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Myosin Phosphatase Rho-interacting Protein Regulates DDR1-mediated Collagen Tractional Remodeling
by
Petar B. Petrovic
A thesis submitted in conformity with the requirements for the degree of Master of Science Graduate Department of Dentistry
University of Toronto © Copyright by Petar B. Petrovic 2017
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Myosin Phosphatase Rho-interacting Protein Regulates DDR1-
mediated Collagen Tractional Remodeling
Petar B. Petrovic
Master of Science
Faculty of Dentistry
University of Toronto
2017
Abstract
Collagen remodeling is essential for preserving the structure and function of highly ordered collagen
arrays, which are important for the normal function of many tissues and organs. Remodeling of fibrillar
collagen is mediated in part by α2β1 and α11β1 integrins, and by the discoidin domain receptors (DDR) 1
and 2, which are receptor tyrosine kinases activated by binding to fibrillar collagen. The mechanisms by
which contractile forces generated by non-muscle myosin IIA (NMIIA) are regulated to mediate DDR1-
dependent collagen remodeling are not defined. We considered that cell-induced tractional remodeling of
type I fibrillar collagen is strongly dependent on DDR1 activity and its interaction with NMIIA. By mass
spectrometry and immunoprecipitation we found a time and collagen-dependent association between
DDR1 and Myosin Phosphatase Rho-interacting Protein (MRIP). This association was strongly reduced
by inhibiting DDR1 tyrosine autophosphorylation. MRIP exhibited strong, time and collagen-dependent
co-localization with actin stress fibers, DDR1 and NMIIA. Deletion of MRIP by CRISPR-Cas9 enhanced
co-localization of DDR1 with NMIIA and increased myosin light chain and DDR1 phosphorylation in a
collagen-dependent manner. Increased expression of DDR1, deletion of MRIP, reducing DDR1
activation, or inhibition of Rho-associated coiled coil kinase, all strongly affected cell adhesion to
collagen and the contraction, alignment and compaction of collagen fibrils. Collectively, these data
indicate that DDR1, NMIIA and MRIP comprise a critical, interacting molecular complex that mediates
collagen remodeling by DDR1 adhesions.
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Acknowledgments
These past two years have probably been the most difficult that I’ve yet to experience. It seems
like it was just last week that I happened to stumble into Chris’ office to talk about getting into
dental school at UofT; a conversation that lasted all of 15 minutes before it devolved into an hour
long conversation about everything and nothing at the same time, like an episode of Seinfeld.
The first, and definitely most important, person I have to thank is my supervisor Chris. In
all honesty, I have no idea where I would be had it not been for his guidance, support, and
wisdom. I was quite fortunate to have a desk close by to Chris and there was never a time where
his door was not open to not only discuss science but life, comedy, politics and everything in
between. I’m truly grateful for everything that he has done for me, I’m very lucky to have been
given the opportunity to work with this lab and group.
I’d like to thank my lab members Dhaarmini, Hamid, Nuno, Pam, Qin, and Wilson who
were absolutely instrumental in helping me achieve the goals I had set out at the beginning of
this journey. I’d also like to thank Jonathan Krieger, a mass spectrometry expert working at
Toronto’s Hospital for Sick Children who helped with the initial experiment which set my
project in motion and for the help he provided in writing up the mass spectrometry protocol.
Usually at the end of these acknowledgement sections people thank their friends and
family but mine didn’t really help with any of the experiments, or writing for that matter…lazy!
So I’m going to take this opportunity to thank myself. Thanks Pete, couldn’t have done it without
you buddy.
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Table of Contents
Abstract……………………………………………………………………………......................ii
Acknowledgements……………………………………………………………………….....…..iii
Tables of Contents…………………………………………………………...……………..…...iv
List of Diagrams……………………………………………………………………………...….vi
List of Figures…….……….……….……….……….……….……….……….……….….........vii
Chapter 1 - Literature review.......................................................................................................1
1. Connective tissues........................................................................................................1
1.1 Roles of connective tissue in health and disease.....................................................1
1.2 Proteins of the extracellular matrix..........................................................................2
1.2.1 Various proteins make up the ECM.............................................................3
1.2.2 Types of collagen.........................................................................................5
1.3 Structure and function of type I collagen.................................................................6
1.3.1 Structure.......................................................................................................6
1.3.2 Function.......................................................................................................7
2. Homeostatic regulation of collagen............................................................................9
2.1 Synthesis..................................................................................................................9
2.2 Collagen remodeling..............................................................................................10
2.2.1 Collagen Proteolysis..................................................................................11
2.2.2 Phagocytosis..............................................................................................12
2.2.3 Traction......................................................................................................13
3. Fibrillar collagen receptors.......................................................................................14
3.1 b1 Integrin..............................................................................................................14
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3.1.1 Structure and isoforms of collagen-binding integrins................................14
3.1.2 Function in collagen remodeling...............................................................16
3.2 Discoidin Domain Receptors (DDRs)...................................................................17
3.2.1 Localization, expression and structure.......................................................17
3.2.2 DDR function in fibrotic disorders............................................................20
4. Cytoskeleton cell-generated collagen remodeling...................................................21
4.1 Actin.......................................................................................................................21
4.1.1 Structure, isoforms and function................................................................21
4.1.2 Actin-binding proteins...............................................................................23
4.2 Non-muscle myosin II............................................................................................24
4.2.1 Structure, isoforms and function................................................................24
4.2.2 Mechanisms of activation..........................................................................25
Statement of the problem............................................................................................................27
Chapter 2 - Introduction.............................................................................................................28
Materials and methods................................................................................................................30
Results...........................................................................................................................................36
Discussion.....................................................................................................................................47
Figures...........................................................................................................................................52
Future directions..........................................................................................................................65
Conclusions...................................................................................................................................66
References.....................................................................................................................................68
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List of Diagrams
Diagram 1. Formation of the supramolecular structure of type I fibrillar collagen………….….10
Diagram 2. The β1 integrin family of extracellular matrix receptors……………………………15
Diagram 3. Structure and isoforms of DDR1 and DDR2……………………………………….19
Diagram 4. Structure of NMII and RLC kinases..........................................................................26
Diagram 5. Physiological mechanism by which MRIP regulates DDR1/NMIIA adhesions in
collagen remodeling.......................................................................................................................67
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List of Figures
Figure 1. .......................................................................................................................................53
A. Immunoblot of b1 integrin
B. Immunoblot of DDR1
Figure 2. .......................................................................................................................................53
A. Tables showing MS/MS experimental set up and protein abundances
B. Histogram of NMIIA abundance
C. Histogram of MRIP abundance
D. Immunoblot of MRIP in DDR1 and NMIIA immunoprecipitates
Figure 3. .......................................................................................................................................54
A. Immunofluorescence images of MRIP and F-actin localization with histogram
B. Immunofluorescence images of MRIP, DDR1, and NMIIA localization with
histogram
C. Immunofluorescence images of DDR1 and NMIIA localization with histogram
Figure 4. .......................................................................................................................................55
A. Immunofluorescence images of MRIP and DDR1 localization with histogram
B. Immunofluorescence images of MRIP and NMIIA localization with histogram
Figure 5. .......................................................................................................................................56
A. Immunoblot of MRIP expression in GD25 and MRIP KO cells
B. Immunoblot of DDR1 expression in GD25 and MRIP KO cells
Figure 6. .......................................................................................................................................57
A. Immunofluorescence images of DDR1 and NMIIA localization with histogram
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Figure 7. .......................................................................................................................................58
A. Immunoblots of GAPDH, Gelsolin and Vimentin in protein fractionation assay
Figure 8. .......................................................................................................................................59
A. Immunoblot of DDR1 fractionation with histogram
B. Immunoblot of MRIP fractionation with histogram
C. Immunoblot of NMIIA fractionation with histogram
Figure 9. .......................................................................................................................................60
A. Collagen bead binding in shear wash assay in the presence of inhibitor compounds
B. Influence of DDR1 and MRIP expression on collagen bead binding in shear wash
assay
Figure 10. .....................................................................................................................................61
A. Collagen gel contraction in the presence of inhibitor compounds
B. Influence of MRIP KO on collagen gel contraction
Figure 11. .....................................................................................................................................62
A. Confocal images of GD25 cells remodeling fibrillar collagen
B. Histogram of collagen fiber alignment and compaction in GD25 cells
C. Histogram of collagen fiber alignment and compaction in B1 MRIP KO cells
Figure 12. .....................................................................................................................................63
A. Immunoblot of total DDR1 expression with histogram
B. Immunoblot of Y792 phospho-DDR1 expression with histogram
C. Immunoblot of Y513 phospho-DDR1 expression with histogram
Figure 13. .....................................................................................................................................64
A. Immunoblot of total NMIIA expression with histogram
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B. Immunoblot of total MLC expression with histogram
C. Immunoblot of S19 phospho-MLC expression with histogram
1
Chapter 1 Literature Review
Connective Tissues 1.1 Roles of connective tissues in health and disease Connective tissues play important functional roles in providing support and attachment for
organs, muscles, bones, ligaments, and fascia (1). The extracellular matrix (ECM) is a major
structural component of many connective tissues and provides scaffolding for cells and the
structures that provide organization for several types of tissue compartments (2). ECM molecules
play vital roles in the differentiation, proliferation, survival, polarity and migration of cells (1).
Some of the proteins that comprise the ECM can act as ligands for cell adhesion and can initiate
intracellular signals from membrane-associated adhesion receptors, which act in a similar fashion
as soluble growth factors that bind their cognate receptors to initiate signal transduction. The
genome of many vertebrates encodes for a large array of ECM molecules that include, but are
not limited to glycosaminoglycans, proteoglycans, laminins, fibronectin and collagens (3), (1),
(4). The structure and function of connective tissues are intimately determined by the
organization and distribution of these ECM molecules.
In health, the maintenance of the structural integrity of the ECM is dependent on a
number of integrated, homeostatic processes. In this context, ECM proteins are synthesized,
secreted and organized into tissue-specific scaffolds, some of which are maintained by
mechanically-mediated remodeling processes. During these remodeling processes, ECM proteins
are frequently degraded; these synthetic and degradative processes are iteratively repeated to
enable the preservation of tissue structure and function (3). Arising from the complex nature of
the mechanisms that mediate tissue homeostasis, there are multiple control points at which these
mechanisms can become dysregulated. For example, at sites of chronic inflammation or when
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there has been repeated structural damage, the homeostatic systems that regulate ECM structure
and function may be perturbed, which frequently is associated with alterations of normal matrix
structure as is seen in fibrosis.
The structural composition of tissues and organs is strongly related to their function. The
kidneys for example require tight regulation of structure, molecular composition and anatomical
arrangement of the glomerulus to properly filter blood. The lungs require a very specific tissue
architecture to facilitate the proper exchange of oxygen and carbon dioxide across the alveolar
wall. In general, the structural, functional and mechanical properties (e.g. elastic modulus) of
each organ are determined in part by the molecular composition and structural characteristics of
the ECM (5).
Disruptions of the structure and composition of ECM proteins or of the signaling systems
that are regulated by ECM-cell interactions can contribute to the pathogenesis of a wide variety
of disorders that include for example, idiopathic pulmonary fibrosis, idiopathic cardiomyopathy
and invasive ductal breast cancer (2), (6, 7), (8). Accordingly, compromises to the composition
and structure of the ECM of multiple organs can profoundly affect organ function and the overall
health of mammals.
1.2 Proteins of the extracellular matrix While the ECM in general is composed of a broad range of different polysaccharides, proteins,
proteoglycans and glycosaminoglycans, each specific tissue is comprised of unique sets of ECM
molecules, each with its own distinct composition and organization. This framework is
determined during tissue development by dynamic and reciprocal processes, which involve
tightly regulated communication between a large array of cell types (e.g. epithelial cells,
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fibroblasts, adipocytes, osteoblasts, chondrocytes) and regulatory molecules such as growth
factors, hormones and prostaglandins (5).
1.2.1 Various proteins make up the ECM Proteoglycans, glycosaminoglycans and a large array of fibrous proteins comprise some of the
most abundant classes of macromolecules that make up the ECM. Collagens, laminins,
fibronectins and elastins contribute to the formation of the main fibrous ECM proteins (3). I will
briefly consider some of these proteins below. Notably, the organization of the interstitial ECM
is affected strongly by the glycoprotein, fibronectin. There are three types of fibronectin (FN):
types I, II, and III, which contribute to cell attachment and the function of many types of
anchorage-dependent cells (9). There are 12 rodent and bovine FN isoforms and 20 isoforms in
humans, which are encoded by a single fibronectin gene. Exon skipping at EIIIA/EDA and
EIIIB/EDB and exon subdivision at the V region/IIICS provides alternative splice variants of
fibronectin (10). In addition to its role in cell adhesion, fibronectin is implicated as a
mechanoregulator, possibly as a result of its force-dependent unfolding, which is involved in the
regulation of fibronectin binding to integrins (α6β1, α8β1, αVβ3) and which may affect cell
behavior (5, 11). Integrin binding stimulates fibronectin self-association, which is mediated by
N-terminal assembly domain and by integrin binding to fibronectin. The assembly of fibronectin
along with its binding to integrins can enable the transmission of cell-generated contractile forces
through integrin-linked actin cytoskeletal proteins. Further, fibronectin is important for cell
migration during developmental and wound healing processes. Fibronectin fulfills many of its
multiple functions in ECM biology by virtue of its ability, which arises from its multi-domain
structure, to adhere to membrane-associated integrins and to other ECM proteins like collagen
(9). Fibronectin is implicated in fundamental, pathological processes, such as tumor metastasis
and pressure overload-associated cardiovascular diseases (12), (13), (14). Notably, fibronectin
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expression levels may contribute to disease processes as in certain types of tumor cells, there are
reduced levels of fibronectin expression whereas fibronectin levels are elevated in tissues
undergoing repair and in fibrotic lesions (9).
Laminins are a family of large (~ 500 – 800 kDa), T-shaped heterotrimeric proteins,
which are comprised of a combination of α-, β-, and γ-chains. These proteins are important
constituents of the basal lamina, an important, ultrastructurally-defined region of the basement
membrane (15). Sixteen trimeric laminin isoforms have been described in mouse and human
tissues, many of which exhibit cell and tissue-dependent specificities (16). Contributing to their
heterogeneity of molecular mass and large size is the fact that all three chains are often highly
and variably glycosylated (17). Binding of the G domain of the α chains to integrins,
dystroglycan, or sulfated glycolipids is the general mechanism by which laminins adhere to cells
and to type IV collagen (18), another important constituent of the basal lamina. The α1 and α2
laminin chains contain N-terminal globular domains that can bind several integrins (α1β1, α2β1,
α3β1, α6β1, α6β4, α7β1and αVβ3) (9) (19), a feature of laminins that enables cell binding to
both ends of laminins. The N-terminal globular domains of some α-chains can also bind
sulfatides (20), a feature that may also enable laminin binding to the cell surface.
Laminins contribute to the structure of the ECM and influence cell adhesion,
differentiation and migration (21), (22). Tight regulation of the structural integrity of laminins is
essential for appropriate tissue maintenance and indeed overall organismal survival.
Inappropriate processing and regulation of laminins has been associated with several tissue
disorders including nephrotic syndrome, Crohn’s and ulcerative colitis, certain forms of
muscular dystrophy and improper healing of wounded tendons (23), (24), (25).
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In addition to the molecules that were briefly considered above, one of the most important and
certainly the most abundant ECM protein of mammals are the collagens (26).
1.2.2 Types of Collagen Collagen is a high abundance, ubiquitously expressed protein in mammalian connective tissues
(27). A large proportion (~80% - 90%) of the ECM in many soft connective tissues of mammals
is composed of type I fibrillar collagen (28). In particular, there are at least 28 known members
of the collagen family of proteins (29, 30). All collagens share a common structural α helix
motif. Collagens are rich in glycine: every third amino acid residue is glycine, which is essential
for the structure and function of the triple helical structure. A collagen molecule is ~300 nm in
length, is organized as a triple helix (two a1 chains and one a2 chain) and typically consists of
~1000 amino acid residues in a repeating Gly-X-Y triplet, where X- and Y- are commonly
proline and hydroxyproline (3), (5), (31), (32). Glycine, the smallest amino acid, is specifically
positioned at every third residue, enabling orientation of its R group (a hydrogen atom) to be
positioned towards the centre of the helix, which enables tight helical packaging.
Collagens can be subdivided into distinct groups based on their structure. Types I, II, III,
V, and XI make up the “fibrillar collagen” group (5, 30, 33) while the non-fibrillar group of
collagen is composed of several sub-categories. FACIT (Fibril Associated Collagens with
Interrupted Triple Helices) collagens include types IX, XII, XIV, XVI, XIX, XX, and XXI.
“Short Chain” collagens include collagens type VIII and X (34). Type IV collagen is considered
a “network-forming” collagen and is an important component of the basement membrane (3),
(5), (35). Type VII collagen, another network-forming collagen, is especially abundant in the
skin, oral mucosa, and in epidermal-dermal junctions (5), (36) and assembles into dimers to form
“anchoring fibrils”. These are particularly important in attaching the basal laminae to the
underlying connective tissue. There are also transmembrane collagens (Types XIII, XV, XVII,
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XXIII, and XXV), which form unique structures (34), (37). While the primary function of most
collagens is to act as the structural support for connective tissues, collagens also act as binding
partners for other ECM proteins like fibronectin (29), (38).
1.3 Structure and function of Type I fibrillar collagen Type I fibrillar collagen is widely expressed in connective tissues throughout the body. The
organization of collagen molecules into fibrils, and then hierarchically of fibrils into fibers, is a
critically important process for vertebrate organisms. At the organizational level of the fibril,
collagen is comprised of parallel arrays of collagen molecules. The synthesis of collagen
molecules, their assembly into collagen fibrils and fibers, and the remodeling of collagen (which
involves synthesis, degradation and fibrillar reorganization), are crucial for embryonic
development, maintenance of tissue architecture, and preservation of organ function (39), (40).
Conversely, inappropriate regulation of collagen assembly and structure is linked to a large array
of human diseases (41).
1.3.1 Structure Type I fibrillar collagen, denoted “collagen I” henceforth, is found primarily in skin, tendon,
ligaments, cornea, and bone. There are at least 45 different genes that code for the various α
chains which make up the different types of collagen. These are designated with Roman
numerals in vertebrates in the same order as their discovery (e.g. Type I, II, XXV, etc.) (9).
Arabic numerals distinguish each α chain, which are encoded by a specific and distinct gene (e.g.
type II fibrillar collagen is a homotrimer composed of three identical α1 (II) chains encoded by
the COL2A1 gene in humans).
At the molecular level, collagen I is a heterotrimer composed of two α1 (I) chains and a
single α2 (I) chain wound in a tight triple helix around a central axis. This macromolecular
structure contains long uninterrupted triple helical domains that form a rod. The triple helix is
7
comprised of three left-handed chains that are arranged as a right-handed helical structure with a
single residue stagger between adjacent chains (34). Collagen I is composed of both collagenous
(COL) and non-collagenous (NC) domains; NC domains participate in both structural assembly
and conferring biological activity. Glycine is located at every third residue in the triple helix,,
which is stabilized by electrostatic interactions between lysine and aspartate residues, hydrogen
bonding between individual chains, and by a high content of proline and 4-hydroxyproline
residues (9), (34), (42). Individual collagen I molecules are configured as rods that associate to
form supramolecular structures organized into collagen fibrils, which when assembled then
constitute the next hierarchical level, collagen fibers (43). Collagen fibrillogenesis involves the
intracellular formation of collagen molecules that are targeted to plasma membrane protrusions
(“fibripositors”) via Golgi-to-membrane carriers (44), (45), (46). In the extracellular
compartment, intermediate fibrils fuse to enable maturation of fibrils. Subsequently, collagen
molecules undergo post-translational modifications that involve hydroxylation of proline and
lysine residues, glycosylation, and cleavage of pro-peptides that are used for initial alignment
and supramolecular organization of collagen fibrils (40). The self-assembly of procollagen
polypeptides into fibers is classically seen in electron micrographs as electron dense, cross-
banded linear structures that exhibit 67 nm spacing in tissue sections of appropriate thickness
(i.e. optimally, sections of gold interference colour)(Diagram 1 below).
1.3.2 Function Early X-ray crystallography studies of mammalian connective tissues provided insights into the
structure of type I collagen in stretched tendons (47). Later work built upon this foundation and
showed the structural role of type I collagen and its contributions to tissue architecture and the
mechanical properties of tissues, including the properties of tensile strength (in skin) and traction
force resistance in ligaments (34), (48). As an essential component of the extracellular matrix,
8
collagen I provides important scaffolding functions upon which specialized cells of
mesenchymal origin such as chondrocytes, fibroblasts and osteoblasts form the tissues that
comprise different types of connective tissues. The principal fibrillar component of tendon and
skin is type I collagen, which is a reflection of its high tensile strength, a property that enables
many tissues to undergo stretching and compression yet maintain their structural integrity. At the
microscopic level, type I collagen fibrils provide conduits for cell migration, which can be
beneficial in wound healing but which can also be detrimental in the context of cancer cell
invasion and metastasis. In the formation of bone, type I collagen fibrils are important for the
assembly of the matrix upon which hydroxyapatite crystals are deposited and are ultimately
mineralized, thereby enabling the formation of mineralized bone (49). The importance of
collagen in bone formation is well-demonstrated in osteogenesis imperfecta, a genetic disorder
characterized by excessively brittle bones. Osteogenesis imperfecta arises in humans carrying
deleterious mutations in the genes that encode the α1 (I) and α2 (I) chains of type I collagen. In
this instance, the abundance and proper folding of type I collagen triple helices is impaired,
which ultimately weakens the structure of bone (49).
More recent work has demonstrated the importance of cell-collagen interactions in
mechanically-mediated maintenance of tissue structure and function. Specialized connective
tissue cells embedded within the matrix such as fibroblasts, form extensions, which adhere to and
exert cell-generated mechanical forces on collagen fibers in the ECM. Fibroblasts express
adhesion receptors to fibrillar collagen, which include the α2β1 and α11β1 integrins and in some
instances, the Discoidin Domain Receptors (DDRs) 1 and 2. These molecules may play a central
role in detecting local deformations in the structure of fibrillar collagen as under certain
circumstances, these adhesion receptors can “translate” specific structural features of collagen
9
fibrils into intracellular signals. DDR1, a receptor tyrosine kinase that is activated by binding to
fibrillar collagen, is overexpressed in certain forms of invasive ductal breast cancers where it
plays a role in promoting mechanical remodeling of the surrounding collagenous matrix,
possibly providing a positive feedback mechanism for cancer cell metastasis (6), (7), (50), (51) .
Homeostatic regulation of collagen 2.1 Synthesis Collagen biosynthesis is a tightly regulated process composed of multiple complex steps. As
described above, collagen I is a triple helical molecule made up of two α1(I) chains and one α2(I)
chain. In humans, the COL1A1 gene is located on chromosome 17 at position 21.33 while
COL1A2 is found on chromosome 7 at position 21.3 (48). In the human genome, ~50 exons
encode the nascent mRNA transcript from which the individual pro α-chains are synthesized.
The pro α chains enter the lumen of the rough endoplasmic reticulum (RER), in part guided by
specific N- and C-terminal amino acid sequences and an N-terminal signal peptide. After entry
into the RER lumen, selective prolyl and lysyl hydroxylation take place and the signal peptide is
cleaved off. These initial post-translational modifications are essential for stabilization of the
procollagen molecule and facilitate the formation of disulphide bridges between the C-terminal
domains of the individual α1(I) and α2 (I) chains. The collagen I triple helix undergoes assembly,
a process that proceeds from the C-terminal domain to the N-terminal domain. In the Golgi
network, packaging of the collagen molecule occurs from where it is secreted into the
extracellular space. There, following collagen molecule alignment, C- and N-propeptides are
cleaved. In the extracellular space collagen molecules self-assemble to form fibrils with
characteristic staggered pattern. This staggered assembly of collagen molecules is responsible for
the 67 nm banding pattern that is seen in electron micrographs and is reflection of the separation
10
of each molecule by 35-40 nm. This arrangement also facilitates strong interactions with adjacent
molecules and accounts for the high tensile strength that characterizes collagen fibers. Inter-
fibrillar covalent cross-links at lysine and hydroxylysine residues stabilize collagen fibrils. The
telopeptide regions of collagen molecules are rich in hydroxylysine and lysine residues; lysyl
oxidase is responsible for their conversion into their aldehyde counterparts (hydroxyallysine and
allysine), which either spontaneously cross-link with one another or other lysine and
hydroxylysine residues on adjacent molecules to form immature crosslinks. These immature
cross-links then mature by forming additional bonds with adjacent amino acid residues (34),
(48), (52)(Diagram 1 below).
Diagram 1. Formation of the supramolecularstructure of type I fibrillar collagen
2.2 Collagen Remodeling Three major mechanisms enable remodeling of collagen fibrils. One mechanism involves
secretion of degradative enzymes such as matrix metalloproteinases that digest the polymers
which comprise the collagenous ECM. The second mechanism involves a cell-mediated process
by which collagen fibrils are partially cleaved on the cell surface, internalized by phagocytosis
The structure and hierarchical assembly of type I collagen ranging from the polypeptide chain to a fully assembled collagen fibril
11
and degraded in phagolysosomes. The third mechanism is an actin-myosin-mediated process by
which cells exert mechanical forces on collagen fibrils to increase the local density and
organization of collagen fibrils.
2.2.1 Collagen Proteolysis Three separate pathways have been described for enzymatically-mediated degradation of ECM
in soft connective tissues: the matrix metalloproteinase (MMP) pathway, the plasmin pathway
and the serine protease pathway (53). Serine proteases are secreted by polymorphonuclear
leukocytes and include molecules such as elastase and cathepsin G, which mediate degradation
of the ECM at a neutral pH in the inflammatory response. Serine proteases can destroy bacteria
in lysosomes (i.e. intracellularly) but can also digest ECM proteins at extracellular sites to clear
damaged tissue or facilitate entry of leukocytes into dense connective tissues. Several different
ECM proteins (e.g. fibronectin, laminin, proteoglycans, collagens) can be cleaved by these
hydrolytic enzymes (54) both at acidic pH (which is more optimal for lysosomal hydrolase
cleavage of substrates) or at juxta-membrane sites where the pH is much closer to pH=7.4,
although catalytic rates are substantially reduced at pH=7.4.
Plasmin is an enzyme with broad substrate specificity that plays a role in matrix
remodeling by virtue of its ability to rapidly cleave peptide bonds between lysine and arginine
residues in ECM molecules like fibrin and fibronectin (53). Currently, there is no evidence that
plasmin can digest type I or type II collagen (abundant in cartilage) but previous work has shown
that plasmins can influence collagen degradation by activating certain types of matrix
metalloproteinases (MMPs) with triple helicase activity (55).
ECM proteolysis is also mediated by a family of zinc-dependent endopetidases (MMPs)
that comprise at least 24 known members. Three of the MMP family, MMP-1, MMP-8, and
MMP-13 are interstitial collagenases that can cleave and degrade triple helical collagen fibrils.
12
Cleavage by these MMPs at specific sites in the collagen molecule generates one quarter and
three quarter length fragments of collagen. These MMPs are often bound to inhibitors (e.g. tissue
inhibitors of metalloproteinases; TIMPs) and are also usually present in tissues as latent
enzymes. As a result of complex and well-described activation processes that are mediated by
other MMPs or certain serine proteases, the catalytic activity of the interstitial collagenases can
be greatly increased. Further, and as touched upon above, the catalytic activity of certain MMPs
is strongly affected by TIMPs and by certain membrane-bound adhesion receptors including β1
integrins (28), (56).
Initial digestion of the collagen triple helix promotes denaturation of the fibrils and
allows further fragmentation by gelatinases at a lower temperature (i.e. <37°C). This step-wise
process allows the normally insoluble collagen molecule to be converted into gelatin. This
degradation process also as indicated above, in turn allows gelatinases like MMP-2 and MMP-9
to further digest the collagen molecule (57). Collectively, these MMP-dependent degradative
processes not only mediate breakdown of the ECM but may also expose cryptic binding sites in
collagen fibrils to facilitate cell attachment by collagen receptors. This exposure in turn controls
cell behavior, such as the promotion of epithelial cell migration in wound healing and cancer
(58), (59).
2.2.2 Phagocytosis Collagen phagocytosis is a cell-mediated, multi-step process that involves cell-dependent
recognition and binding of collagen fibrils, activation of collagen receptors upon binding,
collagen fibrillar internalization and intracellular lysosomal degradation. Collagen fibrils in the
ECM are degraded and internalized by fibroblasts using this phagocytic route, particularly in soft
connective tissues with rapid rates of collagen turnover such as the periodontal ligament (60). At
early stages of collagen phagocytosis, specific amino acid sequence motifs within the collagen
13
molecule are recognized by collagen-binding adhesion receptors, which promote the initial steps
of internalization. Integrins are one family of collagen-binding receptors involved in
phagocytosis; binding promotes local clustering and recruitment of large numbers of receptors
and intracellular molecules like actin filaments, microtubules, and actin-binding proteins to
enable formation of large, multi-protein adhesion complexes (focal adhesions) (61). Protein
clustering enables bi-directional communication between the ECM and the actin cytoskeleton,
which initiates signaling processes that can influence several different cellular processes and the
further recruitment of other types of accessory proteins (62, 63), (64). After fragmentation on the
cell surface, which can be mediated by membrane-type MMPs such as MT-1, the collagen fibrils
are engulfed and internalized by lamellipodial types of structures (65). Subsequently, collagen
fragments enter into lysosomes where lysosomal hydrolases such as cathepsin L, B and N further
digest the fibrils (66).
2.2.3 Traction In addition to the aforementioned proteolytic and phagocytic processes that mediate collagen
degradation there is a third, mechanically-mediated, but not yet fully defined process, that
contributes to collagen remodeling. Remodeling by traction is a process by which cells,
embedded within the ECM, exert cytoskeletally-generated contractile forces on fibrillar matrix
biopolymers like fibronectin and collagen (51). Notably, mechanosensation and metastatic
invasion by cancer cells involve critical, physical interactions of cells with matrix proteins (67),
which mirror several important steps that are also seen in tractional remodeling.
In general, the initial step in tractional remodeling involves binding of fibrillar ECM
proteins by cell adhesion receptors. Specialized receptors can selectively bind collagen fibers by
recognizing specific amino acid motifs embedded in the triple helix. Upon binding collagen, the
adhesion receptors frequently undergo allosteric changes that initiate intracellular signaling
14
processes, and which are associated with receptor clustering, the recruitment of accessory
proteins (e.g. tyrosine kinases such as focal adhesion kinases and phosphatases such as protein
tyrosine phosphatase-a), and cytoskeletal contractile proteins (e.g. non-muscle myosin IIA) to
form large adhesion complexes. While the development of strong adhesions to fibrillar ECM
proteins is essential for cell anchorage in many connective tissues, this process also enables cells
to focus cell-generated mechanical forces on collagen fibers, which thereby enables
reorganization of fibers and their structure and orientation in the extracellular space.
Fibrillar collagen receptors 3.1 β1 Integrin Cell-mediated tractional remodeling of the ECM can increase the mechanical strength of tissues
by, for example, optimization of fiber orientation, a process that ultimately impacts the growth,
survival, and migration of cells in connective tissues (68), (69). There are at least three, critically
important, membrane-bound collagen receptors that contribute to mechanical remodeling of
fibrillar collagen. The first two adhesion receptors, the α2β1 and α11β1 integrins, are fibrillar
collagen-binding, heterodimeric receptors that belong to the integrin family of plasma
membrane-associated proteins (70), (71). Both of these integrins have been implicated in
mesenchymal cell migration, collagen fiber reorganization, collagen gel contraction, and tissue
fibrosis and wound healing processes modeled in vitro (71), (72), (73).
3.1.1 Structure and isoforms of collagen-binding integrins Integrins encompass a family of heterodimeric plasma membrane-associated receptors that
contribute to the adhesion of cells to ECM proteins and to other cells. Integrins consist of two,
non-covalently bound α and β subunits; so far 18 α and 8 β subunits have been discovered.
Globally, there are a total of 24 different combinations of α and β subunits that comprise specific
15
integrins with varying functions and ECM binding specificities (Diagram 2 below) (74). At least
12 α subunits can combine with the β1 subunit. Each of these integrin pairs has a generally non-
redundant function and a measureable ligand binding specificity (29, 71-73, 75). The β subunit is
characterized by a large extracellular domain, a single-pass transmembrane domain, and a short
cytoplasmic tail (76). Association of the N-termini of the α and β subunits enables ligand-
specific ECM binding. Intracellularly, β integrin subunits interact via their C-terminal domains
with the actin cytoskeleton through a large array of actin binding proteins that include filamins,
vinculin, talin, paxillin and a-actinin.
The maintenance of tissue homeostasis is critical in wound healing and developmental
processes, both of which involve important functions fulfilled by collagen-binding integrins (72),
(77) and which are regulated in part by the affinity of integrins for their cognate ligands. In this
context, the binding activity of integrin receptors is modulated by allosteric alterations of integrin
structure, which in turn are regulated by local conditions in the cellular environment and by the
clustering, organization and spatial orientation of integrins in the plasma membrane. Certain
elements of integrin functional activity (e.g. ligand binding, ligand specificity and structural
Diagram 2. The β1 integrin family of extracellular matrix receptors.
The β1 integrin subunit can bind to a variety of a subunits giving rise to a combination of integrin receptors capable of binding to specific
extracellular matrix proteins. Courtesy of (74).
16
stabilization) can be promoted or suppressed by the relative abundance of Ca2+ and Mg2+. These
ions can impair integrin affinity for ligands (i.e. Ca2+) or increase affinity and promote cell
adhesion (i.e. Mg2+) (78), (79). When inactive, integrins are not typically bound by ligands and
exhibit a resting conformational state. Ligand binding induces conformational (allosteric)
changes, which enhance ligand binding and also can promote or inhibit the generation of certain,
downstream, adhesion-dependent signals.
3.1.2 Function in collagen remodeling The α2β1 and α11β1 integrins are the main β1 integrin receptors involved in binding to, and
regulating the structural organization of, type I fibrillar collagen. Other integrins interact with
different collagen types and with other ECM proteins including fibronectin, osteopontin and
laminin (80). Previously, a functional link between integrin expression and disease was observed
that involved the α1β1 and α3β1 integrins in osteoarthritic cartilage in which receptor
overexpression was found on the surface of hypertrophic chondrocytes (81). Stimulation of the
α5β1integrin by presentation with fibronectin fragments, that modeled exposure to damaged
matrix in osteoarthritis, enhanced the production of PGE2 reactive oxygen species, nitric oxide,
and certain MMPs including MMP-1, MMP-3 and MMP-10 (80). Fibronectin fragments also
induce “crosstalk” between α5β1 and α4β1 integrins that promote osteoblast differentiation (82).
The importance of collagen binding integrins in development is readily seen in developing mice:
β1 integrin knockout mice die at a very early age of development. Further, deletion of the β1
integrin in particular affects chondrocytes, which exhibit defective proliferation, migration and
failure to form normal cartilage and bone (83), (84). Keratinocyte migration over connective
tissues and wound re-epithelialization are facilitated by MMP-1-mediated collagen cleavage after
binding to the α2β1 integrin (56), (85), indicating that this integrin is important for matrix and
17
epithelial function. In three dimensional collagen gel contraction assays, α2β1 integrin
expression is directly related to increased gel contraction and the transmission of contractile
forces, which is impaired by treatment with α2 integrin blocking antibodies (86). This binding
mechanism is also relevant for collagen remodeling by phagocytosis as activation of the β1
integrin is important for the initial binding step of collagen phagocytosis (62). β1 integrin-
containing adhesions are also involved in the modulation of the function of other adhesion
receptors and may interact with other receptor families (e.g. Discoidin Domain Receptors;
DDR). Indeed, overexpression of DDR1 enhances the abundance and activation of β1 integrin at
the cell surface. Conversely, β1 integrin-mediated binding to collagen is inhibited by knockdown
of DDR1 expression (87).
3.2 Discoidin Domain Receptors (DDRs) 3.2.1 Localization, expression and structure A more recently discovered family of receptors that helps to maintain the structure and function
of fibrillar collagens are the discoidin domain receptors (DDRs), a unique sub-family of receptor
tyrosine kinases (RTKs) whose initial activation is mediated by binding to fibrillar collagen (i.e.
native, triple-helical form). DDR1 and DDR2 have been identified in vertebrates whereas
homologues of DDRs have been discovered in certain invertebrate organisms (6, 88). DDR1 in
particular is activated by binding to collagens type I-VI and VIII (89). DDR1 and DDR2 exhibit
unique characteristics that differentiate them from classical RTKs. Whereas classical RTKs
exhibit rapid activation (i.e. within seconds of ligand binding), DDR activation (measured by
tyrosine autophosphorylation) (6) peaks at ~90 minutes after initial ligand binding and is
sustained for up to 18 hours (and even several days) after stimulation (2), (90).
18
DDR1 and -2 are transmembrane proteins with two extracellular “discoidin” domains
composed of 160 amino acid residues. These are unusual structures that have not been identified
in other RTKs. DDRs are comprised of a juxtamembrane region and a C-terminal, cytoplasmic
region, which is responsible for initiating intracellular signaling cascades via tyrosine
phosphorylation. The name “discoidin” arises from the observation that DDRs share homology
with the protein discoidin I (found in Dictyostelium discoideum) where it functions as a
galactose-binding lectin (91). Early work using recombinant DDR1 showed that sites of ligand
binding and receptor activation were located at loops 1 and 3 of the extracellular domain while
other studies showed that the large juxtamembrane regions of DDRs may play an auto-inhibitory
role, which contributes to their unusually prolonged activation kinetics (2, 92), (93).
The DDR1 gene, which is composed of 17 exons, is located on human chromosome 6.
DDR1 protein is found at high levels in epithelial cells while there is abundant DDR1 mRNA
expression in spleen, kidney, lung, brain and placenta of mice and humans (94), (95), (96), (97).
There are five different isoforms of DDR1, DDR1a – e (Diagram 3 below) (98), (99), which are
formed via alternative splicing (7). DDR2, which is located on human chromosome one, is
expressed as only one known isoform, which is composed of 19 exons. DDR2 protein is
predominantly found in connective tissue cells that originate from embryonic mesoderm. DDR2
mRNA is highly expressed in a broader array of tissues including heart and skeletal muscle and
in brain, kidney and lung (6), (100), (101), (102). For DDR1, DDR1a and b are the most
commonly expressed isoforms. DDR1c is the longest isoform. DDR1d is missing the entire
intracellular kinase domain. DDR1e is a “kinase dead” isoform that lacks entire sections of the
juxtamembrane region and the ATP binding site (6), (103), (104).
19
DDRs exhibit unique characteristics that are not exhibited by most RTKs. The
extracellular region is uniquely composed of two tightly-linked globular domains: an N-terminal
Discoidin (DS) domain and a Discoidin-like (DS-like) domain. Next is a transmembrane (TM)
region, which is thought to be the key region for the formation of DDR dimers (105), followed
by several large cytoplasmic juxtamembrane (JM) regions. Finally DDRs contain catalytic kinase
domains that are followed by short C-terminal tails (not present in DDR1d) (6).
DDR1 is thought to exist as a stable pre-formed dimer at the cell surface; upon binding to
triple-helical collagen the structural conformation of DDR1 changes triggering intracellular
signaling cascades (106). At the present date there is some contention over the DDR activation
kinetics and more work needs to be done. Some work has shown that dimerization of DDR1
ectodomains was essential for collagen binding as DDR1 dimers originating in the biosynthetic
pathway were stable at the cell surface without the presence of collagen (7), (107), (108).
Conversely, other experimental evidence suggested that collagen-induced DDR1 clustering was
not essential for its activation as even short triple-helical collagen peptides were sufficient for
DDR1 activation (109).
Diagram 3. Structure and isoforms of DDR1 and DDR2
DDR1 has five isoforms formed via alternative splicing while DDR2 has only one. The different isoforms are distinguished by length and size. DDR1a and DDR1b are the
most widely expressed while DDR1d and DDR1e are non-functional. Courtesy of (99).
20
There is some functional interplay between DDR1 and the classic fibrillar collagen
adhesion receptor α2β1 integrin. DDR1, which binds a GVMGFO motif in collagens I-III,
promotes ECM degradation and increases collagen turnover in a β1 integrin-dependent manner
by indirectly stimulating MMP activation. This functional interplay does not apparently rely on
close proximity of DDRs to integrins since DDR1 does not co-localize with classical focal
adhesions containing β1 integrin, talin and vinculin (51), (87), (110).
3.2.2 DDR function in fibrotic disorders Fibrosis occurs when the wound healing response to injury or infection fails to restore the
original tissue architecture and function. Dysregulated collagen remodeling, attributable to
mechanically-induced collagen compaction or the excessive deposition of collagen, is a critical
determinant of tissue fibrosis. Tissues initiate wound healing responses in order to re-establish
homeostasis after an injury but, if the damage is severe or persists chronically, then the
deposition of collagen (and other matrix proteins) exceeds degradation and the tissue repair
process is manifest as fibrosis. On a larger scale, fibrosis plays a negative role by impairing
organ function, especially in several high prevalence diseases (e.g. cardiovascular disease,
pulmonary fibrosis and chronic renal diseases) (111-113).
Although the mechanism is not well-defined, inflammation and collagen I deposition
have been associated with higher levels of DDR1 expression (114, 115). Further, collagen I-
induced DDR1 tyrosine phosphorylation can activate p38 and ERK to promote collagen
synthesis (116). Activation of DDR1 has also been linked with pulmonary fibrosis after
pulmonary injury induced by bleomycin (117). As a regulator of collagen degradation, DDR1
has been functionally linked with MMP-2 and MMP-9 (118) in mouse vascular smooth muscle
cells (VSMCs) and with MMP-1 in human VSMCs (119). DDR1 overexpression has been
21
implicated in the fibrosis seen in several types of cancers as measured by increased collagen
compaction (87) and alignment (51), (120). In cells adhering to, and migrating on collagen,
DDR1 is associated with non-muscle myosin IIA (NMIIA) (121), which established a link with
an earlier finding showing that myosin is required for fibroblast-mediated tractional remodeling
of collagen fibers (122). DDR1 expression and activation are also important for the collagen-
dependent assembly of NMIIA heavy chain into filaments (51) (121).
Cytoskeleton cell-generated tractional remodeling 4.1 Actin Actin is one of the most abundant and highly conserved proteins in eukaryotic cells. The
dynamic regulation of the assembly of actin monomers into filaments is essential for many
cellular processes including transit through the cell cycle, cell attachment to ECM, preservation
of cell structure, mechanotransduction, matrix remodeling and cancer (123), (124), (125), (126).
4.1.1 Structure, isoforms and function Actin is a ~42 kDa protein, which exists in two states that are in constant flux: globular actin
monomers (G-actin) and actin filaments (F-actin). The actin monomer comprises ~375 amino
acids and is usually bound to ATP or ADP. Three small differences in the N-terminal amino acid
sequences of actins of vertebrates are the basis for three broadly expressed actin isoforms. α-
actin is found primarily in muscle cells (α-skeletal, α-cardiac and α-vascular) but also in
specialized types of fibroblasts (α-smooth muscle actin in myofibroblasts). b- and g-Actin (g-
non-muscle and g-smooth muscle) are primarily found in non-muscle cells (127). The actin
filament is an 8 nm-wide structure, which assembles to form a right-handed helix (3). The actin
filament is asymmetrical, due to its assembly from actin monomers, and has structurally distinct
ends: the slower-growing minus (-) (pointed) end and the faster-growing plus (+) barbed end (3).
22
Spontaneous assembly of actin monomers into filaments, particularly at the pointed end, occurs
but contributes only minimally to actin assembly. There are three phases of actin polymerization
that are dependent on ATP hydrolysis: initial monomer addition (e.g. by nucleation), elongation
and stabilization. For actin assembly to occur, multiple subunit-subunit actin contacts must be
formed, which then quickly allow polymer elongation to occur preferentially at the (+) end to
form actin filaments. One of the most common processes for actin filament assembly involves
addition of actin monomers to the (+) end of existing actin filaments, which requires the
exposure of the (+) end by removal of actin capping proteins (e.g. gelsolin). Actin monomers are
then rapidly and preferentially added to the (+) end (the elongation step) until the rate at which
the addition of subunits at the (+) end is in equilibrium with the removal of subunits at the (-)
end, which results in a steady-state (the critical concentration) (3), (128). There are other
important mechanisms by which actin filaments are assembled which involve addition of
monomers on to filaments at 70° branch points by the actin related protein 2/3 (Arp 2/3) complex
and also by a number of nucleation mechanisms mediated by proteins including formins, which
begin with the addition of monomers to a short actin oligomer as the nucleation primer.
Actin is an indispensable protein for cell function; it plays a central role in cytoskeletal
structure and mediates a large number of processes including contraction and motility. In yeast,
clathrin-mediated endocytosis, which is involved in the uptake of receptor proteins and
extracellular fluids, is mediated by dense networks of actin, the assembly of which is controlled
by the Arp2/3 complex (129). Lamellipodia, which are composed of dense, crosslinked networks
of actin filaments, are protrusive cell extensions found at the leading edge of a motile cell. The
polymerization of actin at the leading edge of lamellipodia can generate forces that are exerted
on the ECM (130). Within lamellipodia, cell adhesions connect to non-muscle myosin-IIA and
23
B, which generate forces that are transmitted to ECM proteins (131). Actin networks are
organized in some cell types (e.g. cultured fibroblasts) into contractile actin bundles that
typically insert into adhesions located on the ventral surface of cultured cells (132). These actin-
rich structures (stress fibers) are composed of repeats of myosin-II motor proteins and α-Actinin
(130) and which enable cytoskeletally-generated contractile forces to be transmitted to the ECM
substrate.
4.1.2 Actin-binding proteins As the central element of the actin cytoskeleton, actin filaments play crucial roles in multiple
cellular processes. The dynamic nature by which actin is assembled into filaments and other,
higher order structures is dependent on the interaction of actin with a large family of proteins that
bind actin and regulate its form and function. Some of these actin binding proteins include
filamin, gelsolin, cofilin and thymosin (3), all of which contribute to actin dynamics and
structural regulation. Accordingly, the abundance and activity of these actin binding proteins
influence many cellular processes including motility, phagocytosis, matrix remodeling and
contractility (124), (125), (126). For example, gelsolin is an actin capping/severing-protein that
co-localizes with non-muscle myosin IIA at sites of cell adhesion to collagen to promote
collagen phagocytosis (133). Fascin is an actin bundling protein that is overexpressed in invasive
human melanoma A375MM cells and contributes to ECM invasion by stabilizing actin filaments
in invadopodia (134).
One of the most important actin binding proteins is myosin (found in muscle cells and the
related non-muscle myosin in cells like fibroblasts). Myosin family proteins are actin-dependent
motor proteins that generate contractile forces as a result of force generated by their motor
domains and by interactions through their N-termini with a large number of molecules that
24
include actin, integrins and DDRs. These protein interactions enable myosins to act as
connectors that bridge the actin cytoskeleton and plasma membrane-associated ECM adhesion
receptors and to apply contractile forces like tension to these adhesions.
4.2 Non-muscle myosin II In muscle cells, myosin II forms highly ordered structures with actin to generate contractile
forces that are regulated by Ca2+ and ATP hydrolysis (3). Non-muscle cells contain non-muscle
myosin II (NMII) which also generates contractile forces but mediates contraction by a variety of
means.
4.2.1 Structure, isoforms and function NMII is a two-headed class II myosin that is ubiquitously expressed by metazoans (pictured
below). NMII is a hexamer, composed of two N-terminal heavy chains that form a homodimer
made of globular heads (motor domains) that contain binding sites for other proteins like actin
and for molecules like ATP that are critical for motor activity. There are two essential light
chains (ELCs) and two regulatory light chains (RLCs), which contain tyrosine and serine
residues that upon phosphorylation modulate the contractile activity of NMII (135). There are
two major heavy chain isoforms of NMII in vertebrates, NMIIA (myh9) and NMIIB (myh10); a
third NMIIC (myh14) has also been found in mammalian cells (136), (137) but will not be
discussed here. Most of the differences between NMIIA and NMIIB are located in the non-
helical tail whereas the motor domain and rod region exhibit 85% and 72% amino acid sequence
identity respectively (135). There are diverse patterns of localization for both isoforms and many
cell types express NMIIA and NMIIB (e.g. fibroblasts) while some types of cells exhibit only
one isoform. For example, NMIIA is expressed in chicken intestinal epithelium (138) while
NMIIB is expressed in embryonic cardiomyocytes (139). While the expression patterns of
25
NMIIA and NMIIB are often diverse, their functions appear to be highly specific. During wound
healing when endothelial cells start to migrate into the wound, NMIIA is abundant at the leading
edge of cells and contributes to membrane protrusion whereas NMIIB is predominantly located
at the trailing edge of cells to regulate retraction (140). As an abundant protein in cell adhesion
complexes, NMII can also bind a wide variety of proteins to modulate its own contractile activity
and to transduce mechanical stimuli into biochemical signals (mechanotransduction), a process
which can influence ECM remodeling (121) and gene expression (141).
4.2.2 Mechanisms of activation The main mechanisms by which contractile activity and filament assembly of NMIIA is
regulated involves the phosphorylation and dephosphorylation of specific amino acid residues by
several different kinases and phosphatases. The regulatory light chains and heavy chains of NMII
can be phosphorylated, which influences contractile activity (51), secretion (142) and cytokinesis
(143) and several processes that are linked to disease states (144) (145) (146). Myosin light chain
kinase (MLCK) phosphorylates Ser19 (S19) of the regulatory light chain. As a result of S19
phosphorylation, NMIIA is stimulated to re-fold and form filaments, an important step in force-
generating contractile activity (135). Knockdown of MLCK expression in cultured cells induces
cell rounding and decreased proliferation (147). The Rho associated coiled-coil kinase 1
(ROCK1) regulates NMII activity through phosphorylation of the regulatory light chain. ROCK1
acts downstream of RhoA (a small GTPase) to induce NMII stress fiber assembly in cultured
cells and contraction by phosphorylating the regulatory light chain at S19. This activity of
ROCK1 is two-fold as ROCK1 also phosphorylates the myosin binding subunit of myosin
phosphatase in an inhibitory manner, thereby blocking dephosphorylation of the regulatory light
chain at S19.
26
More recent data have given insight into the orchestration of phosphorylation and
dephosphorylation by identifying novel accessory proteins that indirectly modulate
phosphorylation of the regulatory light chain. Myosin phosphatase Rho-interacting protein
(MRIP) is a 116 kDa signaling molecule that binds RhoA/ROCK1 in an inhibitory manner (148)
and targets myosin phosphatase to the regulatory light chain (149) in order to simultaneously and
indirectly counteract RhoA-induced NMII contractile activity. As a binding partner of NMII that
regulates phosphorylation (148), MRIP may play a central role in the NMIIA-associated
adhesion complex which assembles upon cell binding to ECM proteins like collagen.
Diagram 4. The structure of NMII and RLC kinases
A single NMII fully-assembled protein is pictured here. NMII is a hexamer, the regulatory light chain (pictured in light blue) can be phosphorylated by several different kinases at specific amino acid
residues to regulate protein contraction and relaxation. Threonine 18 and Serine 19 are phosphorylated by ROCK to induce contraction. Courtesy of (144).
27
Statement of the problem Collagen is a high abundance, ubiquitously-expressed protein of mammals that undergoes
extensive remodeling to preserve tissue homeostasis, a process that is mediated by several types
of cell adhesion receptors. Upon binding to fibrillar collagen, a family of receptor tyrosine
kinases (Discoidin Domain Receptors, DDR) is uniquely activated via autophosphorylation and
oligomerization. DDR1 expression is strongly upregulated in aberrant wound healing and in
certain forms of invasive cancer.
In addition to the synthesis and degradation of collagen fibers, appropriate maintenance
of extracellular matrix structure involves remodeling of fibrillar proteins by cell-generated
contractile forces. Non-muscle myosin IIA (NMIIA) is an actin-binding motor protein that
generates contractile forces and binds DDR1. The contractile function of NMIIA is tightly
regulated by several different signaling molecules that determine its relative phosphorylation
state. Currently we do not fully understand the relationship between NMIIA and the molecules
that regulate its relative phosphorylation state and activation, which lead to generation of
contractile forces and that mediate DDR1-dependent collagen remodeling.
Hypothesis:
Myosin Phosphatase Rho-interacting protein (MRIP) regulates NMIIA contractile activity to
affect DDR1-dependent collagen remodeling.
Objectives:
1) Use tandem mass spectrometry of DDR1 immunoprecipitates to identify potential
regulatory proteins that are involved in DDR1-dependent collagen adhesion
2) Assess the spatial localizations and interactions of MRIP with NMIIA, and DDR1
3) Identify the effect of MRIP on the spatial association of DDR1 with NMIIA
4) Quantify the impact of MRIP, NMIIA and DDR1 expression and activity on the regulation
of cell-to-collagen adhesion, collagen contraction, and remodeling.
28
Chapter 2 Introduction
Extracellular matrix (ECM) collagen is the most abundant protein in vertebrates (26) and
undergoes continuous remodeling to enable tissue homeostasis, an important determinant of
health and disease in several different organ systems (69). Collagen remodeling requires tightly
regulated control of biosynthetic and degradative processes including metalloproteinase-
mediated pericellular proteolysis (55) and intracellular, lysosome-mediated degradation of
collagen fibrils as occurs in phagocytosis (60). A third remodeling system involves cell-
generated mechanical forces that affect collagen fiber orientation and regulates the density of
collagen fibers in soft connective tissues (120). In chronic inflammatory lesions, excess collagen
deposition, dysregulated wound healing responses and fiber compaction mediated by cell-
generated tractional forces contribute to tissue stiffening, a hallmark of fibrotic disease and
certain invasive epithelial tumors (7), (8). Tractional remodeling of collagen requires tight
adhesion of cells to fibers, which is mediated by membrane-bound receptors that enable the
application of cell-generated mechanical forces. While the mechanism of force generation by the
actomyosin contractile system is well-described in muscle cells (150), (68), the regulation of
force generation in fibroblasts to mediate collagen tractional remodeling is not well-defined.
The integrins α2β1, α11β1 and the discoidin domain receptors (DDR 1 and 2) are widely
expressed fibrillar collagen adhesion receptors. While much is known about the interaction of β1
integrin adhesions with fibrillar collagen in tractional remodeling (72), there is little definitive
information on the role of DDRs, which are unique receptor tyrosine kinases (RTK). DDRs are
characterized by their unusual activation mechanism, which involves collagen binding-induced
autophosphorylation, a slow process that occurs over the course of minutes (or even hours),
which is in marked contrast to the rapid activation kinetics of classical RTKs (89), (92). DDR1
29
expression is associated with increased cell motility, differentiation, proliferation and MMP-
mediated collagen degradation (6), (93), (121). Several fibrotic conditions of kidney, lung, liver
and cardiac tissue are associated with increased DDR1 expression (8), (110), (114), (117). DDR1
expression is also upregulated in aberrant wound healing and in certain forms of invasive cancer
(151), (152), (98).
In the context of how DDR1 may mediate force-induced collagen remodeling, previous
work showed that collagen-activated DDR1 associates with non-muscle myosin IIA (NMIIA)
(121). NMIIA is an abundant protein in cell adhesion complexes (153), (154). NMII can bind a
wide variety of proteins that modulate its contractile activity and thereby control transduction of
mechanical stimuli into biochemical signals, a process that can strongly influence collagen
remodeling (121). More recent data have defined the structural connections between DDR1 and
non-muscle myosin and its importance in DDR1-mediated collagen remodeling by traction (51).
In spite of these insights, the physiological mechanisms that regulate NMIIA-generated force
transmission through DDR1 is not defined. Here we used tandem mass tagged mass spectrometry
and a cell model that does not express b1 integrins. We identified the myosin-related regulator of
RLC dephosphorylation, Myosin Phosphatase Rho-interacting Protein (MRIP), as a central
determinant of how tractional forces and cell adhesion to collagen mediated by DDR1 are
regulated.
30
Materials and methods
Reagents
Rabbit and goat polyclonal anti-DDR1 (C-20, sc-532) and goat polyclonal anti-MRIP (C-14, sc-
135494) antibodies were from Santa Cruz Biotechnology (Santa Cruz, CA). Rabbit polyclonal
antibody to myosin light chain 2, and rabbit polyclonal to phospho-myosin light chain 2 (ser19)
were from Cedarlane (Burlington, ON). Rabbit anti-IgG antibody was purchased from Abcam
(Cambridge, United Kingdom). Rabbit polyclonal anti-MRIP antibody was from Novus
Biologicals (Littleton, CO). Rabbit monoclonal anti-phospho DDR1 (Y792 and Y513) and rabbit
monoclonal anti-MRIP (D8G8R) antibodies were from Cell Signaling Technology (Danvers,
MA). Rabbit polyclonal antibody to non-muscle myosin was from Biomedical Technologies
(Stoughton, MA). Rat monoclonal anti-mouse CD29 (9EG7) antibody, type I rat tail collagen,
and type I bovine collagen were purchased from BD Biosciences (Mississauga, ON, Canada).
Mouse monoclonal antibody to β-actin, fibronectin from bovine plasma, and puromycin were
from Sigma-Aldrich (St. Louis, MO). G 418 disulfate salt was from Thermo Fisher Scientific
(Waltham, MA). Alexa Fluor® 488 goat anti-mouse, Alexa Fluor® 568 goat anti-rabbit
antibodies, and rhodamine phalloidin, were from Life Technologies (Burlington, ON).
Fluoresbrite YG Microspheres 2.0 m were from Polysciences Inc. (Warrington, PA).
FluoSpheres Carboxylate-modified 2.0 µm microspheres (580/605) were from Molecular Probes
(Eugene, OR). Nilotinib was obtained from Reagents Direct (Encinitas, CA). Y-27632 was from
Calbiochem (Millipore; Billerica, MA).
Cells
Mouse NIH-3T3 were provided by the late Wolfgang Vogel (University of Toronto, Toronto,
Canada). Integrin β1-deficient GD25 cells were provided by Dr. Reinhard Fässler (Max-Planck
31
Institute for Biochemistry, Munich, Germany). Cells were cultured at 37°C in complete DMEM
(10% fetal bovine serum and 10% antibiotics). GD25 WT cells required a selective growth
medium supplemented with 2 µg/mL puromycin. GD25 OE and B1 MRIP KO cell lines required
a selective growth medium supplemented with 200 µg/mL G418. Cells were maintained in an
incubator under 5% CO2 conditions at 37°C, and were passaged with 0.25% trypsin containing 1
mM EDTA.
MRIP deletion by CRISPR/Cas9
The MRIP knockout cell line was generated in mouse GD25 OE (high DDR1 expression) cells
using CRISPR/Cas9 technology (Applied StemCell. Milpitas, CA). In this process, nucleotides
in exon 1 of mouse MRIP were removed, leading to a frame shift of the downstream MRIP
sequence, and the creation of a premature STOP codon, thus producing a knockout cell
model. To obtain this KO cell line, the following steps were conducted. First, a mixture of DNA
plasmids containing active gRNAs and the Cas9 gene was electroporated into GD25 OE cells.
The electroporation was carried out with a neon transfection system at 1450 V/30 ms/pulse
setting with 0.5 µg DNA per plasmid, using a 10 µl tip. Cells were cultured in drug-free medium
for 48 hours, and then selected using 2-3 µg/mL puromycin for a period of 24 to 48 hours. The
cells were then transferred back to drug-free medium. A small portion of the cell culture,
presumably comprising a mixed cell population, was subjected to genotype analysis. If the mixed
culture showed indels (presence of insertions or deletions), indicating Cas9 cleavage and/or
genome modifications, it was subjected to single cell cloning process. Once confirmed as
positive, single cell cloning was carried out by end-point dilution and culture. Cell clones were
then genotyped by PCR and sequencing. Five KO clones were expanded, and the genotypes were
further confirmed by sequencing.
32
Tandem mass tagged mass spectrometry
DDR1 adhesion complexes were prepared as previously discussed (51) using a C-terminal anti-
DDR1 antibody bound to superparamagnetic DynaBeads. GD25 WT and GD25 OE cells were
plated on either collagen or fibronectin for one or eight hours prior to lysis and
immunoprecipitation. Samples were reduced, alkylated, digested, and TMT-labelled according to
the manufacturer’s directions (Thermo Fisher TMT 10 Plex). Labelled peptides from all samples
were combined and lyophilized. Peptides were resuspended in 2.5% TFA, and were then bound
to a homemade Strong Cation Exchange column (SCX). Peptides were eluted off in 1 fractions
(5% Ammonium Hydroxide, 80% ACN), and then lyophilized.
Samples were analyzed on an Orbitrap analyzer (Q-Exactive, ThermoFisher, San Jose,
CA) outfitted with a nanospray source and EASY-nLC nano-LC system (ThermoFisher, San
Jose, CA). Lyophilized peptide mixtures were dissolved in 0.1% formic acid and loaded onto a
75 µm x 50 cm PepMax RSLC EASY-Spray column filled with 2 µM C18 beads (ThermoFisher
San, Jose CA) at a pressure of 800 Bar. Peptides were eluted over 120 min at a rate of 250 nl/min
using a gradient set up as 0% - 40% gradient of Buffer A (0.1% Formic acid; and Buffer B, 0.1%
Formic Acid in 80% acetonitrile). Peptides were introduced by nano-electrospray into the Q-
Exactive mass spectrometer. The instrument method consisted of one MS full scan (525–1600
m/z) in the Orbitrap mass analyzer with an automatic gain control target of 1e6, maximum ion
injection time of 120 ms and a resolution of 35,000 followed by 15 data-dependent MS/MS scans
with a resolution of 35,000, an AGC target of 1e6, maximum ion time of 100 ms, and one
microscan. The intensity threshold to trigger a MS/MS scan was set to an underfill ratio of 1.0%.
Fragmentation occurred in the HCD trap with normalized collision energy set to 27. The
dynamic exclusion was applied using a setting of 20 seconds.
33
Tandem mass spectra were extracted, charge states deconvoluted and deisotoped by
Xcalibur version 2.2. All MS/MS samples were analyzed using Sequest (Thermo Fisher
Scientific, San Jose, CA, USA; version 1.4.1.14) and X! Tandem (The GPM, thegpm.org;
version CYCLONE (2010.12.01.1)). Sequest was set up to search
Uniprot_Mouse_Nov_18_2015.fasta (Downloaded Nov 18 2015, 74993 entries) assuming the
digestion enzyme trypsin. X! Tandem was set up to search the Uniprot_Mouse_Nov_18_2015
database (unknown version, 77779 entries) also assuming trypsin. Sequest and X! Tandem were
searched with a fragment ion mass tolerance of 0.020 Da and a parent ion tolerance of 10.0 ppm.
Carbamidomethyl of cysteine and TMT6plex of lysine and the n-terminus were specified in
Sequest and X! Tandem as fixed modifications. Glu->pyro-Glu of the n-terminus, ammonia-loss
of the n-terminus, gln->pyro-Glu of the n-terminus and oxidation of methionine were specified in
X! Tandem as variable modifications. Oxidation of methionine was specified in Sequest as a
variable modification.
Scaffold (version Scaffold_4.7.5, Proteome Software Inc., Portland, OR) was used to
validate MS/MS based peptide and protein identifications. Peptide identifications were accepted
if they could be established at greater than 95.0 % probability. Peptide probabilities from X!
Tandem were assigned by the Peptide Prophet algorithm (155) with Scaffold delta-mass
correction. Peptide Probabilities from Sequest were assigned by the Scaffold Local FDR
algorithm. Protein identifications were accepted if they could be established at greater than 95.0
% probability and contained at least 1 identified peptide. Protein probabilities were assigned by
the Protein Prophet algorithm. Proteins that contained similar peptides and could not be
differentiated based on MS/MS analysis alone were grouped to satisfy the principles of
parsimony.
34
Shear stress assay
Microspheres (2.0 µm diameter Fluoresbrite YG) were incubated with 1 mL of neutralized (pH
7.4; 0.1 M NaOH) bovine type I collagen (1.0 mg/mL) for 20 min at 37 °C. 2.0 µm diameter
carboxylate-modified fluorescent microspheres (580/605) were incubated with 1mL of 1% (w/v)
BSA for 30 min at 37°C. Beads were pelleted and re-suspended in 1x PBS for storage and use. ~
5.0 x 104 cells were plated and incubated overnight (37°C; 5% CO2) in 24-well tissue-culture
plastic plates. Beads were counted using a hemocytometer and equal quantities were incubated
with the cells in an 8:1 bead-to-cell ratio for 2 hours (37°C; 5% CO2). Medium was carefully
removed and each well received 200 µL of 1x PBS. Cells were then exposed to 200 µL washes
(0, 1, 2, 4, 8, 16) of 1x PBS using a multichannel pipettor from ~1 cm above the plating surface.
Under certain conditions cells were incubated with either Y-27632 (10 M) or Nilotinib (2 µM);
DMSO served as a control. Cells were fixed using 4% paraformaldehyde for 10 min, stained
with DAPI (10 µg/mL) for 15 minutes and then covered with ~ 500 µL 1x PBS. Cells were
identified by DAPI-stained nuclei and a minimum of 30 cells were counted per well. The number
of bound beads associated with each cell was counted using a fluorescence microscope within a
10 x 5-mm rectangular zone in the middle of the well. Four wells were inspected per condition
and each experiment was repeated a minimum of three times.
Collagen gel contraction
Type 1 bovine collagen (3.03 mg/mL) was added to a solution consisting of DMEM, 0.25 M
NaHCO3, FBS, 10X antibiotic, and 0.1 M NaOH. Cells were added to the solution to produce a
final concentration of 1 x 106 cells/mL. 200 µL aliquots of the gel-cell solution were pipetted into
the centre of each well of a 24-well non-tissue culture plate, ensuring that no gel contacted the
sides. The gel-cell solution was polymerized for 5 mins at room temperature before adding 1 mL
35
of growth medium to each well. The gel-cell solutions were incubated for 4 days (37°C; 5%
CO2); gels were checked daily and supplemented with ~250 µL of extra growth medium or
growth medium with inhibitors as needed. Measurements of the collagen gel diameters were
made with a dissecting microscope and an inter-ocular grid starting at time 0 hrs and every 15
minutes thereafter for a total of three hours post-release.
Analysis of collagen alignment and compaction
Glass coverslips (plasma-treated, treated with 2% APTES and 0.1% gluteraldehyde) were coated
with ~120 µL bovine type I collagen (1 mg/mL), neutralized (pH 7.4; 0.1 M NaOH). Cells (~
30,000 per slip) were incubated at 37°C; 5% CO2. After incubation, the samples were rinsed
twice with 1x PBS, fixed in 4% paraformaldehyde, blocked with 1% BSA and stained for 45
minutes at room temperature with rhodamine phalloidin. Images were obtained with a Leica TCS
confocal microscope (Mannheim, Germany). Collagen fibers were visualized using confocal
reflectance microscopy. Analysis and quantification of acquired images was done using Image J.
The fluorescence intensities of collagen in fixed area regions of interest were measured in each
sample. Fast Fourier Transform was used to quantify the alignment of collagen fibrils from
confocal microscopy images as described in (156). Alignment indices were quantified by
calculation of area under the intensity curve within ±10º of the peak. Values represent mean and
standard errors of the mean.
Immunoprecipitation and immunoblotting
Cells were lysed using a commercially purchased lysis buffer (Thermo Fisher RIPA Buffer)
supplemented with protease inhibitor cocktail on ice, sedimented (10,000 rpm at 4°C for 10
minutes) and for immunoprecipitation, 1 mg of solubilized protein was incubated with 0.5 µg
polyclonal anti-DDR1, or anti-NMIIA antibody, along with protein G superparamagnetic
36
DynaBeads at 4°C overnight. Beads were washed three times with buffer and proteins were
eluted with 4x Laemmli buffer or 1 mM DTT in 2x Laemmli buffer and then boiled at 95°C for 7
minutes. Proteins were separated by SDS-PAGE and transferred to nitrocellulose membranes.
The membranes were probed with the indicated antibody followed by goat anti-mouse, goat anti-
rabbit, or donkey anti-goat fluorophore-conjugated secondary antibody. Blot density analysis
was done with a LiCor Odyssey. All western blots were repeated at least three times and
representative data sets are presented.
Protein fractionation experiments
Cells (~ 6.5 x 106 ) were plated on type I fibrillar collagen (1 mg/mL) -coated 150-mm tissue
culture plastic plates and incubated for 24 hours (37°C; 5% CO2). The next day cells were
washed with 1x PBS twice, covered and placed on ice for 20 minutes in a hypotonic buffer
solution (1 mM NaF; 10 mM Iodoacetamide; 10 mM tris). The hypotonic buffer solution
promotes release of the cytosol and represents the “cytoplasmic fraction”. Cells were scraped,
collected, and centrifuged at 10,000 rpm for 4 minutes at 4°C. The supernatant (cytoplasmic
fraction) was collected and the resultant pellet was broken down using RIPA Lysis Buffer
(Thermofisher) containing 1% Triton X-100. Subsequently, the sample was centrifuged at 10,000
rpm for 3 minutes at 4°C. The supernatant was collected, which represents the Triton-soluble
fraction. The resulting pellet from this final centrifugation represents the Triton-insoluble
fraction.
Confocal microscopy and immunostaining experiments
Cells were seeded on either 1 mg/mL type I fibrillar collagen- (neutralized with 1.0 M NaOH) or
fibronectin-coated glass-bottom MatTek dishes (~100-150 µL volume). Cells (~ 30,000 per slip)
were incubated at 37°C; 5% CO2 for specified time points. After incubation, the samples were
37
rinsed twice with 1x PBS, fixed in 4% paraformaldehyde, blocked with 1% BSA in PBS,
permeabilized and stained with DAPI in NP-40 (10 µg/mL) then stained for 45 minutes at room
temperature with rhodamine phalloidin, if identification of F-actin was required. For co-
localization studies of DDR1, MRIP and NMIIA, cells were incubated with the appropriate
primary antibody (in 1% BSA in PBS) for 1h. The corresponding secondary antibody was
incubated with cells in the same manner as noted above for 1h. The cells were washed twice with
1x PBS between primary and secondary incubations as well as after secondary incubation.
Images were obtained at either 40x or 60x magnification with a Leica TCS confocal microscope
(Mannheim, Germany). Scale bars represent 20 µm. Pearson correlation coefficients were
determined using the “colocalization2” plugin in Fiji.
Statistical analysis
For all continuous variable data, mean and standard errors of means were computed.
When appropriate, comparisons between two samples were made by Student’s t-test (unpaired).
For multiple samples, ANOVA was used and differences between groups were assessed with
Tukey’s post-hoc test. Statistical significance was set at a type I error rate of p<0.05. All
experiments were performed in triplicate unless otherwise stated.
38
Results
DDR1-associated adhesion complexes exhibit collagen-dependent protein interactions
While the association of DDR1 with NMIIA is important for cell migration on collagen (121) it
is not currently known whether this association is important for cell-induced mechanical
remodeling of fibrillar collagen. I examined whether there was a collagen-specific repertoire of
proteins that contribute to the function of DDR1-adhesion complexes. For these experiments I
used GD25 cells, which are a β1-integrin knockout (KO) cell line (Figure 1A) (157). As the β1
integrin subunit is expressed in all integrins that directly bind collagen (157), the use of cells that
are null for β1 integrin facilitates definition of the role of DDR1 in adhesive functions. GD25 WT
cells (low constitutive levels of DDR1 expression; Figure 1B) and GD25 OE cells (high DDR1
expression as a result of stable transfection with DDR1b; Figure 1B) (92) were plated on fibrillar
collagen or fibronectin (as a control substrate that is not bound by DDR1) for 1 or 8 hours
(Figure 2). DDR1 immunoprecipitates prepared from these lysates were analyzed by tandem
mass tagged mass spectrometry (TMT/MS) to characterize DDR1-associated proteins. There was
increased abundance (>4-fold) of NMIIA that associated with DDR1 in GD25 OE cells when
plated on collagen compared with the same cells plated on fibronectin at 1 hour. A similar
increase (>3.5-fold) of NMIIA was found upon comparing WT cells plated on collagen for 1
hour or 8 hours (Figure 2).
An unexpected protein that was detected by mass spectrometry in the DDR1
immunoprecipitates and that exhibited similar patterns of change in abundance between
experimental groups was Myosin Phosphatase Rho-interacting Protein (MRIP; Figure 2), which
is an indirect inhibitor of Myosin Light Chain (MLC) phosphorylation. Comparisons of GD25
OE cells plated on collagen for 1 hour versus GD25 WT cells showed that the abundance of
39
MRIP in DDR1-associating proteins increased >10-fold. There was also increased abundance
(3.5-fold) of MRIP in these preparations in cells that had been plated on fibronectin. These
variations were consistent over time: in GD25 OE cells plated on collagen versus fibronectin for
8 hours there was a 4-fold increase of MRIP abundance.
MRIP co-localizes with F-actin in a collagen- and DDR1-dependent manner.
Previous work has shown that MRIP not only co-localizes with the actin cytoskeleton to mediate
stress fiber assembly in non-muscle cells (158) but it acts as an accessory protein to indirectly
induce dephosphorylation and disassembly of NMIIA filaments via targeting of the Myosin
Phosphatase Target Subunit 1 (MYPT1) of Myosin Phosphatase (MP) to the MLC (149). I
examined the effects of DDR1 expression and substrate-specificity on the co-localization of
MRIP with actin filaments by staining with rhodamine phalloidin. GD25 WT and GD25 OE cells
were plated on either collagen or fibronectin for 24 hours. There was no significant impact of
collagen or fibronectin-plating on the co-localization of MRIP (green) with F-actin (rhodamine
phalloidin) (Figure 3A; top-left and bottom-left panels) in GD25 WT cells. GD25 OE cells
showed significantly higher (p< 0.001) co-localization of MRIP with F-actin compared to GD25
WT cells when plating on collagen (Figure 3A; top-left and top-right panels). The effect of
substrate-plating was also strong as GD25 OE cells plated on collagen showed significantly
higher (p< 0.0001) co-localization of MRIP with F-actin compared to GD25 OE cells plated on
fibronectin (Figure 3A; top-right and bottom-right).
MRIP co-localizes with DDR1 and NMIIA on collagen; DDR1 co-localization with NMIIA
is affected by substrate and MRIP expression but not time
I examined whether MRIP co-localizes with DDR1 and NMIIA. In GD25 OE cells (high DDR1
expression) plated on collagen for 24 hours, MRIP strongly co-localized with NMIIA (Figure
3B; left panel) and DDR1 (Figure 3B; right panel; Pearson Correlation Coefficients r ~0.8
40
respectively). Because of the dynamic nature of NMIIA filament assembly and disassembly in
spreading and migrating cells (144), I examined the effect of plating time on the association of
MRIP with NMIIA and DDR1. GD25 OE cells were plated on collagen for 4, 8, and 24 hours
(Figure 4) and immunostained for DDR1 (red) and MRIP (green; Figure 4A). DDR1 and MRIP
strongly co-localized at all time points; the strongest co-localization was at 24 hours (r: ~0.7) but
there were no significant differences between time points. Under the same conditions as
described above, MRIP (green) strongly co-localized with NMIIA (red) at all time points (Figure
4B). The strongest co-localization of MRIP with NMIIA was at 8 hours (r: ~0.7). Time of plating
did not impact the extent of co-localization of MRIP with NMIIA (p>0.2).
To examine the effect of MRIP expression and plating time on the association of DDR1
with NMIIA, an MRIP KO cell line was generated from GD25 OE parental cells using CRISPR-
Cas9 technology (designated as B1 MRIP KO cells; Figure 5). In cells plated on collagen for 4,
8, and 24 hours, DDR1 strongly co-localized with NMIIA for all groups (Pearson Correlation
Coefficients of ~0.6-0.8; Figure 6A). Deletion of MRIP affected the association of DDR1 with
NMIIA since at 4 (p< 0.05) and 8 (p< 0.01) hours, their co-localization was stronger in B1 MRIP
KO cells than GD25 OE cells. Time of plating did not affect co-localization for the various
groups. There was a substrate-dependent effect on the association of DDR1 with NMIIA when
MRIP was deleted in that DDR1/NMIIA co-localization was higher (p< 0.001) when B1 MRIP
KO cells were plated on collagen compared with fibronectin (Figure 3C).
DDR1, NMIIA, and MRIP localize to discrete cellular fractions
The polymerization of cytoskeletal proteins (e.g. actin, non-muscle myosin) is essential for
various cellular processes including determination of cell polarity, migration and force
production (159), (160), (161), (153). I used a cell compartment fractionation protocol, which
41
included the use of specific buffers, detergents and centrifugation protocols as previously
described (162), to assess the effect of substrate and protein expression on the relative abundance
of DDR1, NMIIA and MRIP in cell fractions prepared from GD25 WT, GD25 OE, B1 MRIP
KO and F2 MRIP KO cells plated on collagen. The fractionation protocol enables estimation of
the relative abundance of proteins in 3 specific cellular fractions: a Cytoplasmic Fraction
(denoted “CyPl”), a Triton-soluble fraction (denoted “TS”), and a Triton-insoluble fraction
(denoted “TI”) (Figure 7A). There were no significant differences of the abundance of DDR1,
MRIP and NMIIA in discrete fractions between cell types; plating cells on collagen also had no
significant effect on DDR1 fractionation patterns. Although there were no statistical differences
between the experimental groups, DDR1 was most abundant in the Triton-soluble and Triton-
insoluble fractions of all 4 cell types (Figure 8A). MRIP was more abundant in the Triton-soluble
fraction of GD25 OE cells than the cytoplasmic (p< 0.01) and Triton-insoluble fractions (p<
0.001; Figure 8B). These patterns of abundance were not the same in GD25 WT cells, which
showed a greater relative abundance of MRIP in the cytoplasmic fraction. Finally, NMIIA was
most concentrated in the cytoplasmic fraction of all 4 cell types but there were no significant
differences between cell types. Within each cell type, NMIIA was more abundant in the
cytoplasmic fraction than the Triton-soluble or Triton-insoluble fractions (p< 0.0001; Figure 8C).
DDR1 adhesions to collagen are affected by NMIIA, DDR1 activity and MRIP expression
To assess the effect of DDR1 expression on the formation of strong adhesions to collagen I
incubated cells with fibrillar collagen- or BSA-coated (not shown) fluorescent microspheres and
exposed cells to repeated shear forces (~3.5 Pa/wash) as described (27). On average, GD25 OE
cells exhibited more bound collagen-coated beads per cell than GD25 WT cells (comparisons of
42
washes 0 and 1; p< 0.0001; Figure 9A; black vs grey). Subsequent washes showed no significant
difference between cell types.
Next, to examine the role of DDR1 activation in the formation of strong collagen
adhesions we treated GD25 OE cells with nilotinib (2 µM), a specific inhibitor of DDR1 tyrosine
autophosphorylation and activation (163) or vehicle. GD25 OE cells treated with nilotinib
exhibited impaired binding of collagen-coated beads. There was a marked reduction of bead
binding after treatment at wash 1 (p< 0.0001; Figure 9A; black vs brown); washes 0, 2, 4, 8, and
16 resulted in no differences (p>0.2) between conditions.
NMIIA contractile activity and filament assembly are regulated, in part, by the relative
phosphorylation state of its two regulatory light chains (164). Rho-associated coiled-coil kinase 1
(ROCK1), a serine/threonine kinase acting downstream of RhoA, directly phosphorylates the
regulatory light chains of NMIIA to initiate filament assembly and contraction (165). To
examine the effect of NMIIA filament assembly and activation on the ability of cells to strongly
adhere to collagen-coated beads we treated GD25 OE cells with a potent inhibitor of ROCK1
activity, Y27632 (166), or vehicle. On average, cells treated with Y27632 exhibited decreased
binding to collagen-coated beads compared with vehicle at washes 0 and 1 (p< 0.0001; Figure
9A; black vs red). Subsequent washes showed no significant differences. GD25 WT cells treated
with Y27632 or vehicle (not shown) exhibited no differences (p> 0.2) of collagen-coated bead
binding after exposure to shear stress.
To assess the role of MRIP expression in resistance to shear stress, I compared collagen
bead binding in GD25 OE cells with B1 MRIP KO cells and with F2 MRIP KO cells, a second
MRIP KO cell line generated by CRISPR-Cas9 (Figure 5) (low DDR1 expression; MRIP KO).
There was no difference of bead binding between GD25 OE and B1 MRIP KO cells for all
43
conditions (p>0.2). There was a reduction of bead binding between GD25 OE and F2 MRIP KO
cells at washes 0 and 1 (p< 0.0001) but there were no differences after more washes (p>0.2). I
also found that B1 MRIP KO cells exhibited more bound beads per cell after exposure to shear
force compared with F2 MRIP KO cells at washes 0 (p< 0.0001), 1 (p< 0.0001), and 2 (p< 0.01).
DDR1 expression and activity, NMIIA filament assembly, and MRIP expression regulate
force transmission to fibrillar collagen
Mechanical reorganization of fibrillar extracellular matrix proteins requires actomyosin-
generated contractile forces (51), (93). To examine the effect of DDR1 expression on cell-
generated forces I embedded GD25 WT or GD25 OE cells in anchored fibrillar collagen gels (~1
x 106 cells/mL) (87) (167). After 4 days of incubation collagen gels were released from the
bottom of dishes and gel contraction was measured over a period of 3 hours in 15 minute
increments. GD25 OE cells effectively contracted collagen gels (~9.7 mm to ~6.1 mm, ~26%
reduction in diameter) over a 3 hour period whereas GD25 WT cells contracted gels less
vigorously (from ~10 mm to ~8.7 mm; ~13% reduction; Figure 10A; grey vs black; 15 minutes-
p< 0.01; and p< 0.0001 for later time points).
I examined the effect of DDR1 activation on collagen contraction by treating GD25 OE
cells with nilotinib (2 µM) or vehicle (Figure 10A; black vs brown). Inhibition of DDR1
autophosphorylation and activation reduced collagen gel contraction (p< 0.05 at 45 minutes and
p< 0.0001 for time points thereafter). I blocked NMIIA filament assembly and contractile
activity by treating GD25 OE cells with Y27632 (10 µM) or vehicle (Figure 10A; black vs red).
In GD25 OE cells this treatment impaired collagen gel contraction (~10.5 mm to ~9.6 mm)
which was not observed in the control group (time 0- p< 0.01 and p< 0.0001 for time points
thereafter). I assessed the effect of MRIP expression on cell-generated collagen gel contraction in
44
GD25 OE and B1 MRIP KO cells. Compared with GD25 OE cells, B1 MRIP KO cells exhibited
enhanced collagen gel contraction (~9.2 mm to ~4.6 mm; ~50% reduction in size; Figure 10B;
blue; p< 0.05 at 105 minutes; p< 0.001 at 120 minutes and (p< 0.0001) thereafter.
NMIIA filament assembly and MRIP expression affect fibrillar collagen remodeling
Cell-mediated mechanical remodeling of fibrillar collagen has been quantified from
measurements of collagen alignment and collagen compaction (51), (67), (156). Collagen fiber
alignment is the process by which cells mechanically reorganize collagen fibers into non-random
spatial orientations. Collagen compaction is an apparently distinct dynamic process by which
cells aggregate collagen fibers into clusters in the cell periphery. DDR1 overexpression enhances
collagen fiber alignment and compaction (51). I assessed the role of myosin contractility on
collagen fiber alignment by comparing GD25 OE cells plated on fibrillar collagen gels that were
treated with Y27632 (Figure 11A; right panel) or vehicle (Figure 11A; middle panel) and
analyzed as described (156). There was a 30% reduction of collagen fiber alignment in GD25 OE
cells treated with Y27632 (Alignment Index : ~1.6) compared with vehicle (Alignment Index :
~2.4) (p< 0.01; Figure 11B; left). Similarly, there was a marked decrease in collagen compaction
in treated cells (p< 0.01; Figure 11B; right).
To quantify the effect of MRIP expression on fibrillar collagen remodeling I compared
GD25 OE cells (Figure 11C; black) with B1 MRIP KO cells (Figure 11C; striated). There was
25% higher collagen fiber alignment in MRIP knockout cells compared with GD25 OE cells (p<
0.01; Figure 11C; left). MRIP KO did not have the same effect on collagen fiber compaction
(Figure 11C; right) as GD25 OE cells compacted collagen more effectively than MRIP KO cells
(p< 0.0001).
45
Substrate affects site-specific phosphorylation of DDR1 and MLC; MRIP deletion
enhances DDR1 and MLC phosphorylation.
The physiological mechanisms that mediate DDR1 tyrosine autophosphorylation and activation
are not fully understood. Several key phospho-sites have been identified in the kinase domain
region (activation loop) of DDR1, which corresponds to collagen-induced receptor activation.
One of these residues is tyrosine 792 (Y792) (6). I compared the effect of substrate by plating
GD25 WT, GD25 OE, B1 MRIP KO and F2 MRIP KO cells on collagen or fibronectin for 24
hours (Figure 12). Western blotting showed no effect of substrate-coating on total DDR1
expression, as expected (Figure 12A) but plating cells on collagen enhanced DDR1
phosphorylation (Y792) in GD25 OE cells by >70% compared with GD25 OE cells plated on
fibronectin (p< 0.01; Figure 12B). Compared with GD25 OE cells, B1 MRIP KO cells exhibited
>40% more phosphorylation of Y792 when plated on collagen (p< 0.001; Figure 12B). I also
observed 85% higher Y792 phosphorylation in B1 MRIP KO cells plated on collagen compared
with fibronectin (p< 0.0001; Figure 12B). These phenomena were not observed in a second
tyrosine residue of DDR1 (Y513) that is located in the juxtamedullary (JM) region, and which is
not associated with collagen-induced receptor activation (6). This residue showed no marked
changes of phosphorylation between the different cell types or under conditions when cells were
plated on collagen or fibronectin (Figure 12C).
NMIIA activation and contractility are regulated by several distinct, physiological
mechanisms, which are calcium-dependent (144) and calcium-independent (165), (168), (154).
Phosphorylation of serine 19 (S19) of the regulatory light chain of NMIIA is mediated by several
kinases (e.g. ROCK1), which enhances actomyosin contraction. In contrast, phosphatases (e.g.
Myosin phosphatase) induce actomyosin relaxation. MRIP indirectly regulates phosphorylation
46
of NMIIA (thereby promoting filament disassembly and relaxation) by sequestering
RhoA/ROCK1 and targeting Myosin phosphatase (164), (149), (148) to the regulatory light
chain. To examine the role of MRIP in regulating NMIIA phosphorylation in a DDR1-specific
context, I plated GD25 WT, GD25 OE, B1 MRIP KO, and F2 MRIP KO cells on collagen or
fibronectin for 24 hours. Western blotting showed no substrate or cell type-specific differences
of total NMIIA expression (Figure 13A) or total MLC expression (Figure 13B). There were
DDR1-dependent differences of pMLC S19 phosphorylation: GD25 OE cells showed enhanced
phosphorylation compared to GD25 WT cells when plated on collagen (p< 0.05; Figure 13C).
B1 MRIP KO cells showed 30% higher S19 phosphorylation compared with GD25 OE cells
plated on collagen (p< 0.01; Figure 13C). B1 MRIP KO cells also showed increased S19
phosphorylation when plated on collagen compared with fibronectin (>80%; p< 0.0001; Figure
13C).
47
Discussion
Following injury or the resolution of inflammation, tightly regulated remodeling of fibrillar
collagen is a critical determinant of the structural integrity and homeostatic function of a large
number of tissues and organs (120). Collagen remodeling is mediated by pericellular proteolysis
(56), by intracellular degradation involving the phagocytic route (64) and by mechanically-
driven tractional reorganization in which cells apply actomyosin-generated contractile forces to
compact and reorient fibrillar ECM proteins (122). My principal finding here is that DDR1,
NMIIA and MRIP are enriched in an adhesion complex that mediates collagen remodeling
through the application of tractional forces via DDR1. I found that MRIP is an important
regulator of NMIIA function in these adhesion complexes. Acting together, MRIP and NMIIA
exert a strong impact on DDR1-mediated cell adhesion to collagen and affect the contraction,
alignment and compaction of collagen fibrils. Collectively, these data indicate that MRIP is
positioned at an important regulatory locus in the DDR1 adhesion complex to regulate collagen
remodeling.
Extensive previous data have demonstrated the importance of the fibrillar collagen
receptors, the α2β1 and α11β1 integrins, in maintaining the structural integrity and function of
fibrillar collagen (89). For specific study of the role of DDR1 in collagen remodeling, I used a β1
integrin knockout cell line, GD25 cells (157), which express very low levels of DDR1b (GD25
WT cells) or cells that express high levels of DDR1b (GD25 OE cells) (92), which is
similar to the expression levels exhibited by certain types of invasive cancer cells (7). The GD25
cell models enabled examination of DDR1 function in the remodeling of fibrillar collagen that is
independent of the function of the β1 integrin.
48
For identifying novel proteins that are critical for DDR1-specific adhesive function in
fibrillar collagen remodeling, I prepared DDR1 immunoprecipitates from GD25 WT or GD25
OE cells that had been plated on collagen or fibronectin as a control substrate. I then used
tandem mass-tagged mass spectrometry to define DDR1-associating proteins that are recruited to
a collagen-specific complex. One of my principal findings is that the signaling protein, MRIP, is
enriched in DDR1-adhesion complexes and that MRIP regulates NMIIA function in a collagen-
dependent fashion.
The transmission of cell-generated traction forces through adhesion complexes to the
ECM relies in part on the ability of fibrillar collagen receptors and contractile motor proteins to
interact functionally with actin filaments. Previous data from vascular smooth muscle cells
treated with lysophosphatidic acid showed that MRIP co-localizes with actin filaments (148). I
found by immunoprecipitation that MRIP associates with DDR1 and NMIIA, and that in DDR1-
expressing cells plated on collagen, MRIP co-localizes with actin filaments. The tight spatial
relationships between DDR1, NMIIA, MRIP and actin filaments indicate that these proteins may
aggregate to form macromolecular structures that are analogous to the classical focal adhesion
complexes found in non-muscle cells, such as cultured fibroblasts (169). The assembly of these
complexes appears to be initiated after DDR1 begins to bind to fibrillar collagen.
Integrin-dependent ECM adhesions in cultured cells are dynamic structures (169) that
exhibit extensive and relatively rapid reorganization after plating of cells on ECM proteins (170),
(171). This reorganization relies upon the repeated cycling, association and disassociation of
protein aggregates in the adhesions. I found that for DDR1-dependent adhesions to collagen,
there was strong co-localization of DDR1 with MRIP and of NMIIA with MRIP, but that in
contrast, the extent of co-localization was not affected by the time of plating on collagen.
49
Previous work has demonstrated a general relationship between the solubility of proteins and
their aggregation into supramolecular structures: large, complex structures that contain multiple
proteins are often less soluble and exhibit reduced turnover compared with simpler structures
containing smaller proteins (172), (173). With the use of specific detergent protocols, I found
that DDR1, NMIIA and MRIP were enriched in certain specific cellular compartments based on
detergent solubilities (162) but that these proteins did not aggregate within the same
compartments. The data from these experiments indicated that MRIP is a relatively stable
component of DDR1/NMIIA-enriched adhesions and remains associated with these proteins
under the detergent conditions used here. However, it is evident that their aggregation into
supramolecular adhesion complexes may depend on other, as yet identified factors.
With the use of CRISPR-Cas9 technology, I developed two novel MRIP KO cell lines
that were derived from GD25 OE parental cells. This experimental approach allowed me to
examine the effect of MRIP expression on cell behavior in a model system that was devoid of b1
integrin function. I found that deletion of MRIP did not affect resistance to shear stress in
DDR1/collagen adhesions but that DDR1 overexpression, DDR1 activation (determined with
nilotinib, a potent inhibitor of DDR1 autophosphorylation (163)) and myosin light chain
phosphorylation (assessed with Y27632, a potent inhibitor of ROCK1 activity and of myosin
light chain phosphorylation (166)), all play a central role in the generation of DDR1 adhesions
that resist shear forces. In addition, I found that DDR1 activation, DDR1 expression levels and
myosin light chain phosphorylation also strongly affected cell-generated contraction of collagen
gels as did deletion of MRIP, indicating that MRIP may counteract the accumulation of cell
tension that is required for contraction of fibrillar collagen gels.
50
The compaction and alignment of collagen fibers is mediated by cell-generated
contractile forces (122). Previous work has shown that DDR1 associates with NMIIA to
modulate cell adhesion, migration and collagen compaction (31, 87) (121) but little is known of
how phosphorylation and dephosphorylation of NMIIA influence collagen alignment and
compaction. I found that in GD25 OE cells, impairment of RhoA/ROCK1-associated
phosphorylation of the myosin light chain, which initiates NMIIA filament assembly (144),
induced a marked decrease of collagen fiber alignment and compaction. Notably, deletion of
MRIP by CRISPR was also associated with a marked increase of collagen fiber alignment but
there was no increased collagen compaction. These data suggest that the alignment and
compaction of collagen fibers mediated by DDR1 are regulated by physiologically distinct
mechanisms. The data also suggest that MRIP may counteract the accumulation of cell-generated
tensile forces and that the phosphatase-related activity of MRIP may need to be tightly regulated
to enable collagen fiber compaction.
Finally, compared with wild type cells, I found that in cells with MRIP deletion, there
were marked differences of collagen-induced site-specific phosphorylation of DDR1 and
NMIIA. In DDR1, tyrosine 792 (Y792) is located in the activation loop (6) while tyrosine 513
(Y513) is a secondary phospho-site located in the juxtamembrane region of DDR1 that is not
associated with collagen-induced DDR1 activation. In cells spreading on collagen in which
MRIP was deleted, compared with wild type cells, there was enhanced phosphorylation at Y792
but not at Y513. These data indicate that MRIP is directly involved in the regulation of collagen-
induced DDR1 activation.
Phosphorylation of the myosin light chain at serine 19 (S19) is downstream of RhoA
activation and is induced by ROCK1 (165) to promote NMIIA filament assembly and cell
51
contraction (154). Previous work indicates that MRIP indirectly inhibits MLC phosphorylation
by targeting myosin phosphatase to the MLC, in order to dephosphorylate MLC to inhibit cell
contraction and blocks ROCK1 activity (148). I found that deletion of MRIP was associated with
enhanced MLC phosphorylation at S19 and that this association was dependent on DDR1
expression and collagen. These data further underline the central role of MRIP in regulating
force generation through DDR1-dependent collagen adhesions.
Collectively I show here that DDR1 adhesions to collagen are associated with the
activation of specific signals that impact specific cell behaviors that relate to tractional collagen
remodeling. My data suggest that MRIP is a central molecule in this DDR1 adhesion-related
signaling system and that this system exhibits cyclical feedback and feed-forward mechanisms
that regulate DDR1 activation and its role in collagen remodeling at the cell surface.
52
Figures
A
B
Figure 1. GD25 cells are null for β1 Integrin but have varying patterns of DDR1 expression. A) Western blot showing β1 Integrin expression levels in 3T3/NIH (3T3), GD25 WT (WT) and GD25 OE (OE) cell lines plated on plastic; GD25 cells are a genetic β1 Integrin KO cell line. B) Western blot showing DDR1 expression levels in 3T3/NIH (3T3), GD25 WT (WT) and GD25 OE (OE) cells plated on plastic; GD25 OE cells overexpress DDR1 via an expression plasmid.
53
D
B C
A
Figure 2. Tandem Mass Tagged Mass Spectrometry and Immunoprecipitation experiments show an association between DDR1, NMIIA, and MRIP. A) GD25 WT (low DDR1) and GD25 OE (DDR1 overexpression) cells were plated on Type I fibrillar collagen or fibronectin (control) for 1 and 8 hours; DDR1 was used as the immunoprecipitating antibody. Experiments were done in duplicates and then analyzed via 10-plex TMT/MS. Right panel shows partial list of relevant proteins discovered in TMT/MS. Values represent multiples of log2 fold increases in protein abundance. (B) TMT/MS reveals a collagen-dependent association between DDR1 and NMIIA (myosin heavy chain). (C) TMT/MS reveals a collagen-dependent association between DDR1 and Myosin Phosphatase Rho-interacting Protein (MRIP). (D) TMT/MS results were confirmed by plating GD25 WT (WT) and GD25 OE (OE) cells on collagen for 8 hours using DDR1 as the immunoprecipitating antibody (middle) and NMIIA as the immunoprecipitating antibody (right). A non-specific rabbit antibody directed towards IgG was used as a control (left). Data show mean ± SEM.
130kDa
WT OEIP TCL IP TCL
MRIP–116kDa
β-actin-42kDa
IP:NMIIA
NMIIA–220kDa220kDa
IPOE
MRIP–116kDa
IP TCLTCLWT
β-actin–42kDa
IP:DDR1
130kDa
DDR1–125kDa
130kDa
54
A
B
DDR1NMIIA
CollagenB1MRIPKO
Fibronectin
GD25 OE GD25 WT C
olla
gen
Fibr
onec
tin
MRIP Phalloidin
Collagen-coated
GD25OE
MRIPNMIIA DDR1MRIP
C
Figure 3. MRIP co-localizes with F-actin, DDR1, and NMIIA. MRIP expression levels affect DDR1/NMIIA co-localization. A) Immunohistochemical staining of GD25 cells plated on collagen or fibronectin for 24 hours. MRIP is stained in green, F-actin (phalloidin) is stained in red. MRIP co-localized with F-actin most effectively in GD25 OE cells when plated on fibrillar collagen compared with GD25 WT plated on collagen (p<0.001). MRIP also more effectively co-localized with F-actin when GD25 OE cells were plated on collagen compared with fibronectin (p<0.0001). Data represent mean + S.E.M. B) Immunohistochemical staining of GD25 OE (high DDR1) cells plated on collagen for 24 hours. On the left, MRIP (green) strongly co-localizes with NMIIA (red). On the right MRIP (red) strongly co-localizes with DDR1 (green). C) In the left panel, DDR1 (red) co-localizes with NMIIA (green) in GD25 OE (high DDR1) cells plated on collagen. DDR1 (green) strongly co-localizes with NMIIA (red) on collagen (middle) compared to fibronectin (right). Histograms show Pearson correlation of the spatial co-localization of the indicated proteins. Three grouped symbols are indicative of p<0.001. Four grouped symbols are indicative of p<0.0001. Data show mean ± SEM.
55
A
GD
25 O
E
4h 8h
DDR1/MRIP
24h
B
GD
25 O
E
4h 8h 24h
NMIIA/MRIP
Figure 4. MRIP does not exhibit a time-dependent change of co-localization with DDR1 or NMIIA when plated on collagen. A) Immunohistochemical staining of GD25 OE (high DDR1 expression) cells plated on collagen-coated glass-bottom MatTek dishes for 4 (left), 8 (middle), and 24 hours (right). DDR1 (red) and MRIP (green) did not exhibit significant time-dependent co-localization across conditions. B) Immunohistochemical staining of GD25 OE (high DDR1 expression) cells plated on collagen-coated glass-bottom MatTek dishes for 4 (left panel), 8 (middle panel), and 24 hours (right panel). NMIIA (red) and MRIP (green) did not exhibit significant time-dependent co-localization across conditions. Data show mean ± SEM.
56
A
B
Figure 5. B1 and F2 MRIP null clones exhibit different levels of DDR1 expression. A) MRIP expression patterns in GD25 WT (low), GD25 OE (moderate), B1 (KO), and F2 (KO) cell lines; B1 and F2 are MRIP KO clones generated by CRISPR-Cas9 from GD25 OE parent cells. B) DDR1 expression pattern in GD25 WT (low), GD25 OE (high), B1 (high), and F2 (low) cell lines.
57
A
GD
25 O
E 4h 8h
B1
MR
IP K
O
DDR1/NMIIA
24h
Figure 6. MRIP knockout enhances collagen-induced co-localization of DDR1 with NMIIA at varying times after plating. A) Immunohistochemical staining of GD25 OE cells (high DDR1 expression; top row) on collagen-coated glass-bottom MatTek dishes for 4 (left), 8 (middle), and 24 hours (right). There was no significant time-dependent change of co-localization of DDR1 (red) with NMIIA (green) across the different conditions. In the B1 MRIP KO cell line (MRIP KO; high DDR1 expression; bottom row) there was not a significant time-dependent change of co-localization of DDR1 (red) with NMIIA (green). When comparing the GD25 OE cells (high DDR1 expression; intact MRIP expression) to the B1 MRIP KO cells a significant difference of DDR1 co-localization with NMIIA was observed at 4 hours (p< 0.05) and 24 hours (p< 0.01). There was also a significant difference in DDR1/NMIIA co-localization between GD25 OE cells at 4 hours and the B1 MRIP KO cells at 24 hours (p< 0.05). One symbol is indicative of p<0.05. Two grouped symbols are indicative of p<0.01. Data show mean ± SEM. Data show mean ± SEM.
58
A
CyPl TS TI
GAPDH – 37 kDa
35 kDa
3T3 cells
CyPl TS TI
Gelsolin – 82 kDa
90 kDa
3T3 cells
CyPl TS TI
Vimentin – 57 kDa 55 kDa
3T3 cells
Figure 7. Proteins preferentially partition in response to various buffer conditions and centrifugations allowing for compartment-specific protein aggregations. A) Western blot images providing theoretical proof for a method by which whole cell lysates are fractionated into their constituent cell compartments (based on solubility). Three proteins were used as markers of each compartment. The first panel (top) shows a western blot for Glyceraldehyde 3-phosphate dehydrogenase (GAPDH), a ubiquitously expressed protein involved in glycolysis. In this case it is highly concentrated in the first fraction, the Cytoplasmic compartment (shortened to “CyPl” in subsequent images). The second panel (middle) shows a western blot for Gelsolin, a regulatory protein involved in actin filament assembly. Gelsolin partitions to the triton-soluble fraction, representative of a fraction rich in cytoskeletal proteins (shortened to “TS” in subsequent images). The third panel (bottom) shows a western blot for Vimentin, a type III intermediate filament. Previous studies have shown Vimentin to be largely insoluble by multiple detergents. In this case it represents a triton-insoluble fraction and is associated with a pellet rich in membrane and substrate-rich proteins (shortened to “TI” in subsequent images).
59
Cytoplasmic Triton-soluble Triton-insoluble0.0
0.2
0.4
0.6
0.8
1.0
Fractions
MR
IP fr
actio
n/to
tal M
RIP
GD25 WTGD25 OE
*****
Cytoplasmic Triton-soluble Triton-insoluble0.0
0.2
0.4
0.6
0.8
1.0
Fractions
NM
IIA fr
actio
n/to
tal N
MIIA
GD25 WTGD25 OEB1 MRIP KOF2 MRIP KO
****************
BA
C
A
Figure 8. DDR1 partitions to the cytoskeletal and pellet fractions; MRIP localizes to the cytoskeletal fraction; NMIIA primarily partitions to the cytoplasmic fraction regardless of cell type. A) Image shows a western blot of DDR1 fractionation patterns across GD25 WT, GD25 OE, B1 MRIP KO, and F2 MRIP KO cell lines. Cells were plated on type I fibrillar collagen (1 mg/mL) for 24 hours. DDR1 was found to preferentially partition to both the triton-soluble (cytoskeletal) and triton-insoluble (pellet) fractions while remaining relatively low in the cytoplasmic fraction. B) Western blot of MRIP fractionation pattern across GD25 WT, GD25 OE, B1 MRIP KO, and F2 MRIP KO cell lines. Cells were plated on type I fibrillar collagen (1 mg/mL) for 24 hours. MRIP preferentially partitions to the cytoskeletal fraction compared to the cytoplasmic (p< 0.01) and pellet (p< 0.001). Neither KO cell lines exhibition partitioning of MRIP as expected. C) Western blot of NMIIA fractionation pattern across GD25 WT, GD25 OE, B1 MRIP KO, and F2 MRIP KO cell lines. Cells were plated on type I fibrillar collagen (1 mg/mL) for 24 hours. NMIIA preferentially partitions to the cytoplasmic fraction. Across all cell types the cytoplasmic fractionation pattern of NMIIA was found to be significant (p< 0.0001). This same significance was not exhibited when comparing the effect of cell type within the cytoplasmic fraction. Two grouped symbols are indicative of p>0.01. Three grouped symbols are indicative of p<0.001. Four grouped symbols are indicative of p<0.0001. Data show mean ± SEM.
Cytoplasmic Triton-soluble Triton-insoluble0.0
0.2
0.4
0.6
0.8
1.0
Fractions
DD
R1
frac
tion/
tota
l DD
R1 GD25 WT
GD25 OEB1 MRIP KOF2 MRIP KO
60
A
B
Figure 9. Collagen adhesion is impacted by DDR1 and NMIIA regulatory factors. A) Shear wash assay showing collagen-coated bead binding to cells upon repeated exposure to force (~3.5 Pa/wash); DDR1 expression (black), DDR1 activity, impaired by using 2 µM Nilotinib (brown), and ROCK1 activity, impaired by using 10 µM Y-27632 (red) all impact adhesion strength. Significant differences are noted early on at washes 0 and 1. The various symbols denote comparisons between cell types. Four grouped symbols are indicative of p<0.0001. B) MRIP KO impacts collagen-coated bead adhesion; B1 MRIP KO clone (blue) has high DDR1 expression and F2 MRIP KO clone (green) has low DDR1 expression. Resistance to shear stress appears highest in B1 MRIP KO cells (high DDR1 expression; MRIP KO). Significant differences were most apparent in the F2 MRIP KO cell line, as their ability to strongly adhere to collagen-coated beads was impaired drastically compared to the GD25 OE and B1 MRIP KO cell lines. The results show that expression of DDR1 (a collagen receptor) is a more effective contributor to adhesion strength compared to MRIP expression as no significant differences were noted between the GD25 OE and B1 MRIP KO cells. The various symbols denote comparisons between cell types. Two grouped symbols are indicative of p<0.01 and four grouped symbols are indicative of p< 0.0001.
61
A
B
Figure 10. Collagen contraction is impacted by DDR1 and NMIIA regulatory factors. A) Floating collagen gel contraction assay; DDR1 expression (black), DDR1 activity (brown), and ROCK1 inhibition (red) impact contraction. B) MRIP KO impacts collagen contraction; GD25 OE (black) has high DDR1 expression and moderate MRIP expression, B1 MRIP KO clone (blue) has high DDR1 expression and KO for MRIP. Statistical analysis shows significant differences between GD25 OE and B1 MRIP KO cells at minutes 105 (p= 0.05), 120 (p< 0.001), 135 (p< 0.0001), 150 (p< 0.0001), 165 (p< 0.0001), and 180 (p< 0.0001). One symbol is indicative of p<0.05. Three grouped symbols are indicative of p<0.001. Four grouped symbols are indicative of p<0.0001. Data show mean ± SEM.
62
A
B
C
Figure 11. Effective collagen remodeling requires DDR1 expression and NMIIA filament assembly. MRIP deletion enhances this process. A) Stacked confocal images of Phalloidin-stained cells plated on Type I fibrillar collagen for 24 hours. DDR1 expression promotes effective collagen remodeling, GD25 WT(left) compared to GD25 OE (middle); Collagen remodeling is hindered by impairing NMIIA filament assembly with Y27632 (right). B) Stacked confocal images are analyzed using the Oval Profile ImageJ Plug-in, resulting data (Alignment Index) were plotted and analyzed using GraphPad Prism. GD25 OE cells significantly compact (p< 0.0029) and align (p< 0.0037) collagen fibers in the absence of Y27632. C) MRIP deletion enhanced collagen fiber alignment. B1 MRIP KO cells (grey pattern) align collagen fibers more effectively (p< 0.0039) than GD25 OE cells (black). Collagen compaction was found to be more effective in GD25 OE cells (black) compared with B1 MRIP KO cells (grey pattern) (p< 0.0001). Two grouped symbols are indicative of p>0.01. Two grouped symbols are indicative of p<0.01. Four grouped symbols are indicative of p<0.0001. Data show mean ± SEM.
63
A
B
Plated on collagen Plated on fibronectin
C
Figure 12. MRIP knockout enhances phosphorylation of DDR1 (Y792) in cells plated on collagen versus fibronectin. A) DDR1 expression is highest in GD25 OE cells and B1 MRIP KO cells when plated on collagen (left) for 24 hours compared with fibronectin (middle). GD25 WT cells show mild expression patterns of DDR1 regardless of the substrate they are plated on. B) On collagen, MRIP deletion (B1 cell line) shows enhanced DDR1 phosphorylation at Y792 (left) compared with fibronectin (middle), an activation-specific phospho-site (p< 0.0001). The B1 MRIP KO cells also show stronger phosphorylation of Y792, when plated on collagen, compared to GD25 OE cells (p< 0.001). C) A secondary DDR1 phospho-site, Y513 found in the JM region (bottom row), does not show similar results between cell types nor between varying substrates. Two grouped symbols are indicative of p>0.01. Three grouped symbols are indicative of p<0.001. Four grouped symbols are indicative of p<0.0001. Data show mean ± SEM.
64
B
A
C
Plated on collagen Plated on fibronectin
Figure 13. MRIP knockout enhances phosphorylation of MLC (S19) in cells plated on collagen versus fibronectin. A) NMIIA expression is uniform across cell types when plated on collagen or fibronectin. The difference between substrates is not notable. B) Total expression of the regulatory light chain of NMIIA (MLC) is uniform across cell types when plated on collagen. The difference between substrates is not notable. C) On collagen, the regulatory light chain of NMIIA (MLC) shows enhanced phosphorylation at S19 in the B1 MRIP KO (high DDR1) compared to GD25 OE cells (p< 0.01) and when plating B1 MRIP KO cells on fibronectin (p< 0.0001). S19 phosphorylation is associated with NMIIA filament assembly and cell contraction. S19 is phosphorylated by ROCK1 and dephosphorylated by Myosin phosphatase. One symbol is indicative of p<0.05. Two grouped symbols are indicative of p>0.01 Four grouped symbols are indicative of p<0.0001. Data show mean ± SEM.
65
Future Directions
My data provide novel insights into the physiological mechanisms by which the association of
DDR1 with NMIIA is regulated. In particular, I have discovered a novel function for MRIP in
non-muscle cells, which has previously been studied primarily in smooth muscle. The new data
show that MRIP is a regulator of tractional remodeling of collagen via DDR1 and NMIIA. Part
of my work has shown that there is an association between the expression of DDR1 and the
ability of cells to create adhesions with fibrillar collagen, which resist shear forces. Future
experiments could be conducted to directly quantify the adhesive forces of DDR1 adhesions to-
collagen. These experiments could employ the use of an assay whereby GD25 cells adhering to
magnetic collagen-coated beads are exposed to magnetic fields of varying known intensities,
which could then be used to quantify adhesive forces.
My work has also shown that DDR1, NMIIA, F-actin and MRIP co-localize in a
collagen-dependent manner. Because the assembly and disassembly of adhesion complexes are
fundamentally dynamic processes, using live-cell imaging in conjunction with GFP-tagged
proteins would help to uncover dynamic interactions and the timeframes over which their
association takes place.
Interestingly, my work has shown that knockout of MRIP by CRISPR strongly augments
the ability of epithelial cells to contract collagen gels and align fibrillar collagen yet the opposite
is observed for collagen compaction. These results suggest that compaction and alignment are
two distinct processes which may have distinct molecular regulators. Furthermore, this provides
insight into the dynamic nature of DDR1/NMIIA adhesions in the remodeling of collagen by
traction. These physiological mechanisms could suggest some sort of paralysis of the cell,
whereby the cell is able to form extensions which adhere to fibrillar collagen via DDR1, align
66
fibers using cytoskeletally-generated forces via NMIIA but are then no longer able to repeat this
process as the dynamic molecular machinery has been perturbed and forced to remain engaged.
Using live-cell imaging of MRIP knockout cells remodeling fibrillar collagen would shed light
on these phenomena.
This work has confirmed that MRIP expression is indirectly associated with regulating
the phosphorylation of MLC at S19 but I have also shown here that MRIP appears to influence
the phosphorylation state, and therefore the activation, of DDR1; this has not been observed
before. Using imaging of immunostained cells for phosphorylated DDR1 and knockout of MRIP
would provide insight into the dynamic nature of this phenomenon and the timeframes over
which they occur.
I have also shown that MRIP co-immunoprecipitates with DDR1, which provides
evidence that the two may be physically interacting with one another. It would therefore be
beneficial to define the domains, either on DDR1 or MRIP, which are responsible for this
interaction or look at whether their interaction is dependent upon MRIP interacting with NMIIA
by altering its binding sites with myosin phosphatase and NMIIA.
Conclusions
In conclusion my work has given insight into a novel aspect of the physiological mechanisms
involved in regulating the interaction of DDR1 with NMIIA (pictured below). I found that cell
attachment to collagen promotes a time-dependent association between DDR1, NMIIA and
MRIP. I have shown that MRIP exhibits a time- and collagen-dependent association with actin
stress fibers. I have shown that MRIP knockout enhances both MLC phosphorylation at S19 and
DDR1 phosphorylation at Y792. Finally, I have shown that downregulation of DDR1, MRIP
67
knockout, inhibition of DDR1 autophosphorylation and inhibition of ROCK1 all strongly
affected collagen adhesion, contraction and remodeling.
Diagram 5. Physiological mechanism by which MRIP regulates DDR1/NMIIA adhesions in collagen remodeling
Collagen remodeling by traction is achieved by the contractile ability of NMIIA which becomes phosphorylated, and active, upon collagen-induced activation of DDR1. MRIP presents itself as a critical regulator within the loop which regulates Serine 19 phosphorylation on NMIIA’s RLC. When MRIP is knocked out the cycle is impaired and Serine 19 remains phosphorylated and NMIIA remains contracted.
68
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