to test the following hypotheses:
-The ECD domain of the DDRs contains a DD motif which is similar to other
proteins on the cell surface. These other cell surface proteins bind phospholipids
via their DD motif. My hypothesis is: DD motifs in the DDRs bind phospholipids
based on the DD amino acid content and proposed motif structure (Fig. 1.4). To
28
test this hypothesis, we propose to obtain qualitative characterization of the DDR
ECD by determining the binding properties of phospholipids by using both in
vitro and cell based assays. We will find that those cells over-expressing DDRs
will bind phospholipids with a selectivity based on the protein and phospholipid
utilized.
-Three-dimensional structures of multiple phospholipid binding DD motifs have
been resolved in phospholipid binding proteins. My hypothesis is: the DD motif
of the DDRs will be structurally related to previously solved structures. To test
this hypothesis, we will employ amino acid comparison in coordination with
molecular dynamic simulations of a DD model threaded against previously solved
DD structures. The solved DD structure of the DDRs will provide insight into the
binding of phospholipids as a ligand.
-It has been extensively established that in response to vascular injury, platelets
adhere and aggregate to exposed collagen type I in the subendothelium of the
extracellular matrix. My hypothesis is: platelet aggregation is delayed by DDRs
disrupting normal collagen fibrillogenesis. In addition, DDRs will interfere with
platelet aggregation when DDRs bind to native collagen fibers. To test this
hypothesis, we will use platelet aggregometry in combination with several cell
based assays. Platelet aggregation will be delayed, or not occur at all, due to the
disruption of normal collagen fibrils by DDRs. The newly formed collagen will
29
not be recognized by normal platelet-collagen receptors such as the glycoprotein
(GPIV, GPVI, GPIa-IIa) series located on the surface of platelets.
-The fibrillar states of collagen define the mechanical properties of the ECM and
govern cell-matrix interaction. The interaction of the DDRs with collagen(s), and
their functional implications are not well understood. My hypothesis is: DDR1
binding and collagen-induced tyrosine phosphorylation is dependent on a specific
fibrillar morphology of collagen type 1: monomeric (M), semi-polymeric (SP), or
fibrillar (F). To test this hypothesis, we will use a series of biophysical and cellbased assays. This data will elucidate how DDR1 ECD binds to collagen and
affects DDR1 phosphorylation at the molecular level.
30
CHAPTER 2
FUNCTIONAL ANALYSIS OF DISCOIDIN DOMAIN RECEPTORS BINDING
TO LIPIDS AND COLLAGEN
2. 1
INTRODUCTION
Background
The cell membrane, also known as the plasma membrane, separates the interior of
the cell from its outside environment. The plasma membrane is selectively permeable to
organic molecules and controls the movement of substances into and out of the cell.
Plasma membranes are involved in a variety of cellular processes such as cell adhesion,
and cell signaling. The plasma membrane also serves as an attachment surface for the
extracellular matrix (ECM) in stabilizing cells in structures (89).
Peripheral membrane proteins are proteins which adhere temporarily to the
plasma membrane of the cell. Partners of these proteins can be bound to other cell
membrane proteins (integral proteins), or penetrate peripheral regions of the lipid bilayer.
The attachment of proteins to the plasma membrane has been shown to regulate cell
signaling and many other significant cellular events by multiple mechanisms (90).
Membrane binding may also promote rearrangement, dissociation, or conformational
changes within protein structural domains, resulting in phosphorylation and/or activation
of their biological activity (90,91). Multiple C2 domain containing proteins are involved
in membrane/phospholipid binding. The C2 domain is a protein structural domain
31
involved in targeting proteins to cell membranes. Those proteins are blood coagulation
factors V and VIII, and the milk fact globular protein, lactadherin.
It These soluble proteins (blood factor V and VIII, and lactadherin) bind
phospholipids on the surface of cells, particularly activated platelets, and it is concluded
that they may possess cell adhesion properties or control aggregation events such as
vasculogenesis and/or angiogenesis (35). Factor VIII binds via a stereoselective
interaction with the phospho-L-serine head group of phosphatidylserine (PLS). Factor V
also exhibits this same stereoselective interaction (92). Binding of factor VIII to
membranes is enhanced by not only the presence of phosphatidylethanolamine (PE) in
the membrane itself, but also by membrane curvature (93). The crystal structure of the C2
domains of factors V and VIII, and lactadherin provide the basis for a model lipidbinding mechanism (37-39). Discoidin domains are not true C2 domain proteins,
however, they are referred to as “C2-domain like”. There is currently no evolutionary
relationship between the C2 domain proteins and the discoidin domains. The C2 domains
consist of eight β-strands arranged in a compact β-barrel with four flexible loops
(“spikes”) forming a ligand binding surface (40). All three C2 domains (factor V, VIII,
and lactadherin) display two mainly hydrophobic spikes at the tip of the structure. The
spikes are responsible for the recognition of the binding partners, and in this case, the
binding partner would be phospholipid. The spikes are hypothesized to penetrate the
hydrophobic core of the phospholipid bilayer of the plasma membrane. Site-directed
mutagenesis of the hydrophobic residues in factor V of those residues on the spikes
32
hypothesized to be involved in phospholipid binding, demonstrated a much lower affinity
for the phospholipids themselves (94).
33
Hypothesis
Discoidin domain proteins (DDR1 and DDR2) show a sequence and hypothesized
structural similarity to the C2 domain proteins. Therefore I hypothesize that the DDRs
will demonstrate a marked propensity for binding phospholipids. To test this theory, I
propose to obtain a qualitative characterization of DDRs binding to select phospholipids
by utilizing cell based assays which will be performed using full-length DDR1 and
DDR2 proteins. Surface Plasmon Resonance (SPR) measurements will be performed
using purified ECD of DDRs, which contain the DD region similar to C2 domains on
phospholipid binding proteins factor V, VIII, and lactadherin. We surmise DDRs will
bind phospholipids, and that binding is selective amongst lipid and protein.
34
2.2
MATERIALS AND METHODS
2.2.1 Materials
Chinese Hamster Ovary (CHO), and Human Acute Monocytic Leukemia (THP1)
cells were purchased from American Type Culture Company (ATCC) (Manassas, VA).
FuGENE® 6 transfection reagent, and propidium iodide was purchased from Roche
Applied Science (Indianapolis, IN). Cell Lysis Buffer, and Myc-Tag antibody were
purchased from Cell Signaling Technology (Danvers, MA). Porcine brain L-αphosphatidylserine, Chicken egg L-α–phosphatidylcholine (L-α-PC), and Acyl 12:0 NBD
phosphatidylcholine were purchased from Avanti Polar Lipids, Inc. (Alabaster, AL).
4G10 anti-phosphotyrosine antibody was purchased from Millipore (Billerica, MA).
PureCol® Bovine dermal collagen Type I was purchased from Advanced BioMatrix (San
Diego, CA). Rabbit anti-DDR1 antibody sc-532, anti-mouse and anti-rabbit IgG, HRP
conjugated antibodies were purchased from Santa Cruz Biotech (Santa Cruz, CA). in
vitro experiments utilized DDR1- and DDR2-myc plasmids encoding the entire mouse
DDR1 or DDR2 sequence tagged with triple myc-tag and was from Regeneron
Pharmaceuticals (Tarrytown, NY) (4). Purified human DDR1 Fc chimera, DDR2 Fc
chimera, and TrkB Fc chimera were purchased from R&D Systems, Inc. (Minneapolis,
Mn).
2.2.2 Sequence Alignment
Sequences of the extracellular domain (ECD) of DDR1 (CAI18451), DDR2
(CAI15941), Factor V (NP_005919), Factor VIII (NP_000123), and Lactadherin
(NP_005919) from Homo sapiens were aligned using the CLUSTALW algorithm
35
(Blosum62 scoring matrix) in Bioedit. The alignment was edited using the GeneDoc
multiple alignment editor. The tilde character (~) was inserted to manually edit the
alignment. Loop regions relevant for protein-lipid interactions are represented by green
boxes. Conserved residues are highlighted in RED; partially conserved are in BLUE.
Sequence stretches forming the β-barrel are designated as S. Extra strands represented
with β. Hydrophobic residues implicated in lipid binding to factor V are indicated by *
and highlighted in YELLOW. Numbering of amino acid residues refers to the DD of
DDR1, DD of DDR2, C2 domains of factor V, VIII, and lactadherin.
2.2.3 Preparation of Phospholipid Vesicles (Liposomes)
Stock solutions of
L-α-phosphatidylcholine
(L-α-PC), and phosphatidylserine
(PLS) vesicles were prepared in 1:1 chloroform:methanol (v/v) and mixed in a L-α-PC:
PLS ratio of 1:1, or L-α-PC alone. Methylene chloride was added to the mixture and
evaporated twice under nitrogen gas. Phospholipids were then suspended by gently
swirling ice cold phosphate-buffered saline (137 mM NaCl, 10 mM Phosphate, 2.7 mM
KCl, pH 7.4) over the dried lipid suspension until all lipid was resuspended. Lipid
suspension was subjected to freeze-thawing in a dry ice-ethanol bath three times.
Vesicles prepared this way were used as multilamellar vesicles (95), and used within 24
hours of preparation. Resuspended vesicles were then sonicated in a high intensity bath
sonicator (Laboratory Supplies, Hicksville, NY) (96). L-α-PC vesicles consisted of 100%
L-α-PC
or with 10% NBD-L-α-PC where required for fluorescent studies, while PLS
vesicles had L-α-PC:PLS as 1:1 with or without 10% NBD-L-α-PC for labeled vesicles.
Final concentration of total lipid was not greater than 250 µm where otherwise indicated.
36
2.2.4
Cell Culture and Fluorescence Microscopy
THP-1 cells were grown in a 5% CO2 atmosphere at 37°C in RPMI 1640 medium
plus 2 mM L-glutamine (Life Technologies, Inc.), supplemented with 10% fetal bovine
serum, 50 units/ml penicillin, 100 µg/ml streptomycin, 10 mM HEPES, 1 mM sodium
pyruvate, 4.5 g/L glucose and 0.05 mM 2-mercaptoethonal and passaged 1:3 every 3-4
days. THP1 cells were transfected with either full length DDR1-myc or full length
DDR2-myc (4), and incubated for 24 hours at 37°C. Cells were then observed using
phase-contrast microscopy. After visualization, cells were washed and subjected to SDSPAGE to assess protein expression. Cell count did not exceed 0.5 X 106/mL.
CHO cells were grown under the same incubation conditions, except Dulbecco's
Modified Eagle Medium supplemented with 1.0 g/L D-glucose, 2 mM L-glutamine, 110
mg/L Na-Pyruvate, 0.5% penicillin/streptomycin, 0.1% L-proline, and 10% fetal bovine
serum was utilized. Cells were transiently transfected using expression plasmids for fulllength mouse DDR1 or DDR2 with a myc-tag at the intracellular carboxy terminus
(Regeneron Pharmaceuticals, NY) (4). Cells were then washed and incubated at 4°C for
120 min with labeled phospholipids vessicles. Cells were washed and fresh media cooled
to 4°C was added. Cells were observed with a Nikon fluorescent microscope at 400X
magnification and photographed. Cell count did not exceed 0.5 X106/mL. Cells were
subjected to SDS-PAGE after microscopy to assess protein expression.
37
2.2.5
Flow Cytometry Binding Assay with Fluorescent Phospholipid Vesicles
(Liposomes)
The vesicle binding assay was performed in 0.160 ml of buffer consisting of 10
mM HEPES-Na, pH 7.4, 133 mM NaCl, 5.8 mM KCl, 0.7 mm NaH2PO4, and 5 mM
glucose; NBD-labeled vesicles were added at 10 µM final concentration, and either 0 or
500 µM unlabeled vesicles were added to determine nonspecific binding (2). Transfected
(with either FL-DDR1-myc or FL-DDR2-myc) THP1 cells were resuspended in assay
buffer and added to tubes at a final concentration of 0.5 X 106 cells/ml. Non-transfected
cells were used as controls. Samples were incubated on ice for 120 min; cells were
pelleted at 500 X g, washed once in assay buffer, and resuspended in 0.4 ml of fresh
assay buffer containing 0.01 mg/ml propidium iodide to allow detection of necrotic cells.
Samples were analyzed with a fluorescence-activated cell sorter (FACS) Calibur
instrument, and data was processed with CellQuest Pro software (Becton Dickinson).
Forward and side scatter gates were set to include cells but exclude cellular debris and
phospholipid vesicles not bound to cells. NBD fluorescence was monitored in the FL1
channel and propidium iodide in the FL4 channel. The amount of fluorescent vesicles
bound per cell was then calculated from the mean fluorescence intensity of the gated cell
population. Specific binding was calculated by subtracting nonspecific binding (with
unlabeled vesicles) from total binding (with labeled vesicles only).
2.2.6
Surface Plasmon Resonance
To monitor the molecular interaction between the lipid vesicles and the proteins, a
BIACORE 3000 (Biacore AB, Sweden) equipped with a L1 chip was used. L-α-PC only,
38
and
L-α-PC
/ PLS complex vesicles were captured on the L1 chip surface at a
concentration of 1.5 mM total lipid. Before each vesicle injection, the chip surface was
cleaned three times by injecting 20 mM CHAPS solution (Sigma, St. Louis, MO, USA)
for 1 min at 10 µl/min. Vesicles were injected onto the cleaned chip for 30 min at a flow
rate of 3 µl/min, resulting in a signal of 500 response units (RU). The injected vesicle
concentration and volume were adjusted to a similar immobilization level for each type
of liposome. To remove unstably bound lipid vesicle from the surface, the chip surface
was treated twice with 50 mM NaOH for 1 min at 10 µl/min. To block the liposomeunpacked surface, the chip surface was treated with 0.1 mg/ml fatty-acid free BSA for 1
min at 10 µl/min (97). For vesicle–protein interaction analysis, 125 nM of DDR1 Fc and
DDR2 Fc (R&D Systems, Inc., Minneapolis, Mn), was diluted in a running buffer (10
mM HEPES, 150 mM NaCl, 5 mM CaCl2 at pH7.4) and injected for 1 min at 5 µl/min.
All analysis was performed at 4°C. The entire ECD of the mouse DDRs was fused to the
hinge, CH2 and CH3 regions of human IgG1 via a binding sequence GLY-PRO-GLY as
previously described (4) and is referred to as Fc.
2.2.7 DDR Phosphorylation Studies
CHO cells were transiently transfected with either full length DDR1-myc or full
length DDR2-myc (4). 24 hours post-transfection, serum free media was added to the
cells for a period of 18 hours. Cells were then stimulated with either 10 µg/mL
monomeric collagen type 1, with 20 µM L-α-PC / PLS vesicles, or with both collagen and
lipid vesicles, at 4°C for 90 min. Cells were lysed with cell Lysis buffer (Cell Signaling
Technology (Danvers, Ma.)) containing 20 mM Tris-HCl (pH 7.5), 150 mM NaCl, 2 mM
39
sodium orthovanadate, 1% Triton, 1 mM phenylmethylsulfonyl fluoride, 10 µg/ml
leupeptin, and 2 mM EDTA. Total cell lysates were then subjected to SDS-PAGE and
Western blotting analysis using 4G10 anti-phosphotyrosine antibody at 1 µg/mL to assess
phosphate binding. The choice of using 10 µg/mL collagen to stimulate the cells falls
within reported values and experiments (3,98). It is difficult to assess a µM concentration
of collagen since collagen type I is comprised of one α1(I) chain and one α2(I)
component (71) which are 139 kDa and 129 kDa in the monomeric form and slowly
converts from monomeric to other polymeric forms, thus greater a heterogeneous
mixture..
2.2.8
Vesicle Binding Assay
THP1 cells were transiently transfected with either full length DDR1-myc or full
length DDR2-myc (4). 24 hours post-transfection, cells were lysed with NuPAGE LDS®
sample buffer (Invitrogen) in the absence of ATP and Mg2+ and protein concentration
was determined. PLS or L-α-PC vesicles were added to a final concentration of 2.5
mg/mL, with a total protein concentration of 25 mg/mL of whole cell lysate. Lysates and
vesicles were allowed to incubate 24 hours at 4°C at constant motion. 25 µg of total
protein was removed and subjected to SDS-PAGE using anti-phosphotyrosine 4G10
antibody, followed by DDR antibody to confirm protein expression.
2.2.9
SDS PAGE and Western Blotting
SDS PAGE was performed using 4-12% NuPage Novex Bis-Tris Gels from
Invitrogen (Carlsbad, Ca). The proteins were diluted (final concentration 25-50 µg where
noted) in NuPage LDS® sample buffer (Invitrogen) containing 141 mM Tris base, 20%
40
LDS, 10% Glycerol, 0.51 mM EDTA, 0.22 mM SERVA Blue G250, and 0.175 mM
Phenol Red. Following SDS PAGE, the proteins were transferred onto nitrocellulose
membrane (Invitrogen) and blocked in 0.05% TBS-Tween buffer (20 mM Tris, 0.5 M
NaCl, ph 7.4-7.6, 0.05% Tween) with 5% whole milk. The membranes were then
incubated overnight in TBS-Tween with 0.05% milk in the presence of 1 µg/ml antiphosphotyrosine antibody at 4°C. The next day the membranes were incubated with antimouse IgG horseradish peroxidase (Santa Cruz, Ca) at 0.1 µg/mL and detection was
performed using enhanced chemiluminescence (Amersham Biosciences). For accurate
determination of the molecular weight, protein samples were electrophoresed against
BenchMark Ladder (Invitrogen).
2.3
RESULTS
2.3.1
Evaluation of amino acid Sequences between C2 domains and DD of DDRs
To determine if the amino acid sequences of the DD from DDRs are similar to
other known lipid binding regions in proteins (also called C2 domains), a protein
alignment was performed. Alignment of the DD of DDR1 and DDR2 from Homo sapiens
shows a 58.9% identity between these two proteins (Fig. 2.1). In order to assess the
structural sequence alignment of the DD from DDRs and those C2 domains of other
proteins which bind lipids, a multiple alignment was performed including the C2 domains
of blood coagulation factor V, VIII, and lactadherin. DD of DDR1 is between 25.3% and
30.6% identical to blood factors V, VIII, and lactadherin. The DD of DDR2 is 29.5% to
30.8% identical to those proteins as well. The identity of several conserved amino acids
41
that play a structural role in C2 domains (37) are preserved in the sequences of DDR1
and DDR2. These regions map to those areas which form the eight core β strands
(denoted as S regions in Fig. 2.1) from the known crystals structures of factor V, VIII,
and lactadherin (37-39). The greatest differences are seen outside the core and are
Figure 2.1. Amino acid alignment. ECD of DDR1 (CAI18451), DDR2 (CAI15941),
Blood Coagulation Factor V (NP_005919), Blood Coagulation Factor VIII (NP_000123),
Lactadherin (NP_005919) from Homo sapiens are shown. Loop regions representing
protein-ligand interaction are represented by bold arrows. Conserved residues are
highlighted in RED; partially conserved are in BLUE. Sequence stretches forming the βbarrel are designated S. Residues essential for lipid binding in factor V and VII are shown
with * and high lighted in yellow.
localized to the proposed lipid binding loops shown as green boxes in Fig 2.1. DDR1 is
20.0% to 30.0 % identical, and DDR2 23.5% to 30.0% identical to lipid binding protein
loops found in the C2 domains. In the case of factors V and VIII, it has been determined
42
that two sets of residues (highlighted in yellow) in the C2 domain are crucial for binding
of phospholipids-particularly PLS (99). A conserved hydrophobic tryptophan (W) residue
is found in factor V, VIII, and lactadherin, as well as DDR1 and DDR2 in this lipid
binding loop. The hydrophobic residue W is surrounded by hydrophilic residues. In the
second set of sequences implicated in binding phospholipids (99) which are also
highlighted in yellow, DDRs contain a glycine residue (G), as opposed to leucine (L).
2.3.2 Phase contrast and fluorescence microscopy used to determine binding of
lipids
To assess DDRs binding to phospholipids of the plasma membrane, human acute
monocytic leukemia cells (THP1) were transfected with full length mouse DDR1 and/or
DDR2 (4). THP1 cells were assessed via Western blot prior to experiment and found not
to endogenously express DDRs. Cells were visualized using phase-contrast microscopy
(X10) on plastic dishes. Fig 2.2a represents normal, untransfected THP1 cells. It is
important to note that cells were not grown past 0.5 X 106/mL confluency. Fig2.2b
represents those cells which have been transfected with full-length mouse DDR1-myc
(4), and Fig 2.2c represents THP1 cells transfected with full-length mouse DDR2-myc
(4). DDR2 transfected cells show a marked “clustering” or “clumping” of cells
43
Figure 2.2. Cell microscopy. THP1 Cells were transfected with full length
DDR1-myc (B), or DDR2-myc (C) (4). 24 hours post-transfection, cells were
visualized with phase contrast microscopy. A: untransfected cells. Cells
transfected with DDR2-myc show significant “clumping”. 10X magnification.
compared to untransfected controls. DDR1 transfected THP-1 cells also demonstrate
“clustering”, but is decreased compared to cells expressing DDR2. Following
microscopy, cells were washed and subjected to SDS-PAGE analysis and Western
blotting with either DDR1 and/or DDR2 antibody to assess DDR1/DDR2 protein
expression. Western blotting, using untransfected THP-1’s as a control, indicated a
significant increase in DDR1 and DDR2 protein in the transfected cells (data not shown).
To determine if cells expressing DDR1 or DDR2 bind phospholipids,
fluorescence microscopy was performed on Chinese hamster ovary (CHO) cells
transfected with full-length mouse DDR1 or DDR2 tagged with myc at the carboxy
terminus (4). Assays performed 24 hours after CHO transfection and at 4°C, confirmed
that cells expressing DDR1 bound NBD-labeled PLS. (Fig. 2.3). Cells expressing DDR2
44
showed little to no binding of NBD-labeled PLS. Untransfected cells were visualized as a
control.
2.3.3
Protein-lipid interactions analyzed by flow cytometry
To further investigate the possibility that phospholipid vesicles bind to DDRs, a
flow cytometric assay was developed to measure binding of vesicles to cells. To isolate
the step consisting of binding of vesicles to the cell surface prior to their possible
internalization by endocytic processes, assays were performed at 4°C. Previous work has
shown that binding of vesicles to cells occurs at 4°C, but internalization occurs only at
higher temperatures (90). To keep the labeled phospholipid component constant, both Lα-PC and L-α-PC:PLS vesicles were labeled with 10% NBD- L-α-PC; a final
Figure 2.3. Labeled phospholipids bound to cells. CHO (Chinese hamster ovary) cells
in solution. Cells were transfected with full length DDR1-myc, or DDR2-myc (4). 24
hours post-transfection, cells were incubated with NBD-L-α-PC/PLS for 120 min at
4°C. CHO represents untransfected cells; green represents fluorescent labeled PLS;
DDR1 and DDR2 are those cells transfected with the full length construct. Cell count
did not exceed 0.5 X 106/mL. 100X magnification.
45
concentration of 25 µm, with a total concentration of phospholipid of 250 µm,
comparable to reported values (2). Use of NBD- L-α-PC insured that THP-1 cells were
not labeled due to possible transmembrane transport of the labeled phospholipid by
aminophospholipid translocase. THP-1 cells were transfected with either full-length
DDR1 or DDR2. Negative controls were untransfected THP1 cells. As mentioned
previously, earlier experiments determined that THP1 cells do not endogenously express
DDR1 and/or DDR2. Fig. 2.4a shows the histogram of PLS binding (left), and L-α-PC
binding (right). Gate E on both histograms indicates the regions analyzed to determine
the mean fluorescence of the cell population. The amount of fluorescent vesicles
bound/cell was then calculated from the mean fluorescence intensity of the gated cell
population.
46
Figure 2.4. Flow Cytometry. A: Typical flow cytometry histograms. The assay was
performed in the presence of L-α-PC:PLS (left), and L-α-PC (right) with 10% NBD-Lα-PC. Total lipid concentration was 250 µM (2). Gate indicates the region analyzed to
determine the mean fluorescence of the cell population. B: Mean fluorescence
calculated with increasing addition of phospholipid. THP1 cells transfected with full
length mouse DDR1 (red), DDR2, (blue), and untransfected cells (black). Left
represents L-α-PC:PLS; right side represents L-α-PC. Experiments design, performed,
and analyzed by Dr. Gunjan Agarwal and Dr. Michelle Jon Nauerth.
47
Specific binding was calculated by subtracting nonspecific binding from total binding.
The lower half of Figure 2.4 (Fig. 2.4b), shows an increase in mean fluorescence
with an increase in phospholipid concentrations. PLS binding to cells (left side) shows
PLS having an initial increase in mean fluorescence in those cells which express full
length DDR1 (red line). Cells expressing DDR2 (blue line) also show an increase in
fluorescence with the addition of increasing PLS concentration. Black line represents
untransfected THP1 cells. The right hand side represents L-α-PC vesicles added to THP1
cells. Cells expressing DDR2 show a high affinity for L-α-PC as shown with an increase
in mean fluorescence (blue line). DDR1 expressing cells also show an affinity for L-αPC, but at a much lower fluorescence compared to DDR2 cells (red line). Untransfected
THP1 cells (black line) show little to no affinity for phospholipids..
2.3.4
Surface Plasmon Resonance (SPR) evaluation of DDRs demonstrates
protein-ligand preference
SPR provides a powerful tool for protein-ligand interactions. This is true for low
affinity interactions which are difficult to determine using other methods. SPR also offers
a quick method for evaluation of the functional integrity of recombinant proteins. In this
study, we evaluate the data on a simple qualitative basis only. Fig. 2.5 shows the
relationship between the ligand, PLS, as it associates with the ECD of recombinant DDR1
and DDR2 proteins. Based upon RUs, DDR1 demonstrates an affinity for PLS, as shown
by a maximal RU at the junction of association/dissociation (downward arrow). It is
assumed that this is a simple 1:1 interaction between purified ECD DDR1 and PLS.
48
Response Units (RUs) are significantly lower at this same junction for binding of PLS to
DDR2, TrkB (control protein), and monomeric collagen.
Figure 2.5.
Surface Plasmon Resonance. The phospholipid PLS was
immobilized to a sensor chip and was used to capture purified ECD of DDR1
(red), DDR2 (green), monomeric collagen (brown), and TrkB (blue). The upward
arrow shows the point at which purified proteins were injected. The downward
arrow shows the point at which dissociation started. All experiments were
performed at 4°C. Each sensorgram has been normalized to zero at the minimum
absorbance for each trace to facilitate comparison between the different proteins.
Experiment designed, performed, and analyzed by Dr. Cosmin Mihai
49
2.3.5
Full length DDR1 becomes phosphorylated with PLS
Based on the structure of the DDRs, where a ligand binding extracellular domain
(ECD) has been identified, a transmembrane spanning region, and a cytoplasmic receptor
tyrosine kinase (RTK), it is assumed that if phospholipid vesicles did bind the ECD, that
the internal RTK would become phosphorylated. An experiment was designed in which
Chinese hamster ovary (CHO) cells were transfected with full length mouse DDR1 (4).
24 hours post-transfection, serum free media was added to cells for 12 hours. Cells were
incubated with 10 µg/mL monomeric collagen type I (-/+-, +/+-), 20 µM PLS vesicles (-/+, +/-+), or both (-/++, +/++) for 120 min at 4°C. SDS-PAGE was performed in
conjunction with Western blotting using anti-phosphotyrosine 4G10 antibody. Western
blot analysis (Fig. 2.6) showed no phosphorylation in lanes -/--, -/-+, -/+-, or -/++. These
samples indicate they were not transfected with DDR1, therefore the receptor tyrosine
kinase does not
50
Figure 2.6. SDS-PAGE and Western blotting. 4G10 anti-phosphotyrosine antibody
was used. Cells were stimulated with PLS vesicles, monomeric collagen, or both
vesicles and collagen. As controls, non-transfected and non-stimulated cells (-/--),
and cells transfected with full length DDR1, stimulated with monomeric collagen
(+/+-) were used (3).
Legend: -/--, untransfected, unstimulated; -/-+, untransfected, stimulated with PLS;
+/-+, transfected, stimulated with PLS; -/++, untransfected, stimulated with collagen
and PLS; +/+-, transfected, stimulated with collagen; -/+-, untransfected, stimulated
with collagen; +/--, transfected only; +/++, transfected, stimulated with collagen and
PLS.
exist and phosphorylation should not take place. In previous experiments (data not
shown), it was confirmed that CHO cells do not contain endogenous DDR1, which
confirms no phosphorylation in -/-- lanes. Phosphorylation does occur in lanes +/++, +/+, and +/-+. These lanes do contain the over expressed full length DDR1, which contains
the tyrosine kinase in the cytoplasmic domain. What is interesting to note, +/-- constructs
also show phosphorylation. However, this is a result previously seen in this same type of
experiment-in which DDR1 autophosphorylates (4,22).
51
2.3.6
Vesicle binding assay in whole cell lysates
An additional experiment was designed involving whole cell lysates of those cells
exposed to phospholipids compared to those assays designed with adhered cells. THP1
cells, a non-differentiated cell line, were transfected with mouse full length DDR1, or not
transfected and used as a control. 24 hours after transfection, whole cell lysates were
prepared from each sample and incubated overnight at 4°C with constant motion in a
1:10 ratio of lipid vesicles to protein in NuPage LDS® buffer (Invitrogen), in the absence
of ATP and Mg2+. SDS-PAGE was then performed and phosphorylation was assessed via
Western blotting. Lane 1 (Fig. 2.7) is DDR1 control protein. This control was a protein
sample from a previous experiment. In that experiment, cells were stimulated with
monomeric collagen and a positive result for phosphorylation occurred to confirm
antibody detects phosphorylation. Lysate from untransfected cells (THP1 PS) only
showed no phosphorylation. Whole cell lysate from THP1 cells transected with DDR1
and incubated with PLS or L-α-PC, however, demonstrated that phosphorylation took
place. Previous experiments not presented here indicate L-α-PC may not bind to DDR1.
The cell lysate from cells transfected with DDR2 show no phosphorylation after
incubation with PLS vesicles. Presence of L-α-PC did not increase phosphorylation of
DDR1 above control.
52
Figure 2.7. SDS-PAGE and Western blotting with anti-phosphotyrosine
4G10. Cells were transfected with full length mouse DDR1, with the
exception of untransfected THP1 cells. Lysates were incubated overnight
with L-α-PC or L-α-PC:PLS vesicles. 25 µg of protein was added to each
lane. DDR1 control protein, which was stimulated with monomeric
collagen in a previous experiment, was used as positive control. Lanes
were labeled 1.0 to indicate first vesicle:protein concentration ratio used.
2.4
DISCUSSION
Collagens are abundant components of the extracellular matrix, particularly in
atherosclerotic plaques. Collagens can influence the behavior of smooth muscle cells
(SMCs) and macrophages during plaque development. Adhesion to collagen has also
been shown to promote macrophage differentiation and phagocytosis-the initial step
being exposure of PLS to the outer membrane of the cell (100). The effects of collagen on
the biology of SMCs and macrophages emphasizes the importance of collagen signaling
during atherogenesis and are accomplished through the action of specific collagen
receptors such as the discoidin domain receptors. It remains to be seen if there are other
ligands of the DDRs, particularly phospholipids, and what role, if any, the DDRs play in
53
binding to phospholipids and the contribution to atherosclerotic plaque formation and/or
blood coagulation.
In this study, the binding of DDR1 and DDR2 to phospholipid vesicles has been
investigated on the premise of a qualitative evaluation will be employed. The hypothesis
that DDRs may bind to phospholipids is based on the homology between the discoidintype C2 domains of lactadherin, blood coagulation factor V, and blood coagulation factor
VIII. Of these three, the blood coagulation factors are involved in just that-blood
coagulation. However, lactadherin has been reported to be secreted by macrophages and
is involved in the anchoring of phagocytic cells via PLS, to macrophage integrins (101) –
playing a dual role of binding a phospholipid and a protein. At this stage, it is not clear if
phospholipids are a ligand for the DDRs, and what role, if any, they play in phagocytosis,
and/or blood coagulation. The amino acid sequence of the DDs of the DDRs
demonstrates a high probability that DDRs do bind to phsopholipids. Since the threedimensional structures for lactadherin (16), blood coagulation VIII (38), and blood
coagulation V (37) have been solved, it is easy to compare those amino acid sequences
with those of the DDR DD regions (Fig. 2.1). Just from sequence alignment alone, there
is a strong resemblance to the core region of the phospholipid binding structures.
However, the loop regions (green loops, Fig. 2.1) of the DDs contain key hydrophobic
residues that have previously been implicated in phospholipid binding (39,102,103).
These regions are highly variable and may be a key to phospholipid binding in the DDRs.
The use of THP1 (Human acute monocytic leukemia) cells was to demonstrate a
specific ligand binding pattern of the DDRs. Since THP1’s are a non-differentiated cell
54
line (in the manner they were used in this study), the effect of “clustering” in transfected
cells can be possibly associated with phospholipid binding. THP1 cells transfected with
full length DDR2-myc demonstrated significant clustering compared to DDR1-myc
transfected and non-transfected cells. This “clustering” effect may be due to the DDR2
ECD binding to the L-α-PC in the outer portion of the plasma membrane. It is known that
THP1 cells do express Fc and C3b receptors on the cell outer membrane (104), however,
it has not been demonstrated that the DDR ECDs bind to these receptors. It is interesting
to note that only the DDR2-myc transfected cells demonstrate “clustering”, which was
not observed in the DDR1-myc transfected cells to such a degree. There is a 59% identity
between the two discoidin ECDs, if the phenomenon were attributed to protein ECDECD interaction, the clustering may be observed as well in the DDR1 transfected cells. It
has also not been determined that the DDR proteins do form protein-protein interactions
via their ECDs. It can be concluded there is a likelihood that THP1 cells expressing
DDR2 interact via DDR2: L-α-PC involvement.
Fluorescent-labeled L-α-PC incorporated into a 1:1 mix of L-α-PC: PLS displayed
increased fluorescence in Chinese hamster ovary cells (CHO) which over expressed
DDR1-myc. This response is similar to the membrane binding properties of lactadherin,
blood factor V and blood factor VIII (93,105). In the study presented here, a 50% content
of PLS was employed in the vesicles. In the case of factor V and lactadherin, those
proteins recognize membranes with as little as 4% PLS content (93), but blood factor VIII
does not. Since there is no attachment of the labeled vesicles in those cells which express
DDR2, it can be assumed that binding of DDR1 occurs via PLS interaction in those cells
55
expressing DDR1. This observation, in conjunction with the data acquired by the use of
SPR, also confirms that PLS may be a preferred ligand of DDR1. With that particular
experiment, the purified ECD of DDR1 and DDR2 was passed over immobilized PLS
vesicles. The results of the SPR data are merely qualitative; however, the binding of the
DDR1 ECD to PLS vesicles in the SPR experiment is greater than DDR2 and controls.
This same result is mimicked in the C2 domain of blood coagulation factor V (106).
Blood factor V binds to negatively charged phospholipid membranes and serves as a
cofactor for the activation of prothrombin in the prothombinase complex. This complex
contains the enzyme factor Xa, the protein cofactor blood coagulation factor V, calcium
ions, and a phospholipid membrane surface provided by activated platelets (35,107).
Formation of the prothrombinase complex requires a procoagulant membrane surface.
Activated platelets, platelet microparticles, and other damaged vascular cells provide this
surface (108). It has been reported that factor V binds to phospholipid vesicles containing
25% PLS with a Kd of approximately 10-9 nM (108-110). Further experiments need to be
employed in which the Kd can be determined for phospholipid vesicle binding in DDR1
and DDR2.
To determine if phospholipids are a true ligand of the DDRs, a phospholipid
binding study was employed. This experiment is very similar to those experiments in
which collagen was employed as a ligand (3,4,33). In those experiments, transfected
cells-which do not express DDR proteins endogenously, were transfected with full-length
DDR1 or DDR2, and then stimulated with 10 µg/mL monomeric collagen type I. 1 mM
sodium orthovanadate was also used as a positive control for phosphorylation (3,4,33).
56
The use of 20 µM PLS was used either alone, or in the presence of 10 µg/mL collagen
type I for the experiments in this study. For the most part, those cells which were
transfected with full length DDR1 and exposed to either PLS and/or collagen
demonstrated with a certain validity that phosphorylation takes place with phospholipids
as a ligand. However, those cells which were transfected with DDR1, but not exposed to
either PLS or collagen, also showed phosphorylation. This has been previously shown in
cell stimulation experiments (4,111). This possibly could be an artifact of the transfection
process itself. FuGENE 6.0 (Roche Applied Science, Indianapolis, IN) was applied as a
transfection agent. This agent is somewhat toxic to the cells. Even though the cells were
washed several times after the transfection procedure to eliminate any dead cells, there
possibly could be several remaining. Apoptotic cells tend to expose PLS to the outer
membrane, thus making an apoptotic cell a ligand for DDR1. The results of this
experiment were confirmed with the results of the vesicle binding assay. A positive result
for phosphorylation occurred when whole cell lysates were incubated with PLS. However,
a positive result also resulted when whole cell lysates from cells transfected with DDR2
were incubated with PLS. There is a possibility that DDR2 has a tendency to bind L-α-PC
with the preliminary data which has been presented here. However, the experiments
employed in this study lacked the analytical degree to determine a clear conclusion.
To that end, it is possible that phospholipids are a ligand for the DDRs, and that
the DDRs are lipid specific. However, in this study, the analytical techniques found in the
literature were not employed, and each result was simply qualitative. It is highly
suggestive that phospholipids do bind to the DDRs, but, to further solidify this
57
conclusion, such techniques as a solid-phase ELISA, isothermal titration calorimetry,
and/or surface Plasmon resonance with different phospholipid types and concentrations
would need to be employed.
58
CHAPTER 3
STRUCTURAL PREDICTION OF DISCOIDIN DOMAIN RECEPTORS (DDR1
AND DDR2): A POTENTIAL LIPID BINDING PROTEIN
3. 1
INTRODUCTION
Background
Collagen is a major component of the extracellular matrices. Five different types
of collagen receptors have been identified: integrins, glycoprotein VI, leukocyteassociated IG-like receptor-1, the mannose receptor family, and the discoidin domain
receptor tyrosine kinases-DDR1 and DDR2 (28,112). Discoidin domains (DDs) are found
in a number of functionally unrelated proteins, including blood coagulation factor V and
VIII, lactadherin, and neuropilin-1. Discoidin domains have evolved from a common
ancestor to bind very diverse ligands (collagen, lipids, carbohydrates, or growth factors).
It has been demonstrated, based on structural studies, that the diverse ligand recognition
among the discoidin domain family is mediated by the highly variable loops clustered on
the surface or end region of the discoidin domain (47). The nature and the lengths of
these loops define protein specificity and are responsible for its biological function. It is
known that collagen is a ligand for the DDRs (59). Previously, the region encompassing
59
the putative ligand binding loops in DDR1 and DDR2, L1-L4, have been replaced by
corresponding loops of other discoidin domains (3). The loop chimeras of L1-L4 in both
DDR1 and DDR2 lost the ability to recognize collagen, suggesting that these loops are
responsible for collagen binding.
Blood coagulation factor V is the non-enzymatic cofactor for the prothrombinase
complex-which is required for normal blood hemostasis. The complex itself consists of
blood factors V and X, and a phospholipid membrane from activated platelets
(35,109,113). The complex generates thrombin. Factor V is homologous to factor VIII,
the nonenzymatic cofactor of the enzymatic complex that activates factor X. This
complex also requires a phospholipid membrane for activity (114). Phospholipid binding
occurs through the C2 domain in both factor V and VIII (115,116). With the elucidation
of the crystal structures of factor V and VIII (37,38), phospholipid binding of the C2
domains in these proteins becomes clarified.
The structures of blood factor V, VIII, and lactadherin (39)-another phospholipid
binding protein, have been resolved with the same overall structure. The common feature
is a β-barrel core with three relatively long loops protruding from one end. The loops
form a hydrophobic surface that overlies a ring of basic amino acids. Both C2 domains
from factor V and VIII have 3-4 water exposed hydrophobic residues protruding from the
tips of the long loops, leading to the hypothesis that membrane binding is mediated by
insertion of these residues into the membrane. The basic residues may interact with the
polar head groups of the phospholipids (117). Functionality of these hydrophobic
residues has been confirmed by site-directed mutagenesis (94,99). The crystal structures
60
have also provided a basis for the theory that the amino acids interact with the
hydrophilic head group of PLS. Sequence homology, as well as conformational flexibility
of the protein itself, have been found in factors V and VIII, and also lactadherin (39).
Lactadherin has also adopted the central β-barrel motif and the relatively functional long
loops which display membrane-interactive amino acids.
Hypothesis
Discoidin domain proteins (DDR1 and DDR2) show an amino acid sequence
similarity to the C2 domain proteins. Therefore, my hypothesis is DDR1 and DDR2
should also show a structural homology with those proteins which have solved threedimensional crystal structures: blood factor V and VIII, and lactadherin. To test this
hypothesis, we propose to obtain the homology three-dimensional model of DDR1 and
DDR2 DD using the solved structures of the C2 domain proteins. The validity of the
structure will be assessed using molecular dynamic simulation, and should yield atomic
level information about the solution structure of the DD of DDR1 and DDR2. With the
proposed models, hypothetical conclusions can be drawn as to the function of the DDR
binding regions, and what ligands may bind to the DD in these proteins.
3.2
METHODS
3.2.1 Sequence Alignment
Sequences of the extracellular domain (ECD) of DDR1 (CAI18451), DDR2
(CAI15941),
Factor
V
(NP_005919),
Factor
61
VIII
(NP_000123),
Lactadherin
(NP_005919), and Neuropilin-1 (NP_003864.4) from Homo sapiens were aligned using
the CLUSTALW algorithm (Blosum62 scoring matrix) in Bioedit. The alignment was
edited using the GeneDoc multiple alignment editor. The tilde character (~) was inserted
to manually edit the alignment. Loop regions relevant for protein-lipid interactions are
represented by blue boxes. Conserved residues are highlighted in RED; partially
conserved are in BLUE. Sequence stretches forming the β-barrel are designated as yellow
arrows. Extra strands represented with β-barrel region are in purple. Residues implicated
in lipid binding to factor V are indicated by * and highlighted in YELLOW. Amino acid
residues refers to the DD of DDR1, DD of DDR2, C2 domains of factor V, VIII,
lactadherin, and the FA58C domain of neuropilin-1. Numbering was not included on the
alignment.
3.2.2
Model Construction
Three–dimensional structures of the Discoidin Domains (DD) currently available
were obtained from the Protein Data Bank (PDB) (118). The specific X-ray structures
selected for the DD were those with no ligand attached. When different depositions were
available, the structure with the highest resolution (as observed as lowest Å of resolution)
was selected for use as the template. The PDB identification numbers of the selected DD
structures used in the analysis are presented in Table 1.
The x-ray crystal coordinates of blood coagulation factor V (Protein Data Bank
entry: 1CZT), along with the crystallographic water molecules were used to model the
DD of DDR1. The DD of DDR2 was modeled after the crystal coordinates of human
neuropilin-1 (Protein Data Bank entry: 1KEX). The structures of the DD were
62
constructed by replacing residues of the homologue x-ray crystal structures with the
corresponding DDR residues from the amino acid alignment using SYBYL 6.4 (Tripos,
inc., St. Louis, MO). Inserted loops of the models were roughly accommodated by using
a loop conformational search program (SYBYL/Biopolymer). Major steric interferences
caused by side chains were removed automatically and manually after visual inspection.
3.2.3
Simulation Setup
The molecular model of the DD of DDR1 together with the structural waters
derived from the x-ray crystal structure of blood factor V was subjected to molecular
dynamic minimization. The same occurred with the model of the DD of DDR2, but
utilized the x-ray crystal structure data of human neuropilin-1. Full minimization on the
side chains of the entire protein was performed while fixing the backbone to relieve bad
contacts. Minimization of the entire protein, including the backbone, was performed for
1000 steps. The protein together with the crystal waters was then placed in a box of water
molecules with the box boundaries at least 12.0 Å from any given protein atom. Water
molecules with oxygen atoms closer than 2.0 Å to any protein atom were excluded.
The simulations were performed using the SANDER module of AMBER 9.0
force-field suite of programs (UCSF, CA, USA.). The PROCHECK program was used to
asses the sterochemical quality of the models (119). In the first step, only the added water
molecules were energy minimized at constant volume (10,000 conjugate gradient steps),
then all of the water molecules were subjected to energy minimization (another 10,000
conjugate gradient steps), and finally, the whole system was energy minimized (10,000
conjugate gradient steps). The system was subsequently subjected to a slow heat-up
63
procedure to bring the temperature of the system to 300K. After 25 ps of a constant
volume/constant temperature simulation, the system was reminimized (20,000 conjugate
gradient steps). After another heat-up run of 10 ps to bring the temperature back to 300K,
a constant temperature/constant volume MD run was performed for 25 ps. Finally, a
constant temperature protocol was adopted to simulate 600 ps of dynamics.
The final predicted free solution structure was checked using the program
PROCHECK (120) to evaluate the “goodness” of the geometrical entities such as bonds,
angles, and dihedral angles in the average molecular structure of the solution protein
when compared with those found in the crystal.
3.3
RESULTS
3.3.1
Choice of the Template Structures and Alignment
To select the best template structures for modeling the DD of DDR1 and DDR2,
the crystal structures of discoidin domains available at the time were considered (Table
1). These structures all show a conserved β-sandwich core consisting of 8 strands with 6
relative long loops protruding from one end. Additionally, there are 7 β fold regions in
the surrounding loops which are not part of the sandwich fold. Following the
nomenclature established for blood factor VIII (38), and human neuropilin-1 b1 domain
(45), the sandwich core is designated in green with the numbering system 1-8 in the final
models.
Six juxtaposed loop regions (L1-L6) extend from the top of the β sandwich core.
In the orientation represented here, these loops are at the bottom of the protein structure.
64
Table 3.1. The PDB identification numbers. Selected discoidin structures were
used in the model analysis.
These loops define the limits of a groove or cleft that runs between L1-L3 and L2-L4.
The greatest differences are seen outside of the core region and localized to the loops. It
is apparent that the L2 and L4 loops are shorter than L1 and L3, and it is this region that
is implicated in phospholipid binding. However, these loops are highly variable. On this
basis, the choice for the modeling template is crucial for determining a hypothesized
structure of DDR1 and DDR2. The overall fold characteristics are consistent with
previously published structures. Nevertheless, the length and fold of the connecting loops
and the resulting relative positions of the helices with respect to the different folds of the
loop, as well as the presence and the location of the binding pocket/region, could be
modeled in significantly different ways depending on the structure selected as the
template.
The two most critical issues in homology modeling are the degree of similarity
between the target sequence and the templates and the reliability of the alignment, two
65
aspects that are intrinsically interconnected (121). By applying a recursive PSI-BLAST
search of the DDR1 and DDR2 DD region against the PDB database, the only sequence
producing a significant alignment in the first cycle against DDR1 was that of blood
coagulation V (1CZT) (E-value = 9.95144E-10). Other statistically significant
homologies with the DD of DDR1 are blood coagulation factor VIII (1D7P) (E-value =
3.84406E-10), and human neuropilin-1 (1KEX) (E-value = 2.18089E-8). Human blood
coagulation V appeared to be the more optimal reference sequence/structure for initial
DDR1 alignment. PSI-BLAST search of DD of DDR2 produced a significant alignment
with human neuropilin-1 (1KEX) (E-value = 1.28428E-19), lactadherin (3BN6) (E-value
= 3.64644E-17), and blood coagulation V (1CZT) (E-value = 1.0176E-16). In this case,
human neuropilin-1 was the more optimal choice for modeling the DD of DDR2. The
preferred ligand for neuropilin-1 is vascular endothelial growth factor (VEGF). It has not
been established if neuropilin-1 binds to phospholipids. Neuropilin-1 is not involved in
the blood coagulation cascade. Structurally, this is a better match for DDR2, and
currently there is no hypothesized need for neuropilin-1 to bind to a phospholipid.
3.3.2
Evaluation of amino acid Sequences between C2 domains and DD of DDRs
The known DD sequences were aligned using CLUSTALW. Figure 3.1 represents
the alignment. The pairwise sequence identities and similarities between the DD target
sequences and the DD domains are presented in Table 2. The highest sequence identity
and similarity of the DD of DDR1 was that of human blood coagulation VIII with 28%
66
Fig. 3.1. Alignment of DDR1 and DDR2 DD. Includes discoidin sequence from blood
coagulation factor V, blood coagulation factor VIII, lactadherin, and neuropilin-1. Loop
regions important for protein-ligand interaction are boxed in green. Conserved residues
are highlighted in red; partially conserved residues in blue. Sequence stretches forming
the β-barrel are in yellow; extra β strands are represented in pink. Accession numbers
are: DDR1, CAI18533 residues 31-185; DDR2, CAI15941 residues 30-184; Factor V,
AAB59401 residues 2066-2221; Factor VIII NP_000123 residues 2193-2345;
lactadherin, NP_005919 residues 230-387; neuropilin-1, NP_003864.4 residues 274424. Numbering in this alignment begins at 1. Numbering of sequences where indicated
for protein.
67
Table 3.2. The % identity and similarity between the DD region of DDR1 and all
other proteins evaluated in the study.
Table 3.3. The % identity and similarity between the DD region of DDR2 and all
other proteins evaluated in the study.
identity and 48% similarity. DD of DDR2 had the highest sequence identity with that of
human neuropilin-1 with 32% identity and 47% similarity. However, all sequences
analyzed showed approximately 25%-30% identity with 50% similarity. The data are in
agreement with the knowledge about functional similarities across discoidin domains and
suggest that any member of this family may be a suitable template for modeling.
68
3.3.3
Global aspects of the simulation
Molecular dynamics simulation methods have proved to be a valuable tool for
obtaining molecular level information of protein structures in solution. It is important to
establish that the simulation system is equilibrated prior to the structure being analyzed.
Evaluating RMSDs has been demonstrated to be an appropriate procedure for monitoring
equilibration of the simulated structure (122). Based on this, RMSDs were calculated for
the factor VIII, factor V, and neuropilin-1 crystal structures. In addition, the DD of DDR1
was modeled against factor V and neuropilin-1. Those RMSDs are represented along
with the RMSDs of the DD of DDR2 modeled against neuropilin-1. RMSDs are an
assessment of the constancy in the potential energy measure of the stability of the system.
Simulators use RMSDs with respect to the crystal configuration or to the initial
minimized configuration (123,124). In Fig. 3.2, the RMSDs of the DD region of DDR1
and DDR2 are displayed. Smaller fluctuations in all values of RMSD during the 2000 –
3000 ps segment of the simulation ensure the stability of the system in solution. The
displacements of the backbone atoms of factor V, factor VIII, and neuropilin-1 with
respect to their x-ray crystal positions remain relatively small, with RMSD fluctuations in
the 0.8 – 1.2 Å range. The DD of DDR1 with respect to the factor V crystal structure
shows a slightly higher magnitude of approximately 1.5 – 1.8Å. The DD of DDR2 in
respect to neuropilin-1 and DD of DDR1 with respect to neuropilin-1 x-ray structure
demonstrate large fluctuations in their RMSD values. Such deviations may arise because
of relative motions in the loop region of the protein itself.
69
Fig. 3.2. Root-mean square deviations (RMSDs). Back bone atoms of DDR1
referenced to blood factor V (black); back bone atoms of DDR1 referenced to
neuropilin-1 (red); back bone atoms of DDR2 referenced to neuropilin-1 (green).
Deviations of reference crystal structures; blood factor V (yellow), blood factor VIII
(blue), and neuropilin-1 (purple).
70
The simulation B-factors calculated for the backbone atoms by averaging over the
3000 – 5000 ps segment of the trajectory are given in Fig. 3.3. The B -factor is an
experimental measure of the thermal harmonic motion of the atoms. The larger the value,
the less localized the atom. The (pseudo) B-factors for simulation can be computed from
the fluctuations of atoms about their mean positions between cycles of the molecular
dynamic simulations. Several peaks in the B-factor versus time plots are observed. The
residues in the regions of the peaks are located in the variable loops, and are not a part of
the overall β-barrel fold found in the DD of DDR1 and DDR2. The majority of the
residues that move are found in loop 3 (Fig. 3.5). This loop is one of the longer loops, and
is also implicated in phospholipid binding. The amino acid alignment in figure 3.1
outlines those residues involved in loop 3. It is important to note that in previous studies
with factor V and VIII, mutation of those residues to alanine reduces phospholipid
binding (4,115,125). However, in those same studies, Loop 3 residues are not implicated
in direct binding to a phospholipid, and may play an indirect role in phospholipid
binding.
Figure 3.4 depicts a profile of Cα distance deviations by optimally superimposing
the DD of DDR1 over blood factor V, and DD of DDR2 over neuropilin-1. The Cα
distance was obtained to evaluate the changes in the backbone conformation upon
solvation. In general, the deviations are similar to the B-factor profiles. The greatest
deviations occur in the same region as loop 3 of the model. What is interesting is the
greater deviation occurs when the solvent structures are modeled after neuropilin-1.
Within this loop, most residues are conserved; however, the residues implicated in
71
phospholipid binding here are lysine in neuropilin-1, as opposed to glycine which is
found in DDR1 and DDR2. Based on the size of the residue, this may have an impact on
the movement of this loop.
Fig. 3.3. B-factors calculated for the back bone Cα –atoms of the solvated models.
The simulation B-factors of the backbone atoms were calculated using the x-ray
crystal structure as a reference configuration over the last 500 ps of the simulation.
Highest deviation is noted for Loop 3 in the solvate model.
72
Fig. 3.4. Deviations of the α-carbon atoms of the solvated structures and the x-ray
crystal structures. Greatest deviations occur in the Loop 3 region of the model.
3.3.4
The models
Figure 3.5a represents the backbone alignment of the DD of DDR1 against factor
V. DDR1 is represented as the red ribbon, factor V as green. The core β-barrel sandwich
73
Fig. 3.5. Conformational differences between the modeled DDR1 (red) (left slide)
against blood coagulation factor V (green). Significant tryptophan which has been
implicated in phospholipid binding is shown in both DDR1 and factor V. DDR2 (red)
mapped against neuropilin-1 (green) is shown on the left. Both images show
movement in loop 3.
is conserved. The majority of the variability occurs within the loop regions. Loop 3
(opposite loop 1 which designates important tryptophan residues essential in
phospholipid binding), protrudes in the opposite direction of the crystal structure of factor
V. This would account for the variability seen in the B–factor and Cα distance data. It is
important to realize that loop 1 is truncated in DDR1, as opposed to the same loop in
74
factor V. This would also account for some variability, but not as significant as loop 3.
Fig. 3.5b represents the backbone alignment of DDR2 DD (green) against neuropilin-1
(red). The alignment is much more refined in terms of the loop regions. Loop 3
demonstrates minimal movement, as well as loop 1. Loop 1 is truncated in neuropilin-1
as well as DDR2. In both the DDR1 and DDR2 overlays, significant tryptophan residues
are noted within Loop1. These residues have been implicated in phospholipid binding
(4,115) in factor V.
75
Fig. 3.6. Images of solvated DD of DDR1 taken at several time points during the
simulation. Loop 3 (opposite tryptophan 53) shows variable movement. Tryptophan 53
shows movement in vertical position, with little movement of Loop 1. Right-hand side
figure also demonstrates fluctuations in Loop 3, with horizontal fluctuations of tryptophan
53.
Models were captured at different times during the simulation (in 0.5 ps). Fig. 3.6
represents two views of the modeled DDR1 DD protein. Loop 1 is interesting in that the
loops itself seem to show little to no movement. The essential tryptophan (Trp53) moves
in not just a vertical arrangement, but also in a horizontal form as well. Since the loop is
truncated, this movement of the amino acid may be essential for interaction with the
phospholipid. Loop 3 demonstrates large fluctuation in movement. It is interesting to note
76
that Fig. 3.7 represents the DDR2 DD model imaged at different time points. Loop 1
shows variable movement, however, Trp52 shows little to no displacement. It is assumed
that phospholipid interaction occurs via movement of the loops in DDR2, and not the
movement of the tryptophan itself as compared to DDR1 and as observed in Fig. 3.7.
Loop 3 also shows significant movement in the other region.
77
Fig. 3.7. Solvated structure of DDR2 taken at multiple
time points during minimization. Tryptophan 52 moves in
horizontal position, mainly because Loop 1 shifts
horizontally.
78
Figure 3.8 represents the solved solution structure of the DD of DDR1. The
overall conserved β-sandwich is represented in yellow. This region is conserved and
characteristic of the discoidin family, originating with bacterial galactose oxidase (34).
Figure 3.9 represents the DD of DDR2 structure. Both structures are composed largely of
β strands, β turns, and random coils. The loops connecting the strands are of varying
length and contribute to the variable element of the structure at either end of the barrel,
producing a relatively flat upper surface and an irregular lower surface. The two-three
relatively large loops on the lower surface resemble the corresponding loops of blood
factor V, factor VIII, and lactadherin. These loops carry proven or hypothetical
membrane-interactive hydrophobic amino acids (37).
79
Fig. 3.8. Completed molecular model of the DD of DDR1. Core β-structure is
designated in yellow; additional β-structures in purple. Loop regions are designated in
green. Loop 1 shows essential tryptophan, variable Loop 3 is opposite. Model is in
orientation in which the loops implicated in plasma membrane binding are pointed
down. At the top of the figure are the loops that face the cell wall and connects to the
trans-membrane spanning region.
80
Fig. 3.9. Completed molecular model of the DD of DDR2. Core β-structure is
designated in yellow; additional β-structures in purple. Loop regions are designated in
green. Loop 1 shows essential tryptophan, variable Loop 3 is opposite. Model is in
orientation in which the loops implicated in plasma membrane binding are pointed
down. At the top of the figure are the loops that face the cell wall and connects to the
trans-membrane spanning region.
81
3.4
DISCUSSION
In summary, the modeled structure of the DDs of DDR1 and DDR2 reveal a
predicted discoidin domain fold present in many multidomain extracellular and
membrane proteins involved in cell regulation events. The structures modeled here
represent the most detailed structure of the DDRs to date, and employ analytical methods
to confirm the structure authenticity and reliability. Two other structures have previously
been reported. The Vogel (33), group utilized SWISS-MODEL software to overlay the
DD of DDR1 over the three-dimensional structures of factor V (PDB: 1CZT), factor VIII
(PDB: 1D7P), and neuropilin-1 b1 domain (1KEX). This minimized model was
employed for the rational design of mutations/deletions in an attempt to determine the
collagen type I binding site in the DD region of DDR1. Shimada, et. al (36) determined
the solution structure of the DD of DDR2 to elucidate the collagen recognition
mechanism of the DD region by utilizing multi-dimensional NMR techniques. Both
models reported the same overall fold: eight major β-strands which form the sandwich or
jellyroll barrel common to other DD domains. The bottom of the β-barrel is closed with
interconnecting straight segments, which is seen in all the DD domain crystal structures,
and in the model that is reported here (Figs. 3.8 and 3.9) of the DD of DDR1 and DDR2
(in the notation of the model formed in this study, these loops are shown at the top). At
the top of the β-barrel core protrudes the 6 juxtaposed loops (L1-L6) in which L1 and L3
are the longer, which is also characteristic of what is seen in our model-with the
exception that our model is presented in an opposite orientation. In Figs. 3.8 and 3.9, the
loops are at the bottom of the figure.
82
The structures of the DDs of DDR1 and DDR2 resolved here provide a
foundation and understanding of the general principles involved in DDR molecular
recognition of ligands other than collagens. Appreciation of the three-dimensional
structure allows us to visualize and reconcile how this single domain can function in
specific binding interactions with structurally diverse ligands. The resolved structures
illuminate general regions of the molecule involved in possible binding of phospholipids
as a ligand. In the case of the DD of DDR1, Loop 1 is truncated and forms a flat and/or
scoop surface. On this loop contains an essential tryptophan residue previously identified
in blood factor V and VIII, and lactadherin (36,94,99,117) which is essential for
membrane binding-particularly to phosphatidylserine. It may be likely that some or all of
the residues in loop 1 are solvent exposed, and those interact with the phospholipid head
group. If the essential tryptophan is not solvent exposed, it may insert itself into the
membrane and interact with the plasma membrane hydrophobic tail.
It has not been established among the DDRs if phospholipid is a known ligand.
However, Loop 3 (Figs. 3.1, 3.8, and 3.9), not only shows a high level of similarity
amongst its residues, but a high level of functionality. A possible conclusion or
hypothesis involves the role of Loop 3. In blood coagulation factor VIII, mutation of the
residues in Loop3 leads to a decrease in phospholipid binding (117). Two critical residues
for phospholipid binding in this loop (found in blood factor VIII) are the leucines. These
amino acids have no reactive groups on their side chains and therefore do not react
favorably with water. They react much more favorably with other nonpolar atoms. The
working hypothesis for these residues involves insertion of the loop into the plasma
83
membrane where these residues can interact with the phospholipid tail groups. Both
DDR1 and DDR2 contain glycines in this same position as the leucines are found in
factor VIII. These glycines allow for greater flexibility of the polypeptide backbone.
Possibly the role of this loop is to insert itself into the plasma membrane, or have the
flexibility to interact with the polar head groups of the plasma membrane phospholipids.
Loop 3 may play a role in binding only-as in to serve as an anchor on to the plasma
membrane surface, where Loop 1 plays a role in eventual activation of the downstream
kinase in a full length discoidin protein upon phopspholipid binding.
Regions of the protein predicted to be involved in phospholipid interaction on the
basis of the discoidin domain structures can only be validated upon obtaining structures
of these domains in complex with their binding partners. In this study, we were not able
to complete this task. However, future studies with these models in the presence of a
phospholipid membrane may address the following questions: 1) Are the hydrophobic
residues on the loop regions of the protein buried in the hydrophobic regions of the outer
membrane below the polar head groups; 2) Are the positively charged residues of the
loops located in the polar region of the phosphatidyl head groups of the membrane, or
just above the polar zone; and 3) The DD of DDR1 and DDR2 is orientated perpendicular
to the plane of the plasma membrane. Are the conformations of the charged and nonpolar side chains optimized for binding to a phospholipid membrane? Do some residues
interact with charge groups of the membrane, even if their location is above the plane of
the membrane polar head groups?
84
CHAPTER 4
DRR1 AND DDR2 AFFECT PLATELET-COLLAGEN INTERACTION
4. 1
INTRODUCTION
Background
Blood coagulation is a complex process in which blood forms a thrombus. It is an
important part of hemostasis-in which a damaged blood vessel wall is covered by a
platelet and fibrin coating-preventing further bleeding. Coagulation involves a cellular
process (smooth muscle cell response, platelet activation), and a protein (collagen type I,
coagulation factor) component. Coagulation begins almost instantly after damage to the
vessel wall (endothelium).
Smooth muscle cells (SMCs) are activated after vessel injury, and respond
through proliferation, migration, and extracellular matrix (ECM) synthesis. Smooth
muscle cells produce by far most of the extracellular matrix components (126). Basically,
injury occurs to the vessel at the endothelial surface. Removal of the endothelium occurs,
exposing the basement membranes of the vessel. SMCs migrate from the media of the
vessel wall, either to the plaque which is forming, or to the endothelium. Once exposure
85
of SMCs occurs, fibrillar type I collagen synthesis takes place (127,128) by the SMCs.
Collagens represent up to 40% of the total protein of the vessel wall, in which collagen
type I is the majority of the species present. Not only do collagens provide integrity, but
fibrillar collagen also provides a surface for attachment of platelets. Normally, fibrillar
collagens are located in the ECM underlying vascular endothelial cells and are not
exposed to flowing blood. After injury when exposure of subendothelial structures
occurs, blood will flow directly over the ECM and the exposed collagen. One of the first
events in the thrombotic process is the adherence of platelets to the modified ECM.
Platelets adhere more readily to exposed fibrillar collagen after vascular injury. In
addition, an increase in shear rate will cause more pronounced platelet adhesion. Blood
flow will be slowed in response to plaque build-up, even though there is an increase in
pressure. Once the plaque has ruptured and blood flow increases, pressure decreases.
Platelet adhesion is enhanced by increased presence of red blood cells. In flowing blood,
red cells occupy a position in the middle or central portion of the vessel and push
platelets toward the periphery, increasing platelet-vessel wall contact.
Fibrillar type I collagen is the only ECM protein which supports both platelet
adhesion and platelet activation. When collagen becomes exposed to flowing blood,
platelets rapidly adhere, spread, and become activated to form an aggregate. Damage to
blood vessel walls also exposes subendothelium proteins, most notably von Willebrand
factor (vWF), present under the endothelium, and in coordination with the fibrillar
collagen layer. vWF is a protein secreted by the healthy endothelium, forming a layer
between the endothelium and underlying basement membrane. When the endothelium is
86
damaged, the normally-isolated, underlying vWF is exposed to blood and recruits factor
VIII, collagen, and other clotting factors. Circulating platelets bind to collagen with
surface collagen-specific glycoprotein Ia/IIa receptors. This adhesion is strengthened
further by additional circulating vWF , which forms additional links between the platelets
glycoprotein Ib/IX/V receptor and the collagen fibrils. These adhesions activate the
platelets.
The principal mechanism used to stop bleeding consists of a pair of overlapping
proteolytic cascades called the extrinsic and intrinsic coagulation pathways (129-131).
Fig. 4.1 shows how both pathways merge and lead to the formation of fibrin. In the
intrinsic pathway, all necessary factors are present in the circulating blood, including a
potentially negatively charged surface, such as that of activated platelets. In the extrinsic
activation pathway, it is not a plasma component, but tissue fluid that initiates the blood
coagulation process: after vessel damage, tissue factor, also called tissue thromboplastin,
mixes with the blood and starts the coagulation process. In the location of where these
two pathways merge, blood coagulation factors VIII and V are activated by serine
proteases to interact with blood factor X-which leads to the formation of thrombin, and
eventually activates fibrinogen into fibrin-the fibrillar protein which enhances the platelet
plug. Activated platelets are members of the intrinsic pathway which lead to prothrombin.
Blood coagulation factors V and VIII bind to the activated platelets. When platelets
become activated, the inner region of the plasma membrane flips to expose PLS. Fibrillar
collagen indirectly plays a crucial role in regulating thrombin formation because
negatively charged phospholipids, such as PLS, become exposed on the surface of
87
platelets after an indirect interaction with collagen and form the catalytic surface for the
assembly of active coagulation complexes and thrombin generation (132). Binding of the
coagulation factors found in the blood coagulation pathway to biological membranes is a
pivotal step in the
Fig 4.1. The blood coagulation pathway. Identification of intrinsic and
extrinsic pathways.
88
process of coagulation. The activation of most of these proteins takes place on the
membrane surface where the binding localizes the enzyme-substrate complex. The
phospholipid is required for the prothrombinase complex to function.
It has been well established that both blood coagulation factors V and VIII bind to
PLS on activated platelets. A large body of literature exists on “how” this binding occurs.
However, the milk fat globular protein, lactadherin, also binds to PLS membranes, but
currently has not been linked to the coagulation pathway. Lactadherin functions as a
potent anticoagulant (57) and as a reagent for detection of PLS exposure early in
apoptosis of leukemia cells (49). Homology between the lactadherin C domains and those
of factor V and VIII correlates with the capacity of lactadherin to compete efficiently for
membrane binding sites on PLS containing membranes. The capacity for effective
competition is explained, in part, by steroselective binding of PLS (30). Lactadherin
inhibits the factor Xase complex, in which factor VIII functions, and the prothrombinase
complex, in which factor V functions. Factor V and factor VIII do not compete efficiently
with each other for membrane binding sites (133,134), whereas lactadherin displaces both
with half-maximal displacement at 1-4 nM (57). Lactadherin also competes for
membrane binding sites of vitamin K-dependent coagulation proteins, inhibiting the
factor VIIa-tissure factor complex. The coagulation inhibitory properties and PLS
detecting properties of lactadherin contrast with those of other PLS binding proteins. The
capacity of lactadherin to interact with a range of PLS containing binding sites and to
compete efficiently with coagulation proteins is an unusual feature.
89
Hypothesis
The hypothesis of this study is that the discoidin domain proteins 1 and 2 will
interfere with platelet aggregation. It has been established that the discoidin domain
receptor proteins function as receptors for fibrillar collagen (4,21). However, nothing is
known of the DDRs role or function in the vascular system-particularly in regards to
blood coagulation. We deduce that DDRs will prevent platelet aggregation occurring by
interfering with the formation of fibrillar collagen, or bind to collagen in such a way as to
prevent aggregation from occurring. We will test this theory by using platelet
aggregometry as a technique and observe changes in platelet aggregation when using
collagens with DDRs as an agonist. Other cell techniques will be employed to establish
that platelets do not contain DDRs themselves. We surmise that the DDRs will delay
platelet activation in the presence of collagen.
4.2
MATERIALS AND METHODS
4.2.1 Reagents/Materials
Adenosine diphosphate was purchased by Sigma Chemicals (St. Louis, MO).
Bovine dermal collagen type I, PureCol®, was purchased from Advanced BioMatrix Inc.
(San Diego, CA). Blood draw supplies were purchased from VWR International
(Philadelphia, PA). Platelet aggregometer and software was from Chrono-Log Corp.
(Havertown, Pa). Cuvettes for aggregometry and stir bars specific for 500 µL tubes were
also from Chrono-Log Corp. Purified extracellular domains of DDR1 and DDR2
(recombinant human/Fc chimera, CF), as well as recombinant human TrkB/FC chimera,
90
proteins were obtained from R&D Systems (Minneapolis, MN). Anti-DDR1 and antiDDR2 antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, Ca).
4.2.2 Platelet Draw
Healthy subjects, 18 – 40 years of age, volunteered for whole blood
donation. Written informed consents were obtained from each donor in accordance with
an approved protocol from the institutional review board for human subjects from The
Ohio State University. Phlebotomy was performed in the morning between 7 and 10 AM,
in a fasting state, without caffeine intake 18 hours prior. In addition, each subject
abstained from medication, specifically non-steroidal anti-inflammatory drugs, 10 days
prior to donation. Venous blood samples were collected into 3.8% sodium citrate (9:1,
v/v) tubes. The first tube of drawn whole blood aspirated was discarded. 20 mL’s of
whole blood was then centrifuged at room temperature for 15 min at 200 X g to obtain
platelet rich plasma (PRP) (135,136). Platelets were transferred to polypropylene tubes
and stored at room temperature. A small portion of the PRP was further centrifuged at
room temperature at 1200 X g and plasma was removed from the platelet plug. This
sample was designated as platelet poor plasma (PPP), and would be used as a reference
sample in aggregometry. A standardized platelet concentration of 3.75X108mL-1 for a
500 µl reaction was prepared by dilution of PRP with PPP after a platelet count was
determined with a Z1 Coulter counter (Coulter Corp., Miami, Fl) (137).
91
4.2.3
SDS PAGE and Western Blotting
Activated and inactivated platelets of PRP were pelleted by centrifugation at 500
X g at room temperature for 7 min and subsequently washed two times with Tryode
solution (138) (138 mmol/L NaCl, 2.9 mmol/L KCl, 12 mmol/L NaHCO2, 0.36 mmol/L
Na2HPO4, 5.5 mmol/L glucose, 1.8 mmol/L CaCl2, 0.49 mmol/L MgCl2, pH 7.4).
Platelets were lysed in 20 mM Tris-HCl (pH 7.5), 150 mM NaCl, 2 mM sodium
orthovanadate, 1% Triton, 1 mM phenylmethylsulfonyl fluoride, 10 µg/ml aprotinin, 10
µg/ml leupeptin, and 2 mM EDTA. Protein concentration was assessed and a range of
concentrations were prepared in 1 X NuPage LDS sample buffer (Invitrogen, Carlsbad,
Ca). Electrophoresis was performed using 4 – 12% (w/v) NuPAGE Novex Bis-Tris Gels
from Invitrogen. As a positive control, whole cell lysate (WCL) from HEK293 cells, over
expressing either DDR1 or DDR2 protein, was used as a marker. SDS PAGE was
followed by Western blotting using nitrocellulose membranes (Invitrogen). The
membranes were probed with either anti-DDR1 (sc-532) or anti-DDR2 antibodies and
imaged using enhanced chemiluminescence from Amersham Biosciences (Piscataway,
NJ) after incubation with HRP-conjugated anti-rabbit IgG.
4.2.4
Collagen Preparation
Bovine dermal collagen type 1 (PureCol, Advance BioMatrix, San Diego, Ca.)
was used at a final concentration of 0.5 to 10 μg/mL in PBS. Monomeric (M) collagen
was used by diluting collagen in ice-cold PBS immediately before use. Fibrillar (F)
collagen was obtained by incubating M collagen at 37°C for 4-12 hours, followed by
separating and washing the pellet by centrifugation steps. The F-collagen pellet was
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resuspended in PPP and added as an agonist. Semi-polymeric form of collagen (SP) was
the supernatant collected during the formation of F collagen (139,140). Purified
extracellular domains of DDR1 and DDR2 (recombinant human/Fc chimera, CF), as well
as recombinant human TrkB/FC chimera, CF proteins were obtained from R&D Systems
(Minneapolis, MN). DDRs were mixed with collagens in a 1:5 mass ratio. Unless
otherwise noted, DDRs (or TrkB) and collagen were combined, allowed to incubate at
37°C for 1-6 hrs and the sample was added as an agonist for platelet aggregation.
4.2.5
Platelet Aggregation
A 4-channel platelet aggregometer (Chrono-Log Corp., Havertown, Pa.), was pre-
warmed for 45 min at 37°C. 500 µL siliconized glass tubes specific for aggregometry
were also pre-warmed to this temperature. PPP samples were used as a reference. A stir
speed of 1000 RPM was employed for each sample. PRP and PPP was added to each
cuvette with constant stirring and pre-warmed to 37°C prior to addition of agonist. The
aggregation assay was standard and corresponded to the method of Born (141). In this
assay, 100% aggregation corresponds to the light transmittance signal obtained with a
sample of PPP. Platelet viability was determined for each donation by the use of 10 µM
ADP-a known agonist for platelet aggregation. Any platelet donation less than 60%
aggregation with ADP was eliminated from use. Platelet aggregometry occurred for 30
min unless noted.
4.2.6
Statistical Analysis
A students t-test was performed to determine differences in time to initiation of
aggregation in the presence of collagen only, and collagen with either DDR1 or DDR2. A
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range of collagen concentrations from 0.5 µg/mL – 10 µg/mL was utilized. DDRs were
added in a 5:1 ratio. Statistics were performed on n=5. A two-way analysis of variance
(ANOVA) with repeated measures was also performed between collagen concentrations
and time to aggregation in the presence of DDRs.
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4.3
RESULTS
4.3.1 Platelets do not express DDRs
Platelets were isolated by the standard protocol of Born (141). After they were
washed in Tyrodes’s buffer, cells were subjected to SDS-PAGE analysis to determine if
unactive/active platelets express DDR1 and/or DDR2. Fig. 4.2 represents a western blot
of platelet lysates. A) Western blot probed with anti-DDR1 and anti-DDR2. Lane 1
represents control lysate from cells over-expressing DDR1 and DDR2. This lysate was
utilized to determine viability of the antibody. Lane 2 is a sample of PPP (platelet poor
plasma). This was to insure that the plasma portion of whole blood contains no detectable
amount of the DDR proteins by the use of Western blotting. Lane 3-5 represent different
concentrations of whole cell lysate. Even at a protein concentration of 25 µg/mL, there is
no detectable amount of DDR proteins in the whole cell lysates of unactivated platelets.
B) Western blot of activated platelets probed with anti-DDR1. Lane 1 contains the whole
cell lysate sample from cells over-expressing DDR1. Lane 2-4 represent whole cell
lysates from platelets which were activated by different concentrations of fibrillar
collagen type I. 25 µg/mL of protein was loaded onto the gel. Even though platelets were
activated at concentrations of fibrillar collagen less that 2.0 µg/mL, a range of fibrillar
collagen up to 10 µg/mL shows no detectable expression of DDR1. Lane 5 is a sample of
whole cell lysates from platelets activated with 10 µM ADP. Again, no detectable amount
of DDR1 exists. C) Platelets activated with 5 µg/mL of collagen in the presence of the
ECD of DDR1 (5:1 ratio). DDR1 should delay the normal (142,143) fibrillogenesis and
disrupt the native banded structure of collagen type I. That being the case, collagen in the
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presence of DDR1 should show no activation of the platelets. There is no detectable
DDR1 in the whole cell lysates of this experiment. D) Platelets in the presence of
monomeric collagen. 5 µg/mL of monomeric collagen was used as an agonist for
platelets. Not only did the platelets not aggregate, but there is no expression of DDR1.
The same is true of the negative control which is whole cell lysate from non-activated
platelets.
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4.3.2
Determining platelet aggregation by morphological forms of collagen and the
DDR proteins alone
Fig. 4.3a represents a platelet aggregation experiment. Morphological forms of
collagen were created by incubating collagen type I overnight at 37°C. During this
incubation, fibrillar collagen is formed, while 50% of the sample forms into an
intermediate state (data not shown here) as determined by hydroxyproline assay (144).
Platelets contain known fibrillar collagen receptors-glycoprotein Ia/IIa (also known as
integrin α2β1), and GPIV. The results (A), indicate that the presence of fibrillar collagen
Fig 4.3. A) Platelet aggregation by morphological forms of collagen type I. F form is
equivalent to native fibers which is a known platelet agonist. M form is monomeric.
SP is monomeric to fibrillar intermediate. B) Platelets were subjected to DDR1 ECD,
DDR2 ECD, or TrkB as agonist. Fibrillar (F) collagen was control.
97
is
associated
with platelet aggregation, but monomeric
and
semi-polymeric
(morphological collagen intermediate) do not influence aggregation of platelets. In these
experiments, an indication of platelet aggregation is any measure > 20%. The
morphological forms of collagen were created and used in this experiment to confirm the
preferred agonist for the platelets is the fibrillar form. These results are not novel, and
used here as a basis for further experiments. In order to assess whether or not the ECD of
DDR1 or DDR2 induces platelet aggregation, a concentration of 10 µg/ml of each protein
(DDR1, DDR2 and TrkB) was determined, and added as an agonist to the platelets alone.
As a positive control, 10 µg/ml of fibrillar collagen was used as agonist. Fig. 4.3b show
the DDRs did not induce changes in the platelets as assessed by platelet aggregation.
Therefore, DDRs do not directly activate platelets and inhibit aggregation in the presence
of fibrillar collagen.
It is known that the formation of collagen fibrils is disrupted by the ECD of the
DDRs (142,143). To determine if platelet aggregation is disrupted by this form of
collagen, platelets were subjected to 10 µg/ml of collagen, incubated 5:1 in the presence
of DDR1, DDR2, or TrkB only, overnight at 37°c. Fig. 4.4 represents the aggregation
tracings of these samples. DDR1 and DDR2 incubated with collagen was not an agonist
for platelets. However, fibrillar collagen again showed platelet aggregation, as well as
TrkB, a negative control protein that does not bind to collagen, or disrupt collagen fibril
formation.
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Fig. 4.4. Platelet aggregation in response to fibrillar collagen and fibrillar
collagen in the presence of the ECD of DDR1 and DDR2. 10.0 µg/ml of
collagen was incubated overnight at 37°C in the presence of 1:5 ratio of the ECD
of DDR1` and/or DDR2. As a positive control, collagen only was used, and a
negative control was collagen incubated with TrkB. DDR1 and DDR2
eliminated platelet aggregation.
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4.3.3 DDR1 and DDR2 delay time to aggregation in platelets
To determine any response in time to aggregation, a lag time experiment was
designed. Lag time was calculated as the difference between the time the agonist was
added to the platelet sample, and the increase in aggregation as per the aggregation
tracing. Fig. 4.5 and 4.6 depicts this lag time. DDRs were added to collagen in a 1:5 ratio,
incubated at 37°C for 60 min, and used in aggregation as the agonist. In the case of
DDR1, lag time was increased in those collagen samples that were incubated with the
DDR1 protein. Lag time occurs in as little as 1.0 µg/ml collagen when collagen is
incubated with DDR1. However, as the DDR1:collagen concentration increased, lag time
decreased. A students t-test performed on this experiment by comparing lag time of
collagen incubated with DDR1 and collagen alone, was significant in those
concentrations greater than 2.0 µg/mL. Collagen levels less than 2.0 µg/mL with DDR1
either failed to aggregate platelets (< 20% aggregation), or were not significant. Fig 4.6 is
the aggregation tracing of DDR2 incubated with collagen. This experiment also
demonstrates that DDR2 in the presence of collagen slows the initiation of platelet
aggregation. Aggregation tracings of DDR2 were of 2.0-4.0 µg/mL, as lower values
showed variable results in lag time and % aggregation. It is important to remember that in
this particular experiment, collagen samples were incubated for 60 min in the presence of
the DDRs. The results of this experiment show that during that time, collagen
fibrillogenesis does not move to completion. It is inhibited by the DDRs, but enough
collagen remains to move towards the fibrillar state and induce aggregation. What is
100
interesting to note is the lag times of DDR2 increase with increasing concentrations of
collagen.
Fig. 4.7 is a depiction of the lag times over the course of collagen concentration
increase. Collagen alone, and collagen + TrkB, decrease lag time as the concentration
increases. This is an indication that the collagen is moving to the fibrillar state and is a
true agonist of the platelets. As the concentration of collagen + DDR1 increases, the lag
time does decrease to some extent. DDR1 does disrupt the formation of collagen fibrils,
but in this case collagen fibril formation is moving to completion since aggregation is
occurring. The opposite affect occurs with collagen combined with DDR2. As the
concentration of collagen:DDR2 increases, lag time increases-the opposite effect of
collagen:DDR1. There is a 58% amino acid identity between DDR1 and DDR2.
However, it is clear that two different mechanisms of collagen binding are occurring,
influencing collagen fibrillogenesis, and interfering with platelet aggregation.
Fig. 4.7 shows aggregation occurs for no more than 250 seconds. Percent error is
indicated by black bars, and was calculated via means and standard deviations. A twoway analysis of variation (ANOVA) was performed with replication. P-values were
<0.005, indicating differences in time to aggregation occurred in all concentrations of
collagen:protein (5:1 ratio). In addition, differences also occurred statistically (p<0.005)
amongst collagen in the presence of the DDRs. There is also a statistically significant
interaction that is detected between collagen:protein concentration and type of protein
added (collagen only; collagen + TrkB; collagen + DDR1; collagen + DDR2).
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Fig 4.5. Platelets were exposed to collagen alone, and collagen incubated with the
ECD of DDR1. Time to activation, also known as lag time, increases with increasing
concentration of DDR1 with collagen. Percent aggregations seems to be unaffected.
102
Fig 4.6 Platelets were exposed to collagen alone, and collagen incubated with the
ECD of DDR2. Time to activation, also known as lag time, increases with increasing
concentration of DDR2 with collagen. Percent aggregations seems to be unaffected.
103
Fig 4.7. Comparison of lag times with increasing collagen. Either DDR1, DDR2, or
TrkB were incubated 1:5 with collagen, allowed to incubate overnight, and used as
agonist for platelet aggregation. Lag time decreases with increased collagen
concentration. TrkB is a control protein for the DDRs. However, lag time for
DDR2 increases with concentration.
4.3.4
Platelets do not aggregate when DDRs are added to fibrillar collagen
To resolve any interaction of DDRs with fibrillar collagen, collagen type I was
incubated overnight at 37°C to form the fibrillar state. Once purified and washed to
remove any trapped collagen morphological species, DDRs were added and allowed to
incubate at room temperature for 90 min at room temp. The entire sample was then added
to the platelets. Fig. 4.8 is the aggregation tracing of this experiment. Platelets do not
104
aggregate when DDR1 and DDR2 are combined with fibrillar collagen. In a similar
experiment (Fig. 4.9), collagen again was allowed to form to its fibrillar state overnight.
Fig. 4.8. Collagen was incubated overnight at 37°C. DDR1, DDR2, or TrkB was
added for 90 min, and then added as an agonist. DDR1 and DDR2 inhibited platelets
from aggregating.
105
Fig. 4.9. Collagen was incubated over night. DDR1, DDR2, or TrkB was added to
the collagen and incubated for 90 min. Sample was washed and added to platelets as
an agonist. Samples containing discoidins did not aggregate platelets.
106
This sample was washed several times in PBS, and DDR1, DDR2, or TrkB was
incubated for 90 min at room temp with the collagen. After incubation, the sample was
washed several times to remove excess DDRs/TrkB, and any other collagen species.
Sample was added to platelets as an agonist. Aggregation did not occur in those samples
where DDR1 or DDR2 was combined with fibrillar collagen, as aggregation is
represented as < 20%. TrkB did not interact with the collagen, therefore aggregation took
place. In addition, collagen alone, in the fibrillar state, allowed for platelet aggregation to
take place.
4.4
DISCUSSION
The results of this study are novel and this is the first report involving DDRs in
the role of platelet aggregation. The results of this study support the hypothesis that
DDRs disrupt platelet aggregation/activation by their interaction with collagens.
Specifically, how this is achieved can be based on three conclusions.
1) The discoidin domains are known to bind collagens in their ECD (3,21). It is
also known that the DDRs interact with collagen by disrupting the formation
of the fibrillar form and disrupt the overall banded structure of collagen
(142,143). Platelet aggregation is completely inhibited by the DDR/collagen
structures, as platelets prefer fibrillar collagen to interact with integrin α2β1
and GPVI receptors for activation.
2) Platelet aggregation is delayed by DDR1 and DDR2, most notably by
interfering with the process of fibrillogenesis after the addition to collagen.
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Fibrillogenesis has been monitored by collagen turbidity experiments
(142,143), in which the addition of DDRs delay collagen from forming fibrils,
as monitored visually at 37°C.
3) Even though DDR1 and DDR2 share > 58% residue identity, and both
demonstrate a lag time for platelet aggregation, there may be two mechanisms
at play since lag time for DDR1 is opposite that of DDR2.
4) Platelet aggregation is inhibited by the ECD of DDR1 and DDR2. This is
observed when the DDRs are added to fibrillar collagen. The DDRs bind,
prevent collagen from interacting with the platelets, and the platelets do not
aggregate. Even after washing the collagen:DDR ECD sample, enough of the
DDR is bound to the collagen to prevent aggregation from occurring.
The process of platelet aggregation is an encompassing task in regards to vascular
injury. It is difficult to asses exactly where and what role the DDRs may play in this
process. Currently, no research has been reported in regards to DDRs efforts in collageninduced platelet aggregation. The findings that are reported here are novel and shed some
light as to a possible collagen-platelet interaction. However, larger questions remain.
Since it has been determine by Western blotting, that platelets do not contain DDRs, and
in addition, blood plasma does not contain DDRs as well (when detected by Western
blotting), where do the DDRs exist and originate from? Hou et. al (22), reported that
DDR1 mRNA and protein was increased after injury to the rat carotid artery. It is known
that after vascular injury, smooth muscle cell (SMC) proliferation peaks in approximately
1 or 2 days (145), and the first SMCs begin migrating from the media to the intima of the
108
vessel (146). DDR1 protein expression was expressed in concurrence with this migration
(22). Therefore, there may be an increase in DDRs post vessel trauma as a result of
injury. But what role do the DDRs play in response to trauma? Based on the data reported
here, DDRs may bind to collagen and prevent thrombus formation. This may be a
physiological response for the prevention of vessel occlusion by the thrombus.
Since platelet aggregation is completely impaired when DDRs are combined with
fibrillar collagen, do the DDRs bind to collagen in a location (on collagen) that is
recognized by the platelet collagen receptors- integrin α2β1 and GPVI? Or do DDRS bind
to these receptors without activation of the platelet occurring? To asses this theory,
washed platelets would need to be incubated with the ECDs of DDRs, and then exposed
to fibrillar collagen. Aggregation would be evaluated to ascertain a block for fibrillar
collagen. The lack of aggregation can then be attributed to the DDRs. In our study, we
combined collagen:DDRs in a 5:1 ratio. The stoichiometry of collagen:DDR molecule
relationship has yet to be established. This is a critical molecular relationship as a
saturation of DDR on the collagen may falsely be preventing aggregation to occur and
this response may not mimic what is happening in vivo.
Wound repair and artherosclerosis involves many components and pathways. It is
difficult to speculate what role, if any, the DDRs may play in this process. In the case of
lactadherin, it is believed that this protein interferes with thrombus formation by
competing with the blood coagulation factors for phospholipid binding sites. In addition,
lactadherin can also bind to integrin on macrophages, as mentioned previously. Is there a
similar role for the DDRs? It may be prudent to investigate one small feature at a time109
beginning with plaque formation, DDR expression in apoptotic SMCs, the role of DDRs
in the ECM, etc. A global understanding of the role and DDR-signaling pathways can
then be achieved.
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CHAPTER 5
DISCOIDIN DOMAIN RECEPTOR 1 RECOGNIZES A MORPHOLOGICAL
STATE OF COLLAGEN TYPE I
5. 1
INTRODUCTION
Background
A large variety of organs and tissues have within their structure a complex
network of collagen fibers which govern mechanical properties (68). Collagen must
withstand elastic extension particularly in the vasculature, and in such processes as
wound healing (147,148). Collagen type I in its mature fibrillar state is the major
component of the extracellular matrix (ECM). Collagen fibers interact with cells through
cell surface receptors and soluble proteins, which is integral to cell proliferation, survival,
attachment and cellular differentiation. Collagen is formed from three polypeptide αhelical strands which form a unique triple-helical structure known as monomeric (M)
form. Collagen type I self-assembles into D-periodic cross-striated fibrils (149) where D
= 67 nm, the characteristic axial periodicity of collagen. Each M collagen triple helix is
approximately 300 nm in length and 1.5 nm in diameter. The M form of collagen type I is
driven entropically into microfibrillar structures 4 nm in diameter into a semi-polymeric
111
form (SP), which further condenses into larger fibrils (F) (74,150,151). These fibrils
assemble into bundles to form fibers as well as a large variety of macroassemblies found
in connective tissues. When M collagen moves to the F morphological form, the amount
of fibrillar F collagen formed can be determined by use of the hydroxyproline assay
(144). The amount of collagen that incorporates into F fibers at the saturation
equilibration level is approximately 50%. The other 50% are incomplete formed semipolymeric fibers, SP, and can be identified by atomic force microscopy (AFM) (143).
The assembly of collagen fibers (fibrillogenesis) M to SP to F, is a complex
process regulated by a variety of collagen-binding proteins and other molecules that may
directly or indirectly interact with the collagen molecules and fibrils. The fibrillar states
of collagen define the mechanical properties of the ECM and govern cell-matrix
interactions. This assembly is modulated by the discoidin domain receptor protein 1
(DDR1) (142,143). The interaction of DDR1 with collagen(s), and their functional
implications are not well understood.
Hypothesis
In this study, we investigated if collagen-induced tyrosine phosphorylation of
DDR1 dependent on a specific fibrillar morphology of collagen type 1. Three different
morphological forms of collagen type 1: monomeric (M), semi-polymeric (SP) and
fibrillar (F) collagen were utilized in our cell-based assays. Since DDR1 is a
transmembrane spanning protein containing an intercellular receptor tyrosine kinase,
binding of collagens at the extracellular domain (ECD) will induce phosphorylation,
112
which can be detected by Western blotting. We hypothesize that all three morphological
forms of collagen type I can modulate DDR1 phosphorylation. Therefore it is speculated
that collagen in ECM remodeling is a highly conserved and efficient process.
5.2
MATERIALS AND METHODS
5.2.1 Reagents/Materials
Bovine dermal collagen type I, PureCol®, was purchased from Advanced
BioMatrix Inc. (San Diego, CA). This collagen is 95-98% type I collagen with the
remainder being type III collagen. It is primarily in the monomeric form at a
concentration of approximately 3.2 mg/ml and contains < 1% oligomer. DDR1-MYC
plasmids encoding the entire mouse DDR1 tagged with triple MYC-tag was from
Regeneron Pharmaceuticals (4). Phosphate-buffered saline (PBS) (pH 7.2) was from
VWR (Bridgeport, NJ). Affinity purified rabbit polyclonal anti-DDR1 antibody (sc-532)
raised against the c-terminus of human DDR1 was from Santa Cruz Biotechnology (Santa
Cruz, Ca). Human embryonic kidney 293 cells (ATCC, Manassas, VA) were cultured in
Dulbecco’s modified Eagle medium/F12 nutrient mixture (invitrogen) with 10% (v/v)
fetal bovine serum. Anti-phosphotyrosine (4G10) clone was from Upstate, Temecula,
CA. DDR1 siRNA was purchased from Santa Cruz Biotechnology (Santa Cruz, CA).
5.2.2
Collagen Preparation
Bovine dermal collagen type 1 was used at a final concentration of 0.5 to 12.5
μg/mL in PBS. Monomeric (M) collagen was used by diluting collagen in ice-cold PBS
113
immediately before use. Fibrillar (F) collagen was obtained by incubating M collagen at
37°C for 4-12 hours, followed by separating and washing the pellet by centrifugation
steps. Semi-polymeric form of collagen (SP) was the supernatant collected during the
formation of F collagen (139,140).
5.2.3
Cell Culture
Hek293 cells were transiently transfected with full-length DDR1 plasmid for 24
hours. Cells were then serum starved for 12 hour and stimulated with collagen at either
4°C, or 37°C for 90 min or where specified. Cells were then washed with PBS, lysed, and
total protein concentration was determined in the whole cell lysate (WCL).
DDR1 siRNA was added to confluent Hek293 cells for 8 hours. Cells were then
washed, and serum free media was added for a period of 12 hours. M, SP, or F collagen
type I was added at a concentration of 0.5 μg/mL for 90 min at 4°C. Cells were washed
and whole cell lysate was extracted and assessed for phosphorylation by Western
blotting.
5.2.4
SDS PAGE and Western Blotting
SDS PAGE was performed using 4-12% NuPage Novex Bis-Tris Gels from
Invitrogen (Carlsbad, Ca). The proteins were diluted (final concentration 25-50 µg where
noted) in NuPage LDS® sample buffer (Invitrogen) containing 141 mM Tris base, 20%
LDS, 10% Glycerol, 0.51 mM EDTA, 0.22 mM SERVA Blue G250, and 0.175 mM
Phenol Red. Following SDS PAGE, the proteins were transferred onto nitrocellulose
114
membrane (Invitrogen) and blocked in 0.05% TBS-Tween buffer (20 mM Tris, 0.5 M
Nacl, ph 7.4-7.6, 0.05% Tween) with 5% milk. The membranes were then incubated
overnight in TBS-Tween with 0.5% milk in the presence of 1 µg/ml anti-phosphotyrosine
antibody at 4°C. The next day the membranes were incubated with anti-mouse IgG
horseradish peroxidase (Santa Cruz, Ca) and detection was performed using enhanced
chemiluminescence (Amersham Biosciences). For accurate determination of the
molecular weight, protein samples were electrophoresed with BenchMark Ladder
(Invitrogen).
5.3
RESULTS
5.3.1 DDR1 becomes phosphorylated with morphological forms of collagen
As collagen type I is secreted from the cell as a monomeric (M) form, it
undergoes a transformation at 37°C to an intermediate semi-polymeric (SP) state, to an
eventual fibrillar F form. Figure 5.1 demonstrates that at 37°C, all three forms of this
collagen lead to phosphorylation of the down stream receptor kinase in cells over
expressing full-length DDR1. This experiment was performed to repeat previously
reported data (4). In order to maintain the integrity of the M, SP, and F species, the same
experiment was performed at 4°C. The semi-polymeric form demonstrated an increase in
phosphorylation in cells over-expressing DDR1 than the M or F forms of collagen type I.
115
5.3.2
Time to phosphorylation of semi-polymeric form of collagen
SP form of collagen type I was used to stimulate Hek293 cells over expressing
full-length DDR1. Cells were stimulated at 4°C to maintain the M and SP forms of
collagen. Fig. 5.2 shows that in as little as 15 min, phosphorylation increases in those
cells over
Fig. 5.1. Top Western shows phosphorylation of DDR1 with 0.5 µg/mL M,
SP, F collagens for 90 min at 37°C. -/- untransfected, unstimulated; +/
indicates transfected cells.
Bottom gel is phosphorylation with the presence of morphological forms of
collagen type I for 90 min at 4°C. Cells stimulated with 0.5 µg/mL collagen.
116
expressing DDR1. The Western blot indicates that maximal phosphorylation takes place
in 30 min. This experiment was designed to compare fibrillar collagen response with that
of the SP form. Previous results (4) using 10 µg/mL of monomeric collagen on cells at
37°C show phosphorylation begins at 30 min. Our results show SP collagen type I
demonstrates phosphorylation occurs in as little as 15 min at a concentration of 0.5
µg/mL. Phosphorylation band at t=0 (+/-) is also present in previously described
experiments (4,21).
117
5.3.3
Phosphorylation is not concentration dependent.
Hek293 cells over-expressing full-length DDR1 with a V5 tag were stimulated at
4°C for 90 min. With a collagen concentration as low as 1.0 µg/mL, an increase in
phosphorylation occurs in those cells over expressing DDR1 when exposed to SP form of
collagen. Figure 5.3 shows the phosphorylation pattern at these concentrations.
5.3.4
DDR1 gene silenced cells show increased phosphorylation with semi-
polymeric form of collagen type I
Hek293 cells were transfected with DDR1 siRNA. Fig. 5.4 shows a difference in
DDR1 protein expression with DDR1 siRNA (right side, bottom gel). Cells were then
stimulated with 5.0 µg/mL morphological forms of collagen type I at 4°C for 90 min.
Cells again show an increase in phosphorylation with the SP form of collagen type I,
even with a decrease in overall DDR1 cellular expression. Left side of Western are
Hek293 cells that have not
118
Fig. 5.4. DDR1 siRNA was added to cells. Bottom Western blot is of DDR1
expression. Cells were then stimulated with M, SP, or F collagen for 90 min
at 4°C. -/- HEK293 cells untransfected with siRNA and/or DDR1. Left side of
gel represents untransfected cells. Right side of gel represents cells transfected
with DDR1 siRNA.
been transfected with full-length DDR1. It has been noted (data not present here), that
Hek293 cells do possess endogenous DDR1. Fig. 5.4 is an example of that endogenous
DDR1 protein expression.
5.4
DISCUSSION
The polymeric state of collagen in the extracellular matrix (ECM) strongly
influences cell attachment and behavior. Polymerized fibers of collagen contribute to
mechanical strength of the tissue, induce cell growth, and inhibit cell proliferation.
Soluble monomeric collagen promotes cell cycle progression and proliferation (152). A
proper balance between the polymeric states of collagen is important for tissue
homeostasis, and disruption of this balance is implicated in disease. To restore
119
homeostasis to the cell, it is critical to understand the role of the collagen polymeric
species. DDR1 may play a role in collagen homeostasis.
The discoidin domain receptor tyrosine kinase 1 (DDR1), is a widely expressed
tyrosine kinase found in fibroblasts, smooth muscle cells, macrophages, and monocytes.
While other endocytic receptors specifically recognize collagenase cleaved or fibrillar
collagen, the preliminary data presented provides further evidence that DDR1 may be a
receptor that recognizes a semi-polymeric form of collagen which occurs as an
intermediate product of collagen fibrillogenesis. Collagen type I is the most abundant
protein in the ECM. Collagen secreting cells, such as fibroblasts, are responsible for
maintaining the equilibrium between collagen synthesis and degradation. This involves
secretion of monomeric (M) collagen by cells, its assembly into collagen fibers by
fibrillogenisis, and eventual breakdown by specific matrix metalloproteinases. Both the
synthesis and degradation of collagen fibers is characterized by soluble byproducts such
as excess monomeric (M) collagen and semi-polymeric (SP) collagen which have not
been assembled into fibrillar (F) collagen. It is important for these products to be cleared
rapidly from the ECM for proper connective tissue homeostasis.
Triple helical collagen is the only known ligand thus far for DDR1 (4,21). The
specific binding site on collagen where the ECD of DDR1 interacts is not yet known.
However, it has been determined that DDR1 does bind to overlapping collagen type I
molecules (143). In addition, DDR1 inhibits collagen fibrillogensis by attaching to
collagen molecules (143). It is also known that when collagen binds to the ECD of
DDR1, it induces phosphorylation of the receptor tyrosine kinase region of DDR1. This
120
phosphorylation is unusual as it requires 30 min of collagen stimulation and remains
phosphorylated for over 18 hours (4,21). The reasons behind delayed activation of DDR1
are not understood.
In the first experiment (Fig. 5.1), it was determined that the M, SP and F form of
collagen is a ligand for DDR1-induced phsophorylation. This experiment was performed
at 37 °C in which M collagen undergoes formation of fibrillar (F) collagen via the semipolymeric (SP) intermediate. However, DDR1 phosphorylation caused by stimulation
with M and SP collagen either suggests that at 37°C sufficient F collagen forms to
stimulate DDR1 phosphorylation during the 90 min course of the experiment or either
and/or both the M and SP forms can induce DDR1 phosphorylation. To maintain the
integrity of the collagen species here, conversion is impeded by repeating the experiment
at 4°C. Only the SP form of collagen induced phosphorylation. We can hypothesize that
at this low temperature, the SP species may be random and small enough to bind to the
ECD of the DDR1 and induce phosporylation of the downstream RTK. Even though F
collagen is the preferred ligand for DDR1 phosphorylation, cell and protein (atom
bombardment) movements may be decreased at this temp. Binding may occur, but time
to phosphorylation is decreased. The SP species appears to contain the DDR1 ECD
binding region, thus binding occurs quickly and the result of phosphorylation does not lag
behind.
To test this theory, Hek293 cells over expressing full length DDR1 were
subjected to stimulation with the SP species only at 4°C. Whole cell lysates were
collected every 15 min and subjected to SDS-PAGE analysis, followed by Western
121
blotting using the anti-phosphotyrosine 4G10 antibody. -/- lane represents t=0. Fig. 5.2
shows that in as little as 15 min of stimulation, phosphorylation occurs. Simulation was
performed with 0.5 µg/mL of SP collagen. This concentration of collagen has been used
previously to demonstrate DDR1 phosphorylation (4). One concern is that
phosphorylation occurs rapidly due to the concentration of collagen used, and this
concentration provides a “saturation” effect on the receptor. Fig. 5.3 represents an
experiment in which different concentrations of M, SP, and F collagens were used to
stimulate cells over expressing DDR1 for 90 min at 4°. With as little as 1.0 µg/mL of
collagen, SP is still the preferred ligand for DDR1. It is not known at this time if these
concentrations of collagen mimic the cellular environment. A concentration of 1.0 µg/mL
may still be over saturation level.
One thought to consider with these qualitative experiments is that indeed SP may
be a preferred ligand-but not for DDR1. Cells were transfected with DDR1 siRNA to
knock down DDR1 gene expression. Reducing expression of the DDR1 protein would
also decrease the phosphorylation level as well. In this experiment, DDR1 protein
expression was markedly reduced (Fig. 5.4), however, using 0.5 µg/mL of collagen for
stimulation at 90 min, once again showed that SP was the preferred ligand for
phosphorylation.
It would be prudent to determine, with the use of mass spectrometry or
immunoprecipitation, if the increase in phosphorylation is due to DDR1, and not another
similarly sized protein. The results presented here may be providing a false positive for
the role of morphological collagens and DDR1. Determining the preferred morphological
122
collagen species for DDR1 may provide clues to the regulation of the ECM and the
functioning of the cell.
123
CHAPTER 6
CONCLUSIONS AND DISCUSSIONS
A summary of the main results from the previous chapters is presented here. The
biological significance of the results will be analyzed and future directions of research
will be proposed.
DDR1 and DDR2 may bind Phospholipids as a Ligand
The data presented in the first chapter reveal a qualitative response to
phospholipid binding. It is determined, by the use of flow cytometry and SPR, that cells
over expressing DDR1 possibly show a propensity to bind to phospholipids. Both DDR1
and DDR2 show a high amino acid identity and similarity to those proteins which do bind
phosphatidylserine. However, the experiments presented here do not clarify this issue.
Knowing if phospholipids are a binding partner, the roles DDRs play in blood
coagulation where binding to a phospholipid cofactor is required may be elucidated.
Since the purified ECD of DDR1 and DDR2 are available commercially (R&D
Systems, Minneapolis, MN), a series of experiments can be designed to elucidate the
phospholipid binding partners, as well as generate binding curves and dissociation
124
constants. Such experiment is a solid-phase ELISA where it can be anticipated the
purified DDRs bind to immobilized phospholipids. Two additional experiments in which
we have a working knowledge of are flow cytometry and SPR. A large body of research
currently exists which analysis the phospholipid binding properties of blood coagulation
factor V, VIII, and lactadherin. Using phospholipids immobilized on glass microspheres,
flow cytometry can be used to determine a binding constant. In addition, SPR
experiments can and should be repeated to confirm a KD for the purified protein and the
ligand.
The Modeled Three-dimensional Structure of DDR1 and DDR2 mimics that of other
models
The three-dimensional models presented here mimic those structures in which the
solution structure or x-ray crystal structure exists-for C2 or discoidin proteins. Those
proteins being blood factor V, VIII, lactadherin, and neuropilin-1. The overall core
structure is a similar β-barrel. The loop regions which would interact with a proposed
plasma membrane of the opposing cell show significant variation. Structural data is
critical in that it allows predictions of the mechanism of protein functions.protein
functions. Since there is structural similarity between the already solved structures of the
discoidin family proteins, it can be assumed that DDR1 and DDR2 function in a similar
fashion to those other proteins which bind phospholipids.
To that end, it is imperative to know how the proteins bind to the phospholipid
ligand. It is already determined that a critical tryptophan reside in Loop 1 of the binding
125
region. This residue, when mutated, prevents phosphatidylserine from binding. This data
provides a key to the role of that loop. But what about the surrounding residues? Are
those residues important in head-group binding of the phospholipid as well? To resolve
this issue, mutagenesis can be performed, followed by analytical experiments to confirm
phospholipid binding by flow cytometry or SPR. The modeled loop 1 of DDR1 and
DDR2 indicates the essential tryptophan may be capable of touching the surface or
binding to the phospholipid head group. Where exactly would the tryptophan bind? To
resolve this issue, depth of the tryptophan inserted into the membrane could be measured.
This could be accomplished by using the parallax method (153), which has been utilized
for blood coagulation factor V (154).
It has been proposed that Loop 3 of the binding region is important for insertion
into the plasma membrane. The role of this loop is less resolved in the literature in terms
of function. It would be interesting to note how this loop interacts with the phospholipid
head and tail regions.
DDRs indirectly interfere with Platelet Aggregation
The results of the platelet aggregation experiments show that aggregation is not
affected by the DDRs alone. When DDRs are incubated with collagen, the process of
forming fibrillar collagen is interrupted (143). Impaired fibrillar collagen formation
directly affects platelet aggregation. In addition, the time to aggregation is decreased in
the presence of DDR1, but slowly increases as collagen concentration increases. DDR2 is
atypical in that lag time increases in the presence of increasing concentration of collagen.
126
DDR1 and DDR2 demonstrate > 58% identity in over all amino acid structure amongst
one another. Clearly, both involve different mechanisms as the time to aggregation is
markedly opposite. Fibrillar collagen was formed and incubated with the DDRs. After
incubation, and after washing this sample as agonist, platelet aggregation does not occur.
These results are novel in that no literature exists involving the DDRs and platelet
aggregation. No literature exists as well for the incubation of DDRs with collagen. DDRs
may play a role in the coagulation process by either binding with collagen type I and
preventing fibrillar collagen from forming, binding with the fibrillar collagen of the ECM
and preventing the collagen from interaction with the platelets, or lastly, DDRs may play
a duel role in binding and preventing both phospholipids from activated platelets and
biding of collagen-a role similar to that of lactadherin.
DDR1 binds to a Morphological form of Collagen Type I
Three forms of collagen type I exist in the ECM-M, SP, F. These three forms are
represented in the processing of collagen in the ECM, either during synthesis of M to F
form, or in the breakdown of F to M form. DDR1 may be involved in the processing
these three forms. My results indicated that an increase in phosphorylation occurs with
the SP form rather than the M and F forms. However, it is only assumed that the amount
of collagen molecules is equal in the three morphological forms of collagen. This
assumption is based on the hydroxyproline assay as M collagen moves to SP and F
forms. The SP form may be a small molecule compared to F form, but may contain the
essential sites for binding to DDR1. The site on collagen type I where DDR2 binds has
127
been resolved (41). However, the site on collagen where DDR1 one resides had only been
partially resolved (143) by the use of Atomic Force Microscopy (AFM). Following the
lead of Leitinger (41), AFM would be an appropriate initial method for determining the
DDR1 binding site on collagen.
This work represents a small fraction of roles the DDRs may play in the body. DDRs
may have implications in blood coagulation, coagulation diseases, bleeding disorders,
cell remodeling etc.
128
REFERENCES
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
Vogel, W. (1999) Faseb J 13 Suppl, S77-82
Tait, J. F., and Smith, C. (1999) J Biol Chem 274, 3048-3054
Leitinger, B. (2003) J Biol Chem 278, 16761-16769
Shrivastava, A., Radziejewski, C., Campbell, E., Kovac, L., McGlynn, M., Ryan,
T. E., Davis, S., Goldfarb, M. P., Glass, D. J., Lemke, G., and Yancopoulos, G. D.
(1997) Mol Cell 1, 25-34
Manning, G., Whyte, D. B., Martinez, R., Hunter, T., and Sudarsanam, S. (2002)
Science 298, 1912-1934
Blume-Jensen, P., and Hunter, T. (2001) Nature 411, 355-365
Zwick, E., Bange, J., and Ullrich, A. (2002) Trends Mol Med 8, 17-23
Pawson, T. (2002) Eur J Cancer 38 Suppl 5, S3-10
Oda, K., Matsuoka, Y., Funahashi, A., and Kitano, H. (2005) Mol Syst Biol 1,
2005 0010
Sorensen, O. E., Thapa, D. R., Roupe, K. M., Valore, E. V., Sjobring, U., Roberts,
A. A., Schmidtchen, A., and Ganz, T. (2006) J Clin Invest 116, 1878-1885
Walker, F., Abramowitz, L., Benabderrahmane, D., Duval, X., Descatoire, V.,
Henin, D., Lehy, T., and Aparicio, T. (2009) Hum Pathol 40, 1517-1527
Ornitz, D. M., and Itoh, N. (2001) Genome Biol 2, REVIEWS3005
Riley, B. M., Mansilla, M. A., Ma, J., Daack-Hirsch, S., Maher, B. S.,
Raffensperger, L. M., Russo, E. T., Vieira, A. R., Dode, C., Mohammadi, M.,
Marazita, M. L., and Murray, J. C. (2007) Proc Natl Acad Sci U S A 104, 45124517
Kaess, B. M., Barnes, T. A., Stark, K., Charchar, F. J., Waterworth, D., Song, K.,
Wang, W. Y., Vollenweider, P., Waeber, G., Mooser, V., ZukowskaSzczechowska, E., Samani, N. J., Hengstenberg, C., and Tomaszewski, M. Eur J
Hum Genet 18, 1344-1348
Fujita, N., Imai, J., Suzuki, T., Yamada, M., Ninomiya, K., Miyamoto, K.,
Iwasaki, R., Morioka, H., Matsumoto, M., Chiba, K., Watanabe, S., Suda, T.,
Toyama, Y., and Miyamoto, T. (2008) Biochem Biophys Res Commun 372, 367372
Holmes, K., Roberts, O. L., Thomas, A. M., and Cross, M. J. (2007) Cell Signal
19, 2003-2012
Nakagawara, A. (1998) Med Pediatr Oncol 31, 113-115
Nakagawara, A., Liu, X. G., Ikegaki, N., White, P. S., Yamashiro, D. J., Nycum,
L. M., Biegel, J. A., and Brodeur, G. M. (1995) Genomics 25, 538-546
129
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
34.
35.
36.
37.
38.
39.
40.
41.
42.
Rosen, S. D., Kafka, J. A., Simpson, D. L., and Barondes, S. H. (1973) Proc Natl
Acad Sci U S A 70, 2554-2557
Springer, W. R., Cooper, D. N., and Barondes, S. H. (1984) Cell 39, 557-564
Vogel, W., Gish, G. D., Alves, F., and Pawson, T. (1997) Mol Cell 1, 13-23
Kamohara, H., Yamashiro, S., Galligan, C., and Yoshimura, T. (2001) Faseb J 15,
2724-2726
Alves, F., Vogel, W., Mossie, K., Millauer, B., Hofler, H., and Ullrich, A. (1995)
Oncogene 10, 609-618
Barker, K. T., Martindale, J. E., Mitchell, P. J., Kamalati, T., Page, M. J.,
Phippard, D. J., Dale, T. C., Gusterson, B. A., and Crompton, M. R. (1995)
Oncogene 10, 569-575
Laval, S., Butler, R., Shelling, A. N., Hanby, A. M., Poulsom, R., and Ganesan, T.
S. (1994) Cell Growth Differ 5, 1173-1183
Weiner, H. L., Rothman, M., Miller, D. C., and Ziff, E. B. (1996) Pediatr
Neurosurg 25, 64-71; discussion 71-62
(1999) Nature 401, 921-923
Alves, F., Saupe, S., Ledwon, M., Schaub, F., Hiddemann, W., and Vogel, W. F.
(2001) Faseb J 15, 1321-1323
Perez, J. L., Shen, X., Finkernagel, S., Sciorra, L., Jenkins, N. A., Gilbert, D. J.,
Copeland, N. G., and Wong, T. W. (1994) Oncogene 9, 211-219
Ferri, N., Carragher, N. O., and Raines, E. W. (2004) Am J Pathol 164, 15751585
Karn, T., Holtrich, U., Brauninger, A., Bohme, B., Wolf, G., RubsamenWaigmann, H., and Strebhardt, K. (1993) Oncogene 8, 3433-3440
Lai, C., and Lemke, G. (1994) Oncogene 9, 877-883
Abdulhussein, R., McFadden, C., Fuentes-Prior, P., and Vogel, W. F. (2004) J
Biol Chem 279, 31462-31470
Baumgartner, S., Hofmann, K., Chiquet-Ehrismann, R., and Bucher, P. (1998)
Protein Sci 7, 1626-1631
Kane, W. H., and Davie, E. W. (1988) Blood 71, 539-555
Ichikawa, O., Osawa, M., Nishida, N., Goshima, N., Nomura, N., and Shimada, I.
(2007) Embo J 26, 4168-4176
Macedo-Ribeiro, S., Bode, W., Huber, R., Quinn-Allen, M. A., Kim, S. W., Ortel,
T. L., Bourenkov, G. P., Bartunik, H. D., Stubbs, M. T., Kane, W. H., and
Fuentes-Prior, P. (1999) Nature 402, 434-439
Pratt, K. P., Shen, B. W., Takeshima, K., Davie, E. W., Fujikawa, K., and
Stoddard, B. L. (1999) Nature 402, 439-442
Shao, C., Novakovic, V. A., Head, J. F., Seaton, B. A., and Gilbert, G. E. (2008) J
Biol Chem 283, 7230-7241
Kiedzierska, A., Smietana, K., Czepczynska, H., and Otlewski, J. (2007) Biochim
Biophys Acta 1774, 1069-1078
Leitinger, B., Steplewski, A., and Fertala, A. (2004) J Mol Biol 344, 993-1003
Gaskell, A., Crennell, S., and Taylor, G. (1995) Structure 3, 1197-1205
130
43.
44.
45.
46.
47.
48.
49.
50.
51.
52.
53.
54.
55.
56.
57.
58.
59.
60.
61.
62.
63.
64.
65.
66.
67.
Ito, N., Phillips, S. E., Yadav, K. D., and Knowles, P. F. (1994) J Mol Biol 238,
794-814
Bianchet, M. A., Odom, E. W., Vasta, G. R., and Amzel, L. M. (2002) Nat Struct
Biol 9, 628-634
Lee, C. C., Kreusch, A., McMullan, D., Ng, K., and Spraggon, G. (2003)
Structure 11, 99-108
Vander Kooi, C. W., Jusino, M. A., Perman, B., Neau, D. B., Bellamy, H. D., and
Leahy, D. J. (2007) Proc Natl Acad Sci U S A 104, 6152-6157
Fuentes-Prior, P., Fujikawa, K., and Pratt, K. P. (2002) Curr Protein Pept Sci 3,
313-339
Srivastava, A., Wang, J., Majumder, R., Rezaie, A. R., Stenflo, J., Esmon, C. T.,
and Lentz, B. R. (2002) J Biol Chem 277, 1855-1863
Leitinger, B., and Kwan, A. P. (2006) Matrix Biol 25, 355-364
Agarwal, G., Kovac, L., Radziejewski, C., and Samuelsson, S. J. (2002)
Biochemistry 41, 11091-11098
Konitsiotis, A. D., Raynal, N., Bihan, D., Hohenester, E., Farndale, R. W., and
Leitinger, B. (2008) J Biol Chem 283, 6861-6868
Curat, C. A., and Vogel, W. F. (2002) J Am Soc Nephrol 13, 2648-2656
Bartles, J. R., Frazier, W. A., and Rosen, S. D. (1982) Int Rev Cytol 75, 61-99
Bartles, J. R., Santoro, B. C., and Frazier, W. A. (1981) Biochim Biophys Acta
674, 372-382
Bartles, J. R., and Frazier, W. A. (1980) J Biol Chem 255, 30-38
Koo, D. H., McFadden, C., Huang, Y., Abdulhussein, R., Friese-Hamim, M., and
Vogel, W. F. (2006) FEBS Lett 580, 15-22
Matsuyama, W., Faure, M., and Yoshimura, T. (2003) J Immunol 171, 3520-3532
Foehr, E. D., Tatavos, A., Tanabe, E., Raffioni, S., Goetz, S., Dimarco, E., De
Luca, M., and Bradshaw, R. A. (2000) Faseb J 14, 973-981
Vogel, W., Lammers, R., Huang, J., and Ullrich, A. (1993) Science 259, 16111614
L'Hote C, G., Thomas, P. H., and Ganesan, T. S. (2002) Faseb J 16, 234-236
Olaso, E., Labrador, J. P., Wang, L., Ikeda, K., Eng, F. J., Klein, R., Lovett, D. H.,
Lin, H. C., and Friedman, S. L. (2002) J Biol Chem 277, 3606-3613
Yang, K., Kim, J. H., Kim, H. J., Park, I. S., Kim, I. Y., and Yang, B. S. (2005) J
Biol Chem 280, 39058-39066
Faraci, E., Eck, M., Gerstmayer, B., Bosio, A., and Vogel, W. F. (2003) Matrix
Biol 22, 373-381
Freyman, T. M., Yannas, I. V., Yokoo, R., and Gibson, L. J. (2002) Exp Cell Res
272, 153-162
Ongusaha, P. P., Kim, J. I., Fang, L., Wong, T. W., Yancopoulos, G. D.,
Aaronson, S. A., and Lee, S. W. (2003) Embo J 22, 1289-1301
Dejmek, J., Dib, K., Jonsson, M., and Andersson, T. (2003) Int J Cancer 103,
344-351
Myllyharju, J., and Kivirikko, K. I. (2001) Ann Med 33, 7-21
131
68.
69.
70.
71.
72.
73.
74.
75.
76.
77.
78.
79.
80.
81.
82.
83.
84.
85.
86.
87.
88.
89.
90.
91.
92.
93.
94.
95.
96.
Brown, R. E., Butler, J. P., Rogers, R. A., and Leith, D. E. (1994) Connect Tissue
Res 30, 295-308
Lee, G. M., and Loeser, R. F. (1999) Exp Cell Res 248, 294-305
Prockop, D. J., and Kivirikko, K. I. (1995) Annu Rev Biochem 64, 403-434
Kadler, K. (1995) Protein Profile 2, 491-619
Kivirikko, K. I., and Risteli, L. (1976) Med Biol 54, 159-186
Kadler, K. E., Hojima, Y., and Prockop, D. J. (1987) J Biol Chem 262, 1569615701
Kadler, K. E., Holmes, D. F., Trotter, J. A., and Chapman, J. A. (1996) Biochem J
316 ( Pt 1), 1-11
Parkinson, J., Kadler, K. E., and Brass, A. (1995) J Mol Biol 247, 823-831
Hook, F., Stengel, G., Dahlin, A. B., Gunnarsson, A., Jonsson, M. P., Jonsson, P.,
Reimhult, E., Simonsson, L., and Svedhem, S. (2008) Biointerphases 3, FA108
Chikh, G. G., Li, W. M., Schutze-Redelmeier, M. P., Meunier, J. C., and Bally,
M. B. (2002) Biochim Biophys Acta 1567, 204-212
Jesorka, A., and Orwar, O. (2008) Annu Rev Anal Chem (Palo Alto Calif) 1, 801832
Aktas, M., Wessel, M., Hacker, S., Klusener, S., Gleichenhagen, J., and
Narberhaus, F. Eur J Cell Biol 89, 888-894
Buckland, A. G., and Wilton, D. C. (2000) Biochim Biophys Acta 1483, 199-216
Chaurio, R. A., Janko, C., Munoz, L. E., Frey, B., Herrmann, M., and Gaipl, U. S.
(2009) Molecules 14, 4892-4914
Ross, R. (1993) Nature 362, 801-809
Ross, R. (1999) N Engl J Med 340, 115-126
Finn, A. V., Nakano, M., Narula, J., Kolodgie, F. D., and Virmani, R. Arterioscler
Thromb Vasc Biol 30, 1282-1292
Didangelos, A., Simper, D., Monaco, C., and Mayr, M. (2009) Curr Atheroscler
Rep 11, 188-195
Ruggeri, Z. M. (1997) Thromb Haemost 78, 611-616
Rinehart, J. F., and Greenberg, L. D. (1949) Am J Pathol 25, 481-491
Shyu, K. G., Chao, Y. M., Wang, B. W., and Kuan, P. (2005) Hypertension 46,
614-621
Alberts, B. Mol Biol Cell 21, 3785
Johnson, J. E., and Cornell, R. B. (1999) Mol Membr Biol 16, 217-235
Thuduppathy, G. R., Craig, J. W., Kholodenko, V., Schon, A., and Hill, R. B.
(2006) J Mol Biol 359, 1045-1058
Comfurius, P., Bevers, E., and Zwaal, R. F. (1994) Methods Mol Biol 27, 131-142
Gilbert, G. E., and Arena, A. A. (1995) J Biol Chem 270, 18500-18505
Gilbert, G. E., Kaufman, R. J., Arena, A. A., Miao, H., and Pipe, S. W. (2002) J
Biol Chem 277, 6374-6381
Bangham, A. D., Standish, M. M., and Watkins, J. C. (1965) J Mol Biol 13, 238252
Johnson, S. M., Bangham, A. D., Hill, M. W., and Korn, E. D. (1971) Biochim
Biophys Acta 233, 820-826
132
97.
98.
99.
100.
101.
102.
103.
104.
105.
106.
107.
108.
109.
110.
111.
112.
113.
114.
115.
116.
117.
118.
119.
120.
121.
122.
Saenko, E., Sarafanov, A., Ananyeva, N., Behre, E., Shima, M., Schwinn, H., and
Josic, D. (2001) J Chromatogr A 921, 49-56
Curat, C. A., Eck, M., Dervillez, X., and Vogel, W. F. (2001) J Biol Chem 276,
45952-45958
Kim, S. W., Quinn-Allen, M. A., Camp, J. T., Macedo-Ribeiro, S., Fuentes-Prior,
P., Bode, W., and Kane, W. H. (2000) Biochemistry 39, 1951-1958
Adiguzel, E., Ahmad, P. J., Franco, C., and Bendeck, M. P. (2009) Vasc Med 14,
73-89
Dasgupta, S. K., Abdel-Monem, H., Niravath, P., Le, A., Bellera, R. V., Langlois,
K., Nagata, S., Rumbaut, R. E., and Thiagarajan, P. (2009) Blood 113, 1332-1339
Barrow, R. T., Healey, J. F., Jacquemin, M. G., Saint-Remy, J. M., and Lollar, P.
(2001) Blood 97, 169-174
Pellequer, J. L., Gale, A. J., and Getzoff, E. D. (2000) Curr Biol 10, R237-240
Griffin, F. M., Jr. (1981) Proc Natl Acad Sci U S A 78, 3853-3857
Abbott, A. J., and Nelsestuen, G. L. (1987) Biochemistry 26, 7994-8003
Peng, W., Quinn-Allen, M. A., Kim, S. W., Alexander, K. A., and Kane, W. H.
(2004) Biochemistry 43, 4385-4393
Mann, K. G., Nesheim, M. E., Church, W. R., Haley, P., and Krishnaswamy, S.
(1990) Blood 76, 1-16
Zwaal, R. F., Comfurius, P., and Bevers, E. M. (1998) Biochim Biophys Acta
1376, 433-453
Krishnaswamy, S., and Mann, K. G. (1988) J Biol Chem 263, 5714-5723
Kim, S. W., Ortel, T. L., Quinn-Allen, M. A., Yoo, L., Worfolk, L., Zhai, X.,
Lentz, B. R., and Kane, W. H. (1999) Biochemistry 38, 11448-11454
Hou, G., Vogel, W. F., and Bendeck, M. P. (2002) Circ Res 90, 1147-1149
Leitinger, B., and Hohenester, E. (2007) Matrix Biol 26, 146-155
Rosing, J., Tans, G., Govers-Riemslag, J. W., Zwaal, R. F., and Hemker, H. C.
(1980) J Biol Chem 255, 274-283
van Dieijen, G., Tans, G., Rosing, J., and Hemker, H. C. (1981) J Biol Chem 256,
3433-3442
Ortel, T. L., Devore-Carter, D., Quinn-Allen, M., and Kane, W. H. (1992) J Biol
Chem 267, 4189-4198
Ortel, T. L., Quinn-Allen, M. A., Keller, F. G., Peterson, J. A., Larocca, D., and
Kane, W. H. (1994) J Biol Chem 269, 15898-15905
Lewis, D. A., Moore, K. D., and Ortel, T. L. (2003) Blood Coagul Fibrinolysis
14, 361-368
Berman, H. M., Westbrook, J., Feng, Z., Gilliland, G., Bhat, T. N., Weissig, H.,
Shindyalov, I. N., and Bourne, P. E. (2000) Nucleic Acids Res 28, 235-242
Laskowski, R. A., Moss, D. S., and Thornton, J. M. (1993) J Mol Biol 231, 10491067
Laskowski, R. A. Methods Mol Biol 609, 59-82
Chothia, C., and Lesk, A. M. (1986) Embo J 5, 823-826
Perera, L., Darden, T. A., and Pedersen, L. G. (1999) Biophys J 77, 99-113
133
123.
124.
125.
126.
127.
128.
129.
130.
131.
132.
133.
134.
135.
136.
137.
138.
139.
140.
141.
142.
143.
144.
145.
146.
147.
148.
Perera, L., Li, L., Darden, T., Monroe, D. M., and Pedersen, L. G. (1997) Biophys
J 73, 1847-1856
Li, L., Darden, T., Foley, C., Hiskey, R., and Pedersen, L. (1995) Protein Sci 4,
2341-2348
Novakovic, V. A., Cullinan, D. B., Wakabayashi, H., Fay, P. J., Baleja, J. D., and
Gilbert, G. E. Biochem J 435, 187-196
Amento, E. P., Ehsani, N., Palmer, H., and Libby, P. (1991) Arterioscler Thromb
11, 1223-1230
Majesky, M. W., Lindner, V., Twardzik, D. R., Schwartz, S. M., and Reidy, M. A.
(1991) J Clin Invest 88, 904-910
Strauss, B. H., Chisholm, R. J., Keeley, F. W., Gotlieb, A. I., Logan, R. A., and
Armstrong, P. W. (1994) Circ Res 75, 650-658
Nakagaki, T., Foster, D. C., Berkner, K. L., and Kisiel, W. (1991) Biochemistry
30, 10819-10824
Davie, E. W., and Ratnoff, O. D. (1964) Science 145, 1310-1312
Mann, K. G. (1999) Thromb Haemost 82, 165-174
Kawamoto, Y., and Kaibara, M. (1990) Biochim Biophys Acta 1035, 361-368
Gilbert, G. E., Drinkwater, D., Barter, S., and Clouse, S. B. (1992) J Biol Chem
267, 15861-15868
Gilbert, G. E., Sims, P. J., Wiedmer, T., Furie, B., Furie, B. C., and Shattil, S. J.
(1991) J Biol Chem 266, 17261-17268
Bennett, J. S., Vilaire, G., and Burch, J. W. (1981) J Clin Invest 68, 981-987
Addo, J. B., Bray, P. F., Grigoryev, D., Faraday, N., and Goldschmidt-Clermont,
P. J. (1995) Arterioscler Thromb Vasc Biol 15, 1466-1473
Cooke, G. E., Bray, P. F., Hamlington, J. D., Pham, D. M., and GoldschmidtClermont, P. J. (1998) Lancet 351, 1253
Mustard, J. F., Perry, D. W., Ardlie, N. G., and Packham, M. A. (1972) Br J
Haematol 22, 193-204
Kuznetsova, N., Chi, S. L., and Leikin, S. (1998) Biochemistry 37, 11888-11895
Salchert, K., Streller, U., Pompe, T., Herold, N., Grimmer, M., and Werner, C.
(2004) Biomacromolecules 5, 1340-1350
Born, G. V. (1962) Nature 194, 927-929
Mihai, C., Chotani, M., Elton, T. S., and Agarwal, G. (2009) J Mol Biol 385, 432445
Agarwal, G., Mihai, C., and Iscru, D. F. (2007) J Mol Biol 367, 443-455
Mihai, C., Iscru, D. F., Druhan, L. J., Elton, T. S., and Agarwal, G. (2006) J Mol
Biol 361, 864-876
Clowes, A. W., Reidy, M. A., and Clowes, M. M. (1983) Lab Invest 49, 327-333
Zempo, N., Koyama, N., Kenagy, R. D., Lea, H. J., and Clowes, A. W. (1996)
Arterioscler Thromb Vasc Biol 16, 28-33
Jester, J. V., Petroll, W. M., Barry, P. A., and Cavanagh, H. D. (1995) J Anat 186
( Pt 2), 301-311
Yannas, I. V., Lee, E., Orgill, D. P., Skrabut, E. M., and Murphy, G. F. (1989)
Proc Natl Acad Sci U S A 86, 933-937
134
149.
150.
151.
152.
153.
154.
Veis, A. (1994) Connect Tissue Res 31, i
Silver, D., and Miller, J. (1994) J Mol Graph 12, 139-145
Baldock, C., Gilpin, C. J., Koster, A. J., Ziese, U., Kadler, K. E., Kielty, C. M.,
and Holmes, D. F. (2002) J Struct Biol 138, 130-136
Berrier, A. L., and Yamada, K. M. (2007) J Cell Physiol 213, 565-573
Chattopadhyay, A., and London, E. (1987) Biochemistry 26, 39-45
Majumder, R., Quinn-Allen, M. A., Kane, W. H., and Lentz, B. R. (2005)
Biochemistry 44, 711-718
135