Oligomeric Status of Discoidin Domain Receptor Modulates Collagen Binding, Mechanics, and Receptor Phosphorylation

Dissertation

Presented in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy in the Graduate School of The Ohio State University

David Yeung, M.S.

Graduate Program in Biomedical Engineering

The Ohio State University

2018

Dissertation Committee:

Gunjan Agarwal, Advisor

Gregory Lafyatis

Heather Powell

David Yeung

2018

Abstract

Collagen type I is the most abundant extracellular matrix (ECM) protein found in vertebrates. A number of cell surface receptors and as well as glycoproteins and proteoglycans found in the ECM are known to interact with collagen. The clustering or oligomeric state of these proteins is emerging as an important factor in modulating their interaction with collagen. This thesis attempts to better understand the role of oligomeric status of the collagen receptors, Discoidin domain receptors 1 and 2 (DDR1 and DDR2), in their interaction with collagen. DDRs are widely expressed receptor tyrosine kinases which modulate cells signaling as well as ECM remodeling.

The first aim of this thesis (Chapter 2) was to understand how ligand binding impacts the clustering or oligomeric state of DDRs interaction. AFM imaging provided insight on how the extracellular domain (ECD) of DDR1 but not DDR2 was able to cluster upon collagen binding. Live cell assays where the murine osteoblastic cell line, MC3T3-E1, was transfected to express full length fluorescent DDR1 and DDR2 proteins showed that DDR1 clusters upon collagen stimulation whereas DDR2 does not. Further, receptor phosphorylation after collagen binding occurred in DDR1 clusters. On the other hand,

DDR2 phosphorylation was found in unique filamentous organizations of the protein. iii

Receptor phosphorylation for both DDR1 and DDR2 was observed several hours post collagen binding, consistent with earlier studies.

The second aim of this work (Chapter 3) was to investigate how the oligomeric state of

DDR ECD affects their binding to collagen and the structural and mechanical properties of the collagen network. Solid phase binding was used to observe how DDR oligomerization increased their binding affinity towards collagen. AFM and Confocal fluorescence microscopy imaging of collagen gels revealed that DDR2-Fc dimers, and to an even greater extent, DDR2-Fc oligomers reduced the fibril diameter and disrupted the network architecture. Results from turbidity assays supported oligomeric state of DDR2-Fc was a stronger inhibitor of collagen fibrillogenesis.

Mechanical properties of the collagen networks formed in the presence of dimeric vs. oligomeric DDR2 ECD were examined using macro and micro rheology (Chapter 4). The rheology performed via parallel plate and optical tweezers respectively showed how

DDR2-Fc oligomers increased the shear moduli of collagen gels as well as contributed to a unique strain stiffening pattern not seen in the other samples tested. Our novel active two particle optical tweezer technique showed better agreement with the parallel plate rheology compared to more traditional one particle methods. Finally, we summarize the importance of DDRs and receptor clustering in cell-matrix interactions and scope for future studies

(Chapter 5).

Dedication

To my father, Dr. Kwok Yeung, whose exemplary character and instruction has laid the

foundation for all of my own achievements.

To my mother, Rosita, for giving me the perspective to help me see beyond what was

right in front of me.

To my sister, Alyssa, for teaching me to be proud of who I am.

To John Downing, whose generosity and selflessness have taught me how lives can be

impacted through support and encouragement.

Acknowledgments

I have been fortunate to have had the support and assistance of many people throughout my doctoral studies.

I would like to thank David Chmielewski and Carolyn Wang for assisting me with my

AFM experiments. I would like to express gratitude towards Jack Wellmerling, Brent

Weiss and Nirvan Shankar for their indispensable help with culturing the cells for all the live cell work and protein purification. Without their assistance, the majority of this work would not have been possible. I also thank our collaborators Drs. Rafael Fridman (Wyane

Satte Univrsity) and Dr. Andrew Herr (University of Cincinnati) for DDR2-Fc plasmid and initial help with protein purification respectively.

I would like to thank Dr. Gregory Lafyatis for his guidance with developing the two- particle optical tweezer technique. David Gutschick also played a large role in these experiments and assisted in the data reduction for the microrheology assays. Dr. Peter

Anderson’s knowledge of material mechanics was foundational for the microrheology project and technique development. Initial help in optical tweezer instrumentation and training was performed in conjunction with Tyler Heisler-Taylor. vi

I would also like to thank our collaborators who helped me gather the macrorheology data.

The gel contraction experiments were done in Dr. Heather Powell’s lab, and the parallel plate rheology was performed using the equipment in the Dr. Matthew Reilly’s lab, with the help of his student Archie Tram.

Many of the essential laboratory techniques and requisite knowledge for my project was imparted via my senior doctoral students, Jeff Tonniges and Tanya Nocera along with post- doc Angie Blissett.

Finally, I would like to specially acknowledge my dissertation committee members, Drs.

Heather Powell and Gregory Lafyatis for their help throughout my PhD career. Special gratitude is reserved for my advisor, Dr. Gunjan Agarwal, for her sincere mentorship through the duration of my time at Ohio State.

A major part of this study was made possible by funding provided the NSF project 1201111 and AHA predoctoral fellowship 16PRE31160013.

vii

Vita

May 2007 ...... Penn High School

May 2011 ...... B.S. Biomedical Engineering,

Purdue University

August 2017 ...... M.S. Biomedical Engineering,

The Ohio State University

August 2017 to Present ...... Predoctoral Research Fellow,

Department of Biomedical

Engineering, The Ohio State

University

Publications

Yeung, D., Chmielewski, D., Mihai, C., and Agarwal, G. (2013). Oligomerization of DDR1

ECD affects receptor-ligand binding. J. Struct. Biol. 183, 495–500

Fields of Study

Major Field: Biomedical Engineering

viii

Table of Contents

Abstract ...... iii

Dedication ...... v

Acknowledgments...... vi

Vita ...... viii

Table of Contents ...... ix

List of Tables ...... xiii

List of Figures ...... xiv

Chapter 1: Introduction ...... 1

1.1 Overview ...... 1

1.2 Discoidin domain receptors (DDR1 and DDR2) ...... 3

1.2.1 Structural requirements of DDRs for collagen binding ...... 4

1.2.2 Structural requirements of collagen for DDR binding ...... 7

1.2.3 Structural changes upon DDR-collagen interaction ...... 8

1.2.3.2 Modulation of Collagen fibril structure via DDR binding...... 9

1.2.4. Aim of thesis ...... 10

Chapter 2: Effect of Collagen on the Oligomeric Status of DDR1 and DDR2 ...... 11

2.1 Aims and Rationale ...... 11

2.1.1 Background ...... 11

2.2 Methods ...... 13

2.2.1 Reagents...... 13

2.2.2 DDR expression vectors ...... 13

2.2.3 Expression and purification of recombinant DDR2 ECD proteins ...... 14

2.2.4 Western blot ...... 17

2.2.5 Atomic force microscopy ...... 17

2.2.6 Fluorescence microscopy of cells ...... 18

2.2.7 Immunocytochemistry ...... 19

2.3 Results ...... 20

2.3.1 Western blotting of recombinant proteins ...... 20

2.3.2 Oligomeric status of DDR1 and DDR2 ECD upon collagen binding ...... 21

2.3.3 Oligomeric Status of full length DDR1 and DDR2 upon collagen binding ..... 28

2.3.4 Role of oligomeric status of DDRs in receptor phosphorylation ...... 32

2.4 Discussion ...... 36

Chapter 3: Effect of Oligomeric Status of DDR ECD in Collagen Binding ...... 40

3.1 Aims and Rationale ...... 40

3.2 Methods ...... 41

3.2.1 Solid phase binding (ELISA) ...... 41

3.2.2 Turbidity Assay ...... 42

3.2.3 Confocal Reflectance Microscopy...... 42

3.2.4 Atomic Force Microscopy ...... 43

3.3 Results ...... 44

3.3.1 Effect of DDR oligomeric status on collagen binding and fibrillogenesis ...... 44

3.3.2 Effect of DDR2 ECD oligomeric status on collagen I gel microstructure ...... 49

Chapter 4: Effect of DDR2 Oligomeric Status on Collagen I Gel Mechanics ...... 56

4.1 Aims and Rationale ...... 56

4.2 Methods ...... 57

4.2.1 Collagen Gel Fabrication ...... 57

4.2.2 Microrheology ...... 58

4.2.3 Macrorheology...... 66

4.2.4 Gel contraction assay ...... 67

4.3 Results ...... 68

4.3.1 Viscoelastic nature of collagen gels ...... 68

4.3.2 Effect of DDRs on shear modulus of collagen I ...... 71

4.3.3 Effect of DDRs on gel contraction ...... 79

4.4 Discussion ...... 80

Chapter 5: Conclusions ...... 85

5.1 Comparison of oligomeric status of DDRs with other RTKS post ligand binding . 85

5.2 Comparison of oligomeric status of DDRs with other collagen binding proteins .. 89

5.3 Role of DDRs in matrix mechanics and cell behavior ...... 92

5.4 Future Directions and translational potential ...... 93

References ...... 95

xii

List of Tables

Table 1. AFM height measurements of DDR1 and 2 upon collagen binding...... 28

Table 2. Parameters describing structural properties of collagen gels ...... 52

Table 3. Parameters describing heterogeneity and mechanical properties of collagen gels

...... 84

Table 4. Comparison of Receptor Tyrosine Kinases ...... 87

xiii

List of Figures

Figure 1. Schematic of collagen receptors ...... 2

Figure 2. Schematic of DDR1a and DDR2 ...... 5

Figure 3. Schematic of DDR constructs used for transfection ...... 14

Figure 4. Western blot of DDR2-Fc and DDR2-V5 ...... 21

Figure 5. AFM images of DDR1-Fc binding to collagen ...... 24

Figure 6. AFM images of DDR2-Fc and DDR2-V5 binding to collagen ...... 27

Figure 7. Live Cell fluorescent images of DDR1 and DDR2 after collagen binding ...... 30

Figure 8. Immunocytochemistry of DDR1 after collagen binding ...... 33

Figure 9. Immunocytochemistry of DDR2 after collagen binding ...... 35

Figure 10. Solid phase binding of dimeric and oligomeric DDR1 to collagen...... 45

Figure 11. Solid phase bindng of mono-, di-, and oligomeric DDR2 to collagen ...... 47

Figure 12. Collagen fibrillogenesis assay with DDR2...... 48

Figure 13. Confocal and AFM images of collagen fibrils with DDR2 ...... 50

Figure 14. Quantitative measurements of collagen gels with DDR2 ...... 51

Figure 15. In-phase power spectrum of beads ...... 60

Figure 16. Trap linearity measurements ...... 62

Figure 17. Schematic of bead displacment ...... 63 xiv

Figure 18. Estimation of viscous vs elastic response ...... 69

Figure 19. Collagen gel moduli over strain sweep ...... 70

Figure 20. Collagen gel moduli over frequency sweep ...... 71

Figure 21. 1P moduli varying Poisson's ratio ...... 72

Figure 22. Comparison of 1P and macro rheology ...... 73

Figure 23. 2P microrheology moduli ...... 76

Figure 24. Reciprocal response from 2P measurements ...... 77

Figure 25. Poisson's ratio from 2P measurements ...... 78

Figure 26. Collagen gel area after contraction by cells ...... 80

Chapter 1: Introduction

1.1 Overview

Collagen type 1 is the most abundant extracellular matrix (ECM) protein in mammalian tissues. Interaction of collagen with various biomolecules is important for ECM structure and function as well as for cell-matrix interactions. Not only do collagen fibrils serve as a physical support to cells and regulate biomechanical properties of the underlying tissue, they are also responsible for mechanotransduction events mediated by cells sensing the collagen matrix. The biomolecules that interact with collagen I can broadly classified into glycoproteins, proteoglycans, glycosaminoglycans and cell-surface receptors.

There are four different types of cell-surface collagen receptors as seen in Figure 1. Among these, integrins and DDRs are the most ubiquitously expressed collagen receptors. This thesis focusses on DDRs and their role in regulating collagen biomechanics as well as cell- matrix interactions.

Figure 1. Schematic of collagen receptors adapted from Leitinger 2011 [1]. The integrins which bind to collagen are heterodimers consisting of an α and the β1 subunit. The discoidin domain receptors are homodimers and there are two members of this family.

Glycoprotein VI is also a homo-dimer interspersed with a FcRg. Both DDRs and GPVI consist of two domains in their extracellular regions. The Leukocyte associated immunoglobulin receptor 1 (LAIR-1) is a monomer with just one IG domain. This research was originally published in Annual Reviews [1]. Reprinted with permission.

1.2 Discoidin domain receptors (DDR1 and DDR2)

Discoidin domain receptors, namely DDR1 and DDR2, are part of the family of receptor tyrosine kinases (RTKs). DDRs exist as pre-formed dimers anchored to the cell membrane before binding to their ligand, collagen (including type I) [2]–[4]. Both DDRs are expressed in a wide variety of tissues. DDR1 is expressed in the developing nervous system [5], [6]. as well as the brain, lungs, spleen, kidney and placenta [7]–[11]. DDR2 tends to be found in the cells of connective tissue [12], and its mRNA has been found in tissue such as the cardiac and skeletal muscle, kidney, and lungs. With regard to cell-types, DDR1 is primarily expressed in epithelial [12] and endothelial cells [13]. DDR1 has been found upregulated in breast and glioma cell lines [10], [14] as well as megakaryocytes [15] and peripheral blood mononuclear cells after cytokine stimulation [16]. On the other hand,

DDR2 is expressed in fibroblasts [17], dendritic cells when immature and upregulated in mature ones after TNF-α exposure [18].

Functionally DDRs are involved in cellular processes such as cell-adhesion, proliferation, and migration [19]–[21]. Binding of DDRs to collagen impacts matrix remodeling via modulation of collagen fibrillogenesis [22]. Activation of DDRs after binding collagen has also been implicated in upregulation of matrix metalloproteases More specifically, DDR1 has been shown to induce expression of MMP-1 [23], mediate the production of MMP-2 and 9 [24], [25], and influence the expression of MMP-7 [26]. Separately, DDR2 has been shown to play a role in the expression of MMP-1 [23], MMP-2 [27], MMP-8 [28], and

MMP-13 [29]. Consistent with their role in ECM remodeling and inflammation, DDR expression and activation has been linked to diseases such as cancer, atherosclerosis, osteoarthritis, and fibrosis [24], [30]–[33]. The role of DDRs in health and disease continues to be an active area of research and is being investigated through use of mice models, DDR inhibitors as well as clinical samples.

1.2.1 Structural requirements of DDRs for collagen binding

The Discoidin domain receptors are so named because their N-terminal extracellular domain (ECD) consists of Discoidin (DS) and Discoidin-like (DS-like) domains which are largely conserved between DDR1 and 2. While the DS and DS-like domains are quite different in composition, they are both similarly sized and comprised of eight β-strands forming a barrel structure in both DDRs [34]–[36]. Between these domains and the transmembrane region, resides the extracellular juxtamembrane (JM) domain which is uniquely long amongst other proteins of the RTK family [37]. Both DDR1 and DDR2 are glycosylated proteins. The DS-like domains (Figure 2) contain a conserved calcium binding site [35] which has been shown to be modulate N-glycosylation in the DDRs and their subsequent membrane trafficking [38]. DDR1 has additional glycosylation sites, four

O-glycosylation sites (two on the DS-like domain, and two on the JM domain) and two N- glycosylation sites both on the JM domain. DDR2 also has four O-glycosylation sites (three on the DS-like domain and one on the JM domain) and one N-glycosylation site on the JM domain [39]–[41].

Figure 2. Schematic of DDR1a and DDR2 illustrating the different domains of the DDRs.

The extracellular domains are comprised of the DS, DS-like and extracellular juxtamembrane (EJXM) portions. The transmembrane (TM) domain separates the extracellular portion from the intracellular domain which contains the intracellular juxtamembrane (IJXM) and kinase domains. This research was originally published in the

Journal of Biological Chemistry [42]. Reprinted with permission.

The binding of DDRs to collagen takes place via their DS domains which have been shown to be necessary and sufficient for this interaction to occur [3], [36]. There are three highly conserved loops in these domains of DDR1 and 2 which when co-crystalized with the peptide containing the collagen binding motif helped reveal and confirm the notch where

DDR2 bind to collagen [3], [34], [36], [43]. The bottom and one side of this so-called trench are characterized by the non-polar residues Trp52, Thr56, Asn175, and Cys73- 5

Cys177. The other side is made of a salt bridge between Arg105-Glu113 and Asp69. These amino acids are conserved in the collagen-binding site of DDR1, but there are some differences in DDR2 which differentiate the types of collagen each receptor can recognize

[34].

A long-held belief about the RTK family of proteins is that the binding of their ligand results in the formation of a dimer at the cell surface [44], [45]. To this end, a unique aspect of the DDR subfamily is that they form ligand independent dimers which are present at the cell surface [2], [4], [46]. There are critical domains of the DDRs which have been implicated in their dimerization, particularly the transmembrane domains (TMD) [4], [47].

Using special DDR1 constructs which contained deletions of the extracellular domain or the intracellular domain, Noordeen and others were able to show that two potential dimerization motifs in the TMD existed. The most important of these was a leucine zipper which when mutated disrupted dimerization in truncated DDR1 proteins and activation in the full-length versions. This study also suggests that each segment of the protein (ECD,

TMD, Cytosolic domain) is likely involved in dimerization [4]. Additionally, it has previously been shown that the binding of DDRs to collagen requires dimerization of the

DS domain [3].

1.2.2 Structural requirements of collagen for DDR binding

Both DDR1 and DDR2 recognize the triple helical structure of collagen. For type I collagen, these triple helical monomers are comprised of three α-chains (two α1 and one α2) and are termed ‘heterotrimeric’. The individual monomers are approximately 300nm in length and

1.5nm wide [48]–[50]. While the main focus of this thesis is the binding of DDRs to collagen I, it is important to understand that DDR1 and 2 bind to fibrillar collagens types

2 and 3 as well as to other types of collagen. DDR1 has been shown to bind to the network forming collagens IV and VIII [24]. DDR2 binds to collagen X [51]. It had initially been shown that DDR2 binds to three sites on the collagen I triple helix [52] whereas DDR1 bound to overlapping collagen monomers [53]. Specific binding motifs for DDRs on the collagen triple helix have been identified through the use of collagen peptide toolkits by the Farndale lab, which encompass the entire collagenous domains of the homotrimeric collagens II and III. These data are what have enabled the mapping of the binding sites on the collagen monomers [34], [54], [55]. Both DDRs recognize the sequence of GVMGFO

(where O is hydroxyproline) on collagens I, II, III, and V [54], [55] with DDR2 having additional binding locations on collagens II and III [55].

Although DDRs bind to the monomeric collagen triple helix, it is not clear if DDRs bind to the triple helix present in the collagen fibrils. In the collagen fibril, the monomers are arranged in a specific quasihexagonal pattern [56] which obscures or exposes certain sites on the triple helix. The main point of interest arises because certain high affinity binding

7 sites identified for collagen binding proteins on the monomer could be hidden once incorporated in the collagen fibril [57]. There is still some debate as to how the fibrils are organized to exposed certain binding sites [58]–[60]. Previously, Orgel et al. had assembled a ranking of collagen binding sites based on their accessibility from the surface of the fibril

[57] as well calculated the solvent accessible surface area (SASA) for the matrix metalloprotease (MMP) cleavage site (Perumal 2008). More recently, Hoop et al. have performed a similar set of SASA calculations to understand accessibility of other collagen binding proteins and compared two different fibril organization [61]. Their computations also serve to illustrate that many of the known binding site on the collagen monomer are obscured when in the fibrillar state including the DDR binding motif.

1.2.3 Structural changes upon DDR-collagen interaction

Since ultimately, DDRs are RTK and hence signaling proteins, much research strives to understand this intimate relationship between the ability of these receptors for ligand binding and subsequent phosphorylation. The DDRs have a characteristically slow time to achieve phosphorylation, compared to other members in the RTK family, requiring 60 to

90 minutes after collagen stimulation [62], [63]. The reasons behind this delayed activation of DDRs are not yet understood. In this regard, it is being investigated how binding of

DDRs to collagen results in changes not only to the configuration of the DDRs but also to the collagen structure.

For both DDR1 and 2, it is understood that dimerization [3] and further oligomerization enhances their binding to collagen [52], [53], [64]. There are critical domains of the DDRs which have been implicated in their dimerization, particularly the transmembrane domains

(TMD) [4], [47] as well as the N-glycosylation sites in their ECD (Asn211 or Asn213 of

DDR1 and DDR2 respectively) [41], [65]. A conformational change has not been found in the ligand bound versus unbound state of the DDR1 DS domains [34], [36], [37]. It has been shown that DDR1 undergoes clustering at the cell surface after binding to collagen

[2], [66]. Both full length and kinase dead or depleted versions of DDR1 have the ability to cluster within the cell membrane after collagen binding [2], [65], [67], [68]. Another interesting observation on the mechanism of signal transduction across the cell membrane is that the long extracellular JM domain of DDR1 does not transmit the signal of ligand binding to its intracellular portion. Chimeric versions of the receptor consisting of a DDR1

ECD and an intracellular domain belonging to other proteins from the RTK family indicate that the mechanisms via which DDRs inciting their autophosphorylation may be unique

[66].

1.2.3.2 Modulation of Collagen fibril structure via DDR binding

A unique aspect of DDRs in receptor-ligand interaction is that the state of the ligand also becomes altered. With the use of high resolution microscopy techniques such as atomic force microscopy (AFM) and transmission electron microscopy (TEM), it has been shown that the fibril diameters and D-periodicity of collagen fibrils were disrupted when using recombinant DDR ECD proteins or cells expressing DDR ECDs [53], [64], [69]. Collagen 9 fibrils forms in the presence of DDR2 exhibited decreased persistence length [70]. In terms of overall fibrillogenesis, in vitro turbidity assays showed that DDR2 ECD slowed down the fusion of collagen into fibrils while DDR1 ECD did not [2]. Collagen fibrils in the aorta of DDR1 KO mice also showed an increase in fibril diameter and enhanced depth of D- periodicity [71]. All of these data indicate that DDR ECDs binding to collagen have in impact on the fibril formation and structure.

1.2.4. Aim of thesis

The goal of this thesis is to understand the role of DDR oligomeric status on receptor/ligand function in two separate ways. Firstly, what is the link between receptor clustering and phosphorylation post-ligand binding for DDR1 and DDR2. Secondly, what is the impact of DDR oligomeric status on collagen binding and how it impacts the collagen network microarchitecture and mechanics. The results derived from this thesis will advance our fundamental understanding of the collagen receptors and their role in regulating ECM structure, function as well as cell-signaling.

Chapter 2: Effect of Collagen on the Oligomeric Status

of DDR1 and DDR2

2.1 Aims and Rationale

Oligomerization and/or clustering of cell surface receptors can play a crucial role in modulating receptor-ligand binding, receptor activation, and downstream signaling events as shown for several other members of the receptor tyrosine kinase family [45]. The aim of this study was to understand how ligand (collagen type 1) binding impacts the oligomeric status of DDR1 and DDR2 and its potential role in receptor phosphorylation.

To accomplish this aim, we employed single molecule atomic force microscopy on purified proteins, live cell fluorescence microscopy and immuno-cytochemistry approaches.

2.1.1 Background

The three major structural domains, which can have a putative role in the oligomeric state of a receptor are its extracellular (ECD), transmembrane (TMD), and intracellular (ICD) 11 domains (Figure 2). The specific sites in DDR1 responsible for receptor dimerization have been described to be the leucine zipper motif in the transmembrane domain [4] and the cysteine residues in the JM region of the DDR1 extracellular domain (ECD) [46]. While it is speculated that regions in DDR1 ECD may contribute to receptor dimerization [35], the role of the ECD in receptor oligomerization is not completely understood. In vitro work by us and others revealed that high affinity interaction with collagen requires dimerization and/or pre-oligomerization of DDR1 [3], [46], [72]. It has also been reported that a significant percentage of the DDR1 population forms ligand independent dimers on the cell-surface [2], [4], [46].

The role of oligomerization and/or clustering of DDR2, in mediating interactions with collagen is even less well understood. Current data show that in DDR2, like in DDR1, (i) dimerization [3] and higher-order oligomerization of the DDR2 ECD [52], [64] enhances its binding to collagen; (ii) in cells, DDR2 exists as a constitutive noncovalent homodimer

[4], and like DDR1, the TMD of DDR2 displays a high propensity to self-interact [47]. A juxtamembrane segment in the ICD of DDR2 has also been shown to control receptor dimerization, and thereby regulate collagen-dependent activation[73]. However, unlike

DDR1, studies on the oligomeric state of DDR2 post-ligand binding are lacking.

2.2 Methods

2.2.1 Reagents

Fc-tagged ECDs of human DDR1 and TrkB were purchased as recombinant protein from

R&D Biochemicals, MN and reconstituted in sterile phosphate buffered saline (PBS) at a stock concentration of 100µg/ml. Bovine dermal collagen type I was obtained from

Advanced BioMatrix. Mouse monoclonal anti-DDR1 (against ECD) was from R&D

Biochemicals, MN. Anti-Fc antibody was from Jackson Immunoresearch, (West Grove,

PA). Anti-mouse and anti-goat IgG horseradish–peroxidase-conjugated antibodies were obtained from Santa Cruz Biotech. Dynasore, an inhibitor for dynamin mediated endocytosis (Macia et al., 2006) was purchased from Sigma–Aldrich, St. Louis, MO.

Glass-bottom culture dishes for live cell microscopy were obtained from MatTek

Glassware (Ashland, MA).

2.2.2 DDR expression vectors

For expression of recombinant DDR2 ECDs, we used a DDR2-V5-His construct comprising of the entire mouse DDR2 ECD tagged with a V5 epitope and 6His at the C- terminus as has been described earlier [69], and a DDR2-Fc, consisting of human DDR2

ECD harboring an IgG2 Fc fragment at the C-terminal end cloned into the pcDNA3.1/Myc-

His expression vector as described for DDR1-Fc [65] (Figure 3). For expression of full-

13 length DDRs in cells, we purchased an expression vector containing the human DDR2 cDNA tagged with GFP at the C-terminus from Origene. An expression vector encoding mouse DDR1b tagged with YFP was used as described previously [2].

Figure 3. Schematic diagram showing design of human DDR1-YFP, DDR2-GFP, DDR1-

Fc, DDR2-Fc and mouse DDR2-V5-His constructs. (Manuscript submitted).

2.2.3 Expression and purification of recombinant DDR2 ECD proteins

To obtain DDR2 ECDs displaying different oligomeric states, we used DDR2 ECD constructs in which the C-terminal region of the ECD was fused with either the Fc portion of the IgG molecule, to generate DDR2-Fc or with a V5-His tag, to generate DDR2-V5-

His (Figure 3). The Fc portion of the IgG molecule is known to undergo spontaneous 14 dimerization and to generate oligomeric species in the presence of anti-Fc antibodies. Thus, the recombinant DDR2-Fc ECD fusion protein (DDR2-Fc) is expected to display a dimeric state or allow for the antibody (anti-Fc)-induced oligomeric forms. On the other hand, the recombinant DDR2-V5-His ECD protein is expected to display a monomeric form.

The expression of the recombinant proteins was performed in human kidney HEK293T cells. Briefly, cells were seeded on collagen-coated 100-mm dishes and cultured in

Dulbecco’s Modified Eagle Medium supplemented with 10% fetal bovine serum (FBS) and 1% Penicillin/Streptomycin. Cells grown to 75% confluence were transiently transfected with plasmids encoding either human DDR2-Fc or mouse DDR2-V5 cDNA, using TransIT LT-1 transfection agent (Mirus Bio LLC), according to the manufacturer’s protocol. Twenty-four hours after transfection, the cells were re-fed with fresh serum-free

DMEM. The cells were then cultured for 2-3 days in serum-free DMEM and the conditioned media were collected and replaced with fresh serum-free DMEM, which was collected after another 2-3 days. The media were centrifuged (200g, 5 min) to remove cell debris and stored at 4° C until use.

For purification of DDR2-V5-His protein, the harvested media were dialyzed overnight

(12-14 kDa MWCO) at 4oC in 20 mM Tris-HCl, pH 7.4 with 300 mM NaCl. Thereafter, the media were loaded into a 5 ml His-trap column pre-equilibrated with 20 mM Tris-HCl pH 7.4, 300 mM NaCl. The column was then washed with equilibrium buffer, and the bound protein was eluted using a linear gradient of imidazole in a GE Amersham FPLC system. Fractions eluted between 120-150 mM imidazole were analyzed using SDS-PAGE 15 followed by Coomasie Blue staining and/or Western blotting to detect purified DDR2-V5-

His. Selected fractions were pooled, and subjected to a buffer exchange using a

Superdex200 gel filtration column equilibrated in TBS (20 mM Tris –HCl, pH 7.4, 150 mM NaCl). DDR2-Fc was purified using a procedure very similar to that used for human

DDR1b-Fc, as described [65]. Briefly, the harvested media were dialyzed in a solution of

25 mM NaH2PO4(pH 7.5) and 150 mM NaCl for 24 hours, and then loaded onto a 5 ml

HiTrap Protein A Column (GE Healthcare Life Sciences) pre-equilibrated with 25 mM

NaH2PO4(pH 7.0) and 150 mM NaCl. The column was then washed with 25 mM

NaH2PO4 (pH 7.0) and 150 mM NaCl and the bound protein was eluted using glycine buffer (pH 2.5). The eluate fractions (2 ml) were collected into tubes containing 200 μl of

1M Tris pH 8.0 and resolved by SDS-PAGE followed by Coomasie Blue staining and/or

Western blotting. The fractions containing DDR2-Fc were pooled and dialyzed against

PBS (pH 7.5). Purified proteins were concentrated using an Amicom Ultra Centrifugal

Filter Device (Millipore) with a 10k MWCO, and the protein concentrations were determined using absorbance at 280 nm. The monomeric/dimeric state of the recombinant proteins was determined by SDS-PAGE under reducing and non-reducing conditions followed by Western blotting, using antibodies against the DDR2 ECD or the epitope tag

(anti-Fc or anti V5), as previously described [64].

2.2.4 Western blot

To determine the molecular mass of the purified fusion proteins, SDS-PAGE was performed using 4–12% (w/v) NuPAGE ® Novex Bis-Tris Gels from Invitrogen

(Carlsbad, California). The proteins were diluted (to 10 ng) in NuPage LDS sample buffer

(Invitrogen) containing 141 mM Tris base, 2% LDS, 10% (v/v) glycerol, 0.51 mM EDTA,

0.22 mM SERVA® Blue G250 and 0.175 mM Phenol Red. Following SDS-PAGE, the proteins were transferred onto nitrocellulose membrane (Invitrogen) and blocked in TBS-

Tween buffer (20 mM Tris (pH 7.4–7.6), 0.5 M NaCl, 0.05% Tween) with 5% (w/v) milk.

The membranes were then incubated overnight in TBS Tween, 1% milk in the presence of

0.5μg/ml of either anti-Fc, anti-V5, or anti-DDR2 antibodies at 4 °C. The next day, the membranes were incubated with appropriate anti-Ig horseradish peroxidase (Santa Cruz,

CA) and detection was done using enhanced chemiluminescence (Amersham Biosciences).

For accurate determination of the molecular mass, protein samples were run on similar gels with BenchMark Protein Ladder (Invitrogen) and stained with Coomassie type stain, Safe

Stain (Invitrogen). Protein samples were also run under reducing conditions by adding β-

Mercaptoethanol (Sigma) to the samples before electrophoresis.

2.2.5 Atomic force microscopy

Collagen (1.0 µg/ml) was mixed with recombinant dimeric DDR1-Fc (0.2 µg/ml), monomeric DDR2-V5-His, or dimeric DDR2-Fc proteins (1.0μg/ml DDR2-V5-His, or

17 dimeric DDR2-Fc) in ice cold PBS and incubated at 4 C. As a control DDRs or collagen alone samples were also incubated in PBS under similar conditions. At specific time points

(0, 1, 2, 4 and 24 h), the samples were aliquoted onto chilled and freshly cleaved mica substrates, incubated for 5 min, washed and air dried and subjected to AFM imaging using the Multimode AFM (Digital Instruments, Santa Barbara, CA). AFM imaging was performed in tapping mode in ambient air using NSC15 cantilevers (Micromasch, Estonia) with a nominal spring constant of 40 Nm-1. Both height and amplitude images were recorded at 512 lines per scan direction. Topographic heights of DDR1-Fc in samples with or without collagen were measured from AFM images, by the section analysis feature of the Nanoscope software. At least n = 50 particles were analyzed per sample (except for

DDR2-V5-His particle bound to collagen for which n=25 due to a low number of binding events). Student’s unpaired two-tailed t-test was used to determine statistically significant differences across samples. A p<0.05 was considered significant.

2.2.6 Fluorescence microscopy of cells

Mouse osteoblast MC3T3-E1 cells obtained from ATCC were cultured in modified minimal essential medium alpha (Gibco) with 50 μg/ml ascorbic acid supplemented with

10% fetal bovine serum and antibiotics, as described earlier[2]. Cells were seeded on glass bottom culture dishes (MatTek Glassware, Ashland, MA), for live cell imaging and on polylysine-coated cover-slips for immuno-cytochemistry (ICC). Transient transfection with DDR2-GFP or DDR1b-YFP cDNA constructs was performed using TransIT LT-1

18 transfection reagent (Mirus Bio LLC). As a control, a set of cells were non-transfected.

Forty-eight hours after transfection, the cells were serum starved for ~ 12 hours and thereafter stimulated with collagen type 1 (20 µg/ml). For live cell imaging, the cells in glass-bottom dishes were imaged 48 hours after transfection, before and after collagen stimulation at times indicated.

2.2.7 Immunocytochemistry

Immunocytochemistry (ICC) was performed to evaluate receptor phosphorylation in cells expressing DDR1b-YFP or DDR2-GFP. Forty-eight hours after transfection, the cells were serum starved for ~ 12 hours and thereafter stimulated with collagen type 1 (20 µg/ml). At specific times (0, 1, 4 and 24 hrs) after collagen stimulation the cells were washed (2 times in PBS) and thereafter fixed in 4% paraformaldehyde (in PBS) for 10 minutes at room temperature. Thereafter the cells were washed (3 times in PBS) and permeabilized using

0.05% Triton X (in 2% BSA with PBS) for 5 min at room temperature. After subsequent washing, the cells were blocked with 2% BSA in PBST (PBS with 0.1% tween) for 1 hr at room temperature and then incubated overnight with primary antibody (anti-DDR1- phospho or anti-DDR2-phospho) as indicated. Thereafter, the cells were washed, incubated with secondary antibodies (in 2% BSA in PBST) for 1 hr at room temperature, washed and mounted onto glass slides using Sigma mounting media containing DAPI nuclear stain. All images were acquired using on a Zeiss Axiovert light microscope equipped with appropriate filter cubes for YFP, GFP, TRITC and DAPI fluorescence. Wide field

19 fluorescence images were acquired using a 63X water immersion objective lens and a

Hamamatsu camera coupled to the microscope as described earlier [2].

2.3 Results

2.3.1 Western blotting of recombinant proteins

The monomeric/oligomeric state of DDR2-V5-His and DDR2-Fc were confirmed by

Western blotting under reducing and non-reducing conditions (Figure 4). DDR2-V5-His exhibits a relative molecular mass of ~60 kDa under both reducing and non-reducing conditions, consistent with this protein being in a monomeric state. In contrast, DDR2-Fc displays a molecular mass (~190 kDa) when resolved under non-reducing conditions as compared to ~90 kDa under reducing conditions, characteristic of a Fc-tagged dimer [64].

The oligomeric state of DDR2-Fc, induced by the presence of anti-Fc antibodies, was determined earlier using size-exclusion chromatography [64].

Figure 4. SDS-PAGE and Western blotting of DDR2-V5-His and DDR2-Fc proteins under reducing or non-reducing conditions using anti-epitope or anti-DDR2 antibodies as indicated. (Manuscript submitted).

2.3.2 Oligomeric status of DDR1 and DDR2 ECD upon collagen binding

To examine the oligomeric state of DDR1 ECD upon collagen binding, we performed single-molecule studies using AFM on recombinant proteins. AFM is especially useful to resolve single-molecule interactions and oligomer formation as it can quantify particle sizes with sub-nm resolution and does not require labeling or fixing of biomolecules. AFM

21 imaging has been previously used to study oligomerization of various proteins such as

EGFR receptors[74], amyloid β protein[75], matrix protein M1[76], c rings of F-ATP synthases[77], and apoferritin[78] among others. For AFM studies, DDR1-Fc and DDR2-

Fc dimers as well as DDR2-V5 monomers were incubated with collagen in solution and thereafter immobilized on a mica surface. For comparison, recombinant proteins and collagen alone samples were also imaged using AFM.

Dimeric DDR1-Fc imaged as a globular protein with topographic height ranging from 1 to

4 nm, with an average height of 2.23 ± 0.6 nm. Incubation of DDR1-Fc alone for up to 4 h in solution did not change the size distribution or average height (2.14 ± 0.72) of particles

(Figure 5). Upon collagen binding a significant alteration in the morphology of DDR1-Fc was observed in a time-dependent manner. Very few binding events were observed in AFM images when DDR1-Fc and collagen samples were incubated for less than 1 h (data not shown). The number of binding events increased with incubation times >1 h. Particle size analysis from AFM images showed that the DDR1-Fc particles bound to collagen were not uniform in size and exhibited heterogeneity in both topographical height as well as lateral width. The topographic height of globular structures bound to collagen was analyzed for various time points as shown in Figure 5. Our AFM analysis confirmed that dimeric DDR1-

Fc binds to collagen as the majority of particles at the 1 h time point exhibited a topographic height corresponding to that of DDR1-Fc alone. However, a small percentage of particles exhibited a particle size >4 nm, suggesting the formation of DDR1 oligomers. At longer incubation times (>2 h) the particle size distribution shifted towards larger sizes and particles 10–15 nm in height were also observed binding to collagen. The particle size 22 distribution reached saturation after 4 h of incubation with no significant differences observed between 4 h and 24 h (data not shown) of incubation. The average particle size of collagen bound DDR1-Fc after 4 h of incubation was significantly different than the unbound DDR1-Fc (p<0.0001). Table 1 summarizes how the average height of DDR1-Fc increased upon collagen binding. Our results thus show that collagen binding led to oligomerization of the dimeric DDR1Fc post ligand-binding.

Figure 5. DDR1-Fc oligomerizes upon collagen binding. AFM height images and topographic height measurements of pure DDR1–ECD (DDR1-Fc) (0.2lg/ml) without and

24 with incubation with collagen type 1 for times indicated. DDR1-Fc images as a globular protein 2–4 nm in size. Collagen binding resulted in oligomerization of DDR1-Fc (which can be seen as white globular structures binding to filaments of collagen). Quantitative analysis of particle height of DDR1-Fc bound to collagen is presented at various time points. Collagen-bound DDR1-Fc exhibited a significant increase in particle height as compared to DDR1-Fc alone [79]. This research was originally published in the Journal of

Structural Biology [79]. Reprinted with permission.

We next performed single-molecule AFM imaging and analysis of monomeric DDR2-V5-

His and dimeric DDR2-Fc before and after binding to collagen, using a protocol similar to that used for DDR1-Fc. As shown in the AFM images of Figure 6, in the absence of collagen I, both DDR2-V5-His and DDR2-Fc imaged as a globular protein with a single lobe. AFM height measurements revealed that DDR2-Fc was ~ 0.2 nm larger in size

(p<0.0001) when compared to DDR2-V5-His (Table 1), consistent with the SDS-PAGE results in Figure 1. Upon incubation with collagen I, globular particles binding to collagen filaments could be easily identified in AFM images. Quantitative analysis of AFM images revealed that both the monomeric and dimeric proteins exhibited an increase in particle height upon collagen binding when compared to their respective unbound states (Table 1, p<0.0001) when measured with respect to the underlying mica substrate. However, unlike

DDR1-Fc which formed clusters over two-fold in size upon collagen binding, the increase in height of DDR2-ECD proteins bound to collagen was only ~ 1 nm. To determine whether the increase in the protein size in the presence of collagen I represented an additive effect caused by the collagen filaments, we also determined the height of bound DDR2 proteins 25 with respect to collagen filament. As shown in Table 1, both DDR2-V5-His and DDR2-Fc when measured with respect to its bound collagen displayed a height similar to that of unbound proteins (p>0.3), indicating that the collagen values accounted for the increase in height observed for the bound proteins. Thus, the size of the DDR2 monomers or dimers did not change significantly upon binding to collagen I, and unlike DDR1-Fc[79], DDR2

ECD did not form clusters or higher-order oligomers post ligand binding.

Figure 6. AFM height images of monomeric DDR2-V5-His and dimeric DDR2-Fc before and after binding to collagen as indicated. DDR2-V5-His and DDR2-Fc particles bound to 27 collagen are indicated by black and white arrows respectively. Particle size distribution and average sizes are indicated in the accompanying histograms and Table 1. (Manuscript submitted).

Relative to substrate (mica) Relative to collagen

-collagen +collagen +collagen

DDR2-V5-His 0.92 ± 0.26 1.53 ± 0.25 * 0.97 ± 0.22

DDR2-Fc 1.17 ± 0.40 2.01 ± 0.49 * 1.24 ± 0.48

DDR1-Fc 2.14 ± 0.72 4.88 ± 2.76 * N/A

Table 1. Topographic height (in nm) of DDR1 and DDR2 ECD proteins from AFM images.

Asterisks (*) indicate a significant difference (p<0.0001) in the collagen bound height of each protein to their respective unbound height.

2.3.3 Oligomeric Status of full length DDR1 and DDR2 upon collagen binding

To examine the effect of collagen on full length receptor distribution and/or clustering we used mouse osteoblasts (MC3T3-E1 cells) transiently transfected with vectors encoding the full length DDR2-GFP or DDR1-YFP[2]. Extensive cell spreading and the associated low height profile of these cells make them well-suited for wide-field fluorescence microscopy. Receptor distribution before and after collagen stimulation was followed by

28 fluorescence microscopy in real time in live cells. As shown in Figure 7, both DDR1-YFP and DDR2-GFP exhibited a uniform distribution of fluorescence on the cell surface before collagen stimulation. The control un-transfected cells showed no signal (data not shown).

Figure 7. Live cell imaging of DDR1b-YFP and DDR2-GFP transfected MC-3T3 cells using wide-field fluorescence microscopy, before and after addition of collagen (C) as 30 indicated. DDR1b and DDR2-GFP exhibit a uniform distribution on the cell surface before collagen stimulation. DDR1b-YFP results in cluster formation upon collagen stimulation whereas DDR2-GFP does not. Presence of filamentous structures was observed in DDR2-

GFP expressing cells at prolonged (4 hr) collagen stimulation time. Insets show selected regions (11.7 µm in size) which have been magnified to visualize receptor clustering. The location of these selected regions on the cell surface is indicated by dashed boxes.

(Manuscript submitted).

However, within a few minutes after the addition of collagen I, DDR1-YFP displayed punctuate and noticeable redistribution and clustering on the cell surface, resulting in

‘squiggle’ like structures. At prolonged stimulation, a more heterogenous distribution of size and shape of DDR1-YFP clusters was observed which consisted of both squiggles and globular structures. In contrast to DDR1-YFP, DDR2-GFP showed no obvious signs of receptor redistribution or clustering in response to collagen. However, at prolonged stimulation, DDR2-GFP expressing cells, revealed the formation of filamentous structures with a fraction of DDR2-GFP assembling along or associating with these filaments. Thus,

DDR1 and DDR2 exhibited differences in their clustering behavior upon collagen stimulation.

2.3.4 Role of oligomeric status of DDRs in receptor phosphorylation

To ascertain if the phosphorylation of DDRs was dependent on cluster formation, we examined spatial distribution of receptor phosphorylation in DDR1-YFP or DDR2-GFP expressing cells using immuno-staining with antibodies specific to phosphorylated DDR1 or DDR2. As shown in Figure 8b, DDR1 phosphorylation was co-localized with clusters present after ~4 hours of collagen stimulation. However, not all DDR1b-YFP clusters were phosphorylated at 4 hours and very little co-localization was present at earlier time points

(supporting information). DDR2 phosphorylation on the other hand co-localized with the filamentous structures formed after 4 hrs of collagen stimulation (Figure 9b) with very little basal phosphorylation in other parts of the cell or at earlier time points (supporting information). As observed for DDR1 clusters, not all filamentous structures co-localized with DDR2 phosphorylation. Thus, clustering of DDR1-YFP and assembly of DDR2-GFP into filamentous structures were the major sites of receptor phosphorylation.

Figure 8. Immuno-cytochemistry of DDR1b-YFP expressing cells to examine spatial distribution of receptor phosphorylation. Total receptor is indicated in YFP channel (green) while staining for phopho-DDR1 is shown by TRITC (red). Co-localization of total and phosphorylated receptor is shown in yellow. Blue represents nuclear (DAPI) staining. (a)

Little to no phosphorylation or co-localization signal was detected, 30 min after collagen stimulation despite the presence of DDR1b clusters. Insets show selected regions (11.7 µm 33 in size) which have been magnified to visualize receptor clustering and/or co-localization.

The location of these selected regions on the cell surface is indicated by dashed boxes. (b)

At prolonged collagen stimulation (4 hrs) a number of DDR1b clusters co-localized with phosphorylation. The bottom row consists of selected regions from various cells showing presence of clusters with or without phosphorylation. (Manuscript submitted).

Similar to these observations with DDR1b, little to no phosphorylation was observed for

DDR2-GFP at the early (30 min) time point (Figure 5a). After 4 hrs of collagen simulation, the bulk of phosphorylated DDR2 co-localized with the GFP positive filamentous structures (Figure 5b), with minimal presence of phosphorylated receptor in other parts of the cell. Indeed, not all filamentous GFP positive structures co-localized with phosphorylated DDR2. Thus, clustering of DDR1b-YFP and assembly of DDR2-GFP into filamentous structures were the major sites of collagen I-induced receptor phosphorylation in transfected MC3T3-E1 cells.

Figure 9. Immuno-cytochemistry of DDR2-GFP expressing cells to examine spatial distribution of receptor phosphorylation. Total receptor is indicated in GFP channel (green) while staining for phopho-DDR2 is shown by TRITC (red). Co-localization of total and phosphorylated receptor is shown in yellow. Blue represents nuclear (DAPI) staining. (a)

Little to no phosphorylation or co-localization signal was detected, 30 min after collagen stimulation. Insets show selected regions (11.7 µm in size) which have been magnified to 35 visualize receptor distribution and/or co-localization. (b) At prolonged collagen stimulation

(4 hrs) a fraction of DDR2-GFP was observed to associate or organize with filamentous structures. A fraction of these filamentous structures co-localized with DDR2 phosphorylation. The bottom row consists of selected regions from various cells showing presence of filamentous structures with or without phosphorylation. (Manuscript submitted).

2.4 Discussion

DDRs are type I membrane proteins in which their ECD is exposed to the extracellular milieu, ready to interact with collagens. On the membrane, DDRs are displayed as a mixture of monomeric and homodimeric forms, and thus are unique RTKs because they exist as inactive preformed non-covalent homodimers[4].

Using AFM, we show that the recombinant DDR1 ECD (which lacks the transmembrane and intracellular domains) undergoes oligomerization upon binding to collagen in vitro.

Although we could not determine the stoichiometry of DDR1 oligomers formed, our AFM results show that the mode value for DDR1-Fc size distribution before and after collagen binding was 2–3 and 4–5 nm respectively. Based on the previously determined topographic heights of antibody-induced oligomers of DDR1 [72], we estimate that ligand binding results in formation of tetramers, hexamers or octamers consisting of two to four DDR1-

Fc dimers. However, besides tetramers–octamers, higher order oligomers of DDR1-Fc

(10–15 nm in height) are also formed upon collagen binding.

Several putative sites in the Discoidin (DS) domain, DS-like domain and the JM region of

DDR1 ECD may mediate oligomerization of DDR1-Fc dimers. In a recent study, Carafoli et al. (2012) [35] have shown that monoclonal antibodies (mAbs) that bind to the DS-like domain of DDR1, inhibit collagen-induced receptor activation. They propose that mAbs prevent the proximity of the two DS-like domains and the JM regions in the collagen-bound, signaling, state of the DDR1 dimer. A conserved patch between the DS and DS-like domain is understood to mediate protomer contacts in the signaling DDR1 dimer, either by forming a direct DS-DS interface or by providing a secondary collagen-binding site. In addition,

Arg32 and Leu152 in the DS domain were also shown to mediate dimer formation in the crystal state and were required for DDR1 signaling, even though they are not part of the primary collagen-binding site. Thus far, soluble versions of monomeric DDR1 ECD have shown little [3] or reduced [43] binding to collagen in solid-phase binding assays. It remains to be investigated if monomeric DDR1 ECD can undergo ligand-induced oligomerization as elucidated for DDR1-Fc dimers.

A notable feature in our findings was the dissimilarity in the clustering ability of DDR1-

Fc vs. that of DDR2-Fc post ligand binding, which was evident in the results from both

AFM experiments and cell-based studies. While DDR1-Fc spontaneously clustered to form high-order structures upon collagen binding [79], no such feature was observed for DDR2-

Fc. Measurement of particle sizes from AFM images revealed that DDR2-Fc preserved its 37 size post-collagen binding. Furthermore, live-cell imaging showed that while DDR1b-YFP underwent a re-distribution and cluster formation after collagen stimulation [2], DDR2-

GFP maintained a homogenous distribution on the cell surface with no clustering. While we cannot completely rule out the possibility of contributions from the TMD or ICD domains of DDR2 in small cluster formation, which could not be resolved by wide-field light microscopy of cells, our results suggest that unlike DDR1, DDR2 is not able to organize into large clusters upon ligand binding. We thus elucidate that DDR2 ECD does not mediate receptor oligomerization post ligand binding.

Our results also provide insights into the spatiotemporal relationship between collagen I- induced DDR phosphorylation and clustering. Earlier study, from our laboratory showed that DDR1b-YFP undergoes rapid receptor clustering and internalization into endocytic vesicles after collagen stimulation [2]. Further, we demonstrated that the bulk of phosphorylated DDR1b is found within the intracellular compartment [65]. Consistent with these findings, we now demonstrate that DDR1b clustering precedes its collagen I-induced phosphorylation at Y513. Indeed, DDR1b phosphorylation primarily occurred on preformed clusters at prolonged collagen stimulation times. Studies have identified the tyrosine kinase, Src, to be important to achieving maximum phosphorylation in DDR2 by working on the tyrosines in the DDR2 activation loop which leads to further autophosphorylation [80], [81] This mechanism has also been implicated in DDR1 [82],

[83]. Since the bulk of the receptor is localized within the intracellular compartment as a consequence of ligand-induced endocytosis, it is tempting to speculate that DDR1 phosphorylation and signaling may also take place within the endosomal environment, as 38 demonstrated for other members of the RTK family [84], [85]. While this is possible for

DDR1, based on our data, this may not be the case for DDR2. Indeed, the spatial distribution of phosphorylated DDR2 in response to collagen was strikingly different from that of DDR1. At prolonged collagen stimulation times, DDR2 was found to associate with or assemble into filamentous structures resembling intermediate filaments in the cell cytoskeleton, and thus it did not display a pattern consistent with vesicular locales [86].

However, similar to DDR1 clusters, only a fraction of these filamentous structures co- localized with DDR2 phosphorylation, suggesting putative role(s) of intracellular factors in mediating DDR2 phosphorylation post-ligand binding.

Taken together our results elucidate how clustering of ECD (and likely other domains [4],

[68]) in DDR1 regulate receptor phosphorylation. Thus, whereas DDR1 may undergo phosphorylation in the absence of ligand-induced conformational changes, DDR2 phosphorylation is likely modulated by conformational change(s) accompanying ligand binding [34], followed by intracellular factors which dock the receptor at specific cellular site(s). Indeed, our data indicate that the subcellular locations associated with receptor phosphorylation appear to be distinct for DDR1 and DDR2. While DDR1b phosphorylation occurs on clusters (in endocytic vesicles) [2], DDR2 phosphorylation is localized to cytoskeletal filaments. Recent years have witnessed a number of intracellular kinase inhibitors for DDR1 and DDR2 [87]. Our results suggest that further investigations in identifying intracellular co-factors and subcellular-locations may aid in the design of specific and more effective inhibitors for DDR1 and DDR2.

Chapter 3: Effect of Oligomeric Status of DDR ECD in

Collagen Binding

3.1 Aims and Rationale

Independent studies have shown how the oligomeric state of DDR1 and DDR2 ECD enhances their collagen binding ability. However, a quantitative comparison of the monomeric, dimeric and oligomeric state in collagen binding has been lacking. To gain more insight into the role of the oligomeric state of the DDR ECD in its interaction with collagen, we utilized ECD constructs of DDR2 capable of displaying monomeric, dimeric or oligomeric form as well as similar DDR1 ECD constructs in its dimeric or oligomeric state. Solid phase binding assays and collagen turbidity measurements were used to ascertain collagen binding and fibrillogenesis. The microstructure of the collagen networks formed was evaluated using confocal reflectance and atomic force microscopy.

3.2 Methods

3.2.1 Solid phase binding (ELISA)

A monomeric solution of bovine dermal collagen type I in ice-cold PBS was immobilized in 96-well micro-titer plates by incubating the wells with 100µl of 25 µg/ml of collagen, overnight at 4oC. Thereafter, the plates were washed three times with 200 µl TBS (Tris

Buffered Saline) (Bio-Rad, Hercules, CA) containing 0.05% Tween 20(GE Healthcare,

Uppsala, Sweden), followed by blocking with 300 µl of 1% bovine serum albumin (Santa

Cruz Biotechnology, Santa Cruz, CA) with 0.05% tween overnight at 4 oC. The wells were washed again three times with TBS-tween and incubated with 100 µl of recombinant

DDR2-V5-His, DDR2-Fc, or DDR1-Fc (before and after pre-oligomerization) at concentrations ranging from 0 to 10 µg/ml, overnight at 4 oC. To induce oligomerization of DDR1-Fc and DDR2-Fc, the proteins were incubated individually with an equal mass of anti-Fc antibody in PBS at 4 oC overnight, as previously described [64]. To detect binding of recombinant proteins to collagen, the plates were washed and thereafter probed with the appropriate anti-DDR monoclonal antibodies, followed by washing and incubating with HRP conjugated secondary antibodies. Bound protein was detected by adding 100 µl of 3,3′,5,5′-tetramethylbenzidine to each well for 20 min at room temperature, and no light exposure. The reaction was stopped using 1N HCl (Sigma Aldrich, St. Louis,

MO) solution, and the absorbance was recorded at 450 nm using a spectrophotometer. All

41 experiments were performed at least three times. IC50 for binding was determined as described earlier [79].

3.2.2 Turbidity Assay

Collagen fibrillogenesis was assessed by measuring the turbidity of neutralized collagen I solutions with a rise in temperature. To this end, 250 µl of ice-cold collagen solutions (100

µg/ml) in PBS (pH 7.5) containing 20 µg/ml of recombinant DDR2-V5-His or DDR2-Fc proteins (before and after pre-oligomerization) were incubated in 96-well Costar clear polystyrene plates at 37 oC. A collagen solution with or without anti-Fc antibody (20 µg/ml) was used as a control. The absorbance values were monitored every hour at 405 nm for up to 6 hours using a spectrophotometer.

3.2.3 Confocal Reflectance Microscopy

Confocal reflectance microscopy on collagen gels devoid of beads was performed using an incident wavelength of 488 nm on an Olympus FV1000 inverted microscope with a water immersion 40x objective lens with NA 0.8. Stacks of n=26 slices were obtained from each sample type (with n=3 gels per sample) with a slice thickness of 1.17 µm. The stacks were processed with ImageJ in order to calculate the three-dimensional mesh size, ξ, between the collagen fibrils using the plugin BoneJ [88] which, after thresholding, fits the largest sphere possible in to each open space. The area fraction (Af) of collagen from each slice

42 was calculated by thresholding the images in ImageJ and analyzing the area occupied by collagen. These measurements were averaged over multiple slices of the entire sample to produce the final percentage. In addition, collagen density (ρ) was calculated by creating a maximum intensity projection of all stacks in a series followed by analyzing the intensity per 1µm2 area via ImageJ.

3.2.4 Atomic Force Microscopy

Collagen gels were prepared as described above except without the incorporation of beads.

Collagen gels were polymerized in a 48 well plate overnight in a humidified chamber at

37°C. After polymerization, the gels were washed 3 times with PBS to remove unbound proteins and thereafter fixed with 3% glutaraldehyde (in PBS) for 2 hours at room temperature. After fixation the gels were subjected to 5 washes, each of 20 minutes in distilled water at room temperature. Thereafter, the gels were gently released and removed from the wells, placed on to freshly cleaved mica and kept in a desiccator overnight.

Dehydrated collagen gels were imaged using a Multimode AFM equipped with a JV scanner and a Nanoscope IIIa controller (Bruker Inc) as described in our earlier studies[79].

AFM imaging was performed in tapping mode in ambient air using HQ NSC15 cantilevers

(Micromasch, Estonia) with a nominal spring constant of 40 Nm-1. Both height and amplitude images were recorded at 512 lines per scan direction with a scan speed of 2 lines per second. To avoid tip-convolution effects, the same probe was used to image all sample types. In addition, each sample was imaged using three new probes. Diameter of at least

43 n=25 collagen fibrils was ascertained from AFM height images using the section analysis feature of Nanoscope software.

3.3 Results

3.3.1 Effect of DDR oligomeric status on collagen binding and fibrillogenesis

To confirm the ability of DDR1 ECD to bind to collagen, dimeric DDR1-Fc or the control protein, TrkB-Fc were incubated over immobilized collagen in solid-phase binding assays and the bound protein detected using anti-Fc antibodies. As shown in Figure 10a, DDR1-

Fc specifically bound to collagen while the control protein TrkB-Fc showed no significant binding. To examine if pre-oligomerization of DDR1 ECD impacts its ability to bind to collagen, similar assays were performed by incubating DDR1-Fc before and after pre- oligomerization to collagen coated wells and the bound protein detected using anti-DDR1 antibodies. The DDR1-Fc dimers and antibody-induced oligomers of DDR1-Fc used in our samples had identical concentrations of DDR1 ECD. Our solid-phase binding assays showed a rapid saturation of the binding signal for oligomerized DDR1-Fc as compared to the dimeric samples (Figure 10b). The IC50 for binding of dimeric and oligomeric DDR1-

Fc to immobilized collagen type 1 was determined to be 553 ± 179 ng/ml (7.85 ± 2.54 nM)

44 and 176 ± 44.9 ng/ ml (2.5 ± 0.64 nM) respectively using the approach of Orosz and Ovádi

(2002).

Figure 10. Binding of DDR1-Fc to immobilized collagen type 1. (A) DDR1-Fc dimers bind to collagen but the control protein TrkB-Fc does not. The bound proteins were detected using anti-Fc antibodies. (B) Antibody (anti-Fc) induced oligomers of DDR1-Fc bound to collagen with a higher affinity than the dimers alone. The amount of DDR1-Fc

45 was identical in dimeric and oligomeric samples; bound proteins were detected using monoclonal anti-DDR1 antibodies.

The ability of the recombinant DDR2 ECDs to bind to immobilized collagen I was also evaluated using a similar approach. As shown in Figure 11, the binding of the DDR2 ECD to the immobilized collagen I was dependent on its oligomeric state. The antibody induced- oligomeric DDR2-Fc displayed the highest binding followed by the dimeric DDR2-Fc and the monomeric DDR2-V5-His form. The anti-Fc antibody alone showed no binding to collagen I (data not shown). Quantitative analyses showed that, at equal DDR2 ECD molar concentrations, the IC50 for the oligomeric DDR2-Fc was 32 ng/ml, about 10-fold lower than that of dimeric DDR2-Fc (310 ng/ml). The IC50 for DDR2-V5-His could not be determined as the binding curve did not reach saturation under the conditions tested.

Figure 11. Solid phase binding of DDR2 ECD proteins to immobilized collagen I as indicated. Binding was detected using antibodies against DDR2 ECD. (Manuscript submitted).

Collagen I fibrillogenesis can be induced in vitro by incubating monomeric chains of acid- solubilized collagen I in neutral pH at physiological temperature [64]. We therefore examined the extent of collagen I fibrillogenesis as a function of the oligomeric state of the

DDR2 ECD proteins. As shown in Figure 12, a solution of collagen I alone displayed a time-dependent increase in turbidity (absorbance) in neutralized solution, consistent with formation of fibrils. Under similar conditions, addition of oligomeric DDR2-Fc to the collagen I solution completely blocked fibrillogenesis, as determined by the lack of turbidity in the solution up to 5 hours. As a control, addition of the anti-Fc antibody to the collagen solution had no effect on turbidity, demonstrating that the effect observed on 47 fibrillogenesis was specific to the DDR2 ECD protein. Dimeric DDR2-Fc also inhibited fibrillogenesis albeit was only ~ 80% efficient as compared to the oligomeric form. In contrast, the monomeric DDR2-V5-His had a less pronounced effect on fibrillogenesis, with only ~35% inhibition at 5 hours. Taken together, these studies elucidate that the increasing the oligomeric state of the DDR2 ECD enhanced its ability for collagen I binding and inhibition of fibrillogenesis.

Figure 12. Inhibition of collagen I fibrillogenesis assessed using collagen turbidity measurements. DDR2 ECD proteins (20μg/ml) as indicated were incubated with 100 μg/ml of collagen type 1 in 96-well plate. (Manuscript submitted).

3.3.2 Effect of DDR2 ECD oligomeric status on collagen I gel microstructure

To evaluate how DDRs influence the structural changes in the collagen network, we examined the collagen gel microarchitecture using confocal reflectance microscopy (CRM)

(Figure 13a) consistent with our micro-mechanical characterization, CRM images of collagen gels show a heterogeneity in the distribution of collagen for decoron samples.

Quantitative analysis of CRM images was performed to ascertain local collagen density ()

3 in 1 μm voxels across each image, the mesh size, ξ, and the area fraction (Af) of collagen in each slice. As shown in Figure 14, BSA samples had a lower Af, consistent with their increase in measured [89]. AFM imaging of collagen gels revealed that collagen fibril diameter was unaffected in BSA samples whereas DDR2 samples had a significantly reduced fibril diameter ‘d’(Figure 13b). All values are listed in Table 2.

Figure 13. (a) Confocal reflectance microscopy (CRM) images of collagen gels formed with various NCPs as indicated. Inset shows CRM image of a bead inserted in collagen gel. 50

(b) AFM amplitude images of dehydrated collagen gels elucidating fibril morphology.

Fibril diameters were estimated from AFM height images. (Manuscript submitted).

Figure 14. Quantitative estimates of collagen density () distribution, mesh sizes () and area fraction (Af) ascertained from 3D stacks of CRM images. * denotes p<0.05.

(Manuscript submitted). 51

PBS BSA O. DDR2 DDR2 Parameters describing structural properties of collagen gels (average ± SD) 1 µm2 Intensity 0.55±0.09 0.52±0.11 0.57±.09 0.53±0.09 Distribution 1 µm2 Intensity 15.5% 21.6% 15.3% 17.9% Distribution CV Mesh size ξ (in μm) from 0.75±0.33 0.74±0.29 0.52±0.27 0.82±0.3 CRM Area fraction Af from 30.1±1.10 25.2±0.35 31.1±1.0 31.2±0.52 CRM Fibril dia. d (in nm) from 53.7±11.7 56.4±10.2 39.0±9.7 AFM A2: % of original gel area 20 18 1 20

Table 2. Parameters of different structural properties of collagen gels. (Manuscript submitted).

3.4 Discussion

Our results demonstrate that oligomerization of DDR1 ECD is crucial for high-affinity receptor–ligand binding. Previous studies by us [72] and others [3] have shown conflicting results where DDR1-Fc dimers failed to bind to collagen in one study and did bind in another. These discrepancies were likely due to differences in the techniques used (surface plasmon resonance (SPR) vs. solid-phase binding assays) for analyzing receptor binding.

In this study, by utilizing solid-phase binding assays we confirm that dimeric DDR1-Fc does indeed bind to collagen. The IC50 determined for binding of DDR1-Fc dimers (7.5 nM) to bovine dermal collagen type 1 in this study was comparable to that for rat tail collagen type 1 (10 nM) as reported earlier [3]. We also demonstrate that pre-

52 oligomerization of DDR1-Fc enhances its binding to collagen (IC50 for oligomers was three times lower than that for DDR1 dimers) consistent with our earlier SPR results [72].

Our results elucidate similarities and differences between the dimeric ECDs of DDR1 and

DDR2 in their interactions with their common ligand collagen I. Dimeric DDR1 and DDR2

ECD both bind to the triple helical collagen I and exhibit enhanced binding upon pre- oligomerization. However, as shown in chapter 2, only the ECD of DDR1 is capable of clustering upon ligand binding. Clustering of DDR1 ECD likely sequesters the available receptor domains, thereby reducing the number of binding events on the collagen triple helix as compared to DDR2 ECD. Consistent with this hypothesis, the IC50 for DDR1-Fc was much higher than that for DDR2-Fc in both the dimeric and the oligomeric forms.

Furthermore, the IC50 for oligomeric DDR1-Fc was only ~ 3 times lower than its dimer whereas the oligomeric DDR2-Fc form exhibited an IC50 ~ 10 times lower than its dimeric form, indicative of more frequent binding events for oligomeric DDR2-Fc. Clustering of receptor ECD can have important functional consequences, and in the case of DDRs may regulate the organization of the collagen matrix. Indeed, our previous studies indicate that the DDR2 ECD [64] is a stronger inhibitor of collagen I fibrillogenesis than DDR1 [72],

[90].

We also performed a more comprehensive evaluation of isolated ECDs of DDR2, capable of displaying monomeric, dimeric or oligomeric forms, on collagen binding in vitro as these recombinant proteins could be purified in-house. We found that oligomeric and dimeric DDR2 ECD species showed the highest affinity towards immobilized collagen I, 53 when compared to the monomeric form. These results are in agreement with earlier reports showing that a dimeric state of DDR2 ECD is required for high affinity binding to collagen

I [3] and oligomeric state of DDR2 ECD enhanced its binding to collagen [64]. Our results also help resolve the discrepancies due to the different binding assays utilized in these earlier reports. Further, pre-oligomerization of DDR2-Fc enhanced collagen binding in solid-phase binding assays in a manner similar to that observed for DDR1-Fc [79].

Conversely, monomeric DDR2-V5-His exhibited reduced binding, consistent with the findings for monomeric His-DS2 comprised only of the DDR2 Discoidin domain [3], [36], which is unable to oligomerize. It is important to note that in an earlier study, aminoterminal tagged His-DDR2 ECD was characterized to be a non-covalently linked dimer [3]. This difference in the oligomeric state of His-tagged DDR2 ECD could arise due to different sites for epitope tagging and/or differences in fractions collected during protein purification.

One possible explanation for differences in the binding ability of monomeric vs. dimeric forms of DDR2 ECD to collagen, could be that the monomeric form only binds to the primary GVMGFO site whereas dimeric (and oligomeric) DDR2 ECD bind to additional sites on the collagen triple helix. Our results from AFM imaging support this hypothesis because the number of binding events observed for dimeric DDR2-Fc were more frequent than those for monomeric DDR2-V5-His on the collagen triple helix, under identical experimental conditions. Our earlier AFM studies had elucidated three possible binding sites for DDR2-Fc oligomers on the collagen I triple helix [52]. Recent studies using col II and col III toolkit peptides have identified four additional DDR2 binding sites besides the 54 primary GVMGFO site [54], [55]. Molecular modeling [54] and X-ray crystallographic studies[34] have elucidated how monomeric and dimeric forms of the DDR2 ECD can bind to the GVMGFO site but no such studies exist for the additional DDR2 binding sites on the collagen triple helix. Although not discussed by the authors of these studies, it is interesting to note that in their toolkit studies, the various DDR2 ECD variants (His-DDR2,

DS2-Fc and DDR2-Fc) show different relative affinities to these additional binding sites

[54], [55]. For instance, while DDR2-Fc recognizes the toolkit peptide II-5 [55], this site is not recognized by DDR2-His or DS2-Fc [54]. Thus, it is likely that binding of dimeric

(and oligomeric) DDR2 ECD forms to additional sites on the collagen triple helix could account for their stronger inhibition of collagen fibrillogenesis when compared to the monomeric DDR2 ECD form.

The effect of binding of dimeric vs. oligomeric DDR2 ECD was also manifested in the resulting collagen network architecture. Our imaging from CRM showed the ability for collagen incubated with DDR2 ECD in its dimeric and oligomeric form to result in disrupted fibril morphology with barely resolvable fibrils consistent with previously known effects of DDR2 ECD on collagen fibrillogenesis. Oligomeric DDR2 resulted in a smaller mesh size. AFM results revealed a decrease in collagen fibril diameter with both dimeric and oligomeric DDR2. These structural features of collagen gels suggest that DDR2 ECD may enhance the stiffness of collagen gels.

Chapter 4: Effect of DDR2 Oligomeric Status on

Collagen I Gel Mechanics

4.1 Aims and Rationale

As shown in Chapter 3, the oligomeric status of DDR2-ECD impacted the collagen fibril and network microarchitecture. To understand how these structural changes modulate the mechanical properties of collagen gels, we evaluated both the micro- and macro- mechanical properties of the collagen networks formed in the presence of dimeric or oligomeric DDR2 ECD. This is especially important as both micro [91], [92][92]–[96] as well as macro[97]–[99] mechanical properties of collagen gels are understood to influence cell-matrix interactions and cell behaviors. We utilized optical tweezer-based active microrheology to evaluate micro-mechanical properties of collagen gels with/out non- collagenous proteins (NCPs). Two types of microrheology measurements were performed by using a calibrated, low frequency optical force. The first one-particle (1P) microrheology involved driving a micro-bead by the oscillating optical force and

56 measuring the response of that bead as has been used by others for collagen gels[93], [100]–

[102]. We also performed two-particle (2P) microrheology in which one bead was driven with the oscillating optical force, but now the response of a second, satellite bead was used to characterize the coarse-grained mechanical property of the material between the two beads. Macrorheological properties of identically prepared collagen gels were evaluated using a parallel-plate rheometer. We compared the 1P, 2P and macro-rheological moduli for each sample and across samples to understand how DDR2 dimer vs. oligomer modulates the mechanical properties of collagen gels. A non-collagen binding NCP, namely bovine serum albumin (BSA) was used as a control protein.

4.2 Methods

4.2.1 Collagen Gel Fabrication

Collagen solutions (2 mg/ml) were prepared by successive addition of 34.5% phosphate buffer saline (PBS) (7.5 pH), 1% 1N NaOH solution, and 64.5% vol/vol of 3.1 mg/ml bovine dermal collagen type 1 (PureCol from Advanced Biomatrix, San Diego). Collagen gels were also made with the addition 0.67 mg/ml of NCPs to achieve a NCP:collagen ratio of 1:3 w/w. Recombinant DDR2-Fc was purified in-house using plasmids encoding the human DDR2 ectodomain fused with a C-terminal Fc-tag as described in Chapter 2. Bovine serum albumin (BSA), (Sigma Aldrich, St. Louis) was used as a control, non-collagen binding protein. For microrheology measurements, 2µm diameter carboxyl-coated

57 polystyrene microspheres (ThermoFisher) were added to the solution prior to the polymerization process. All components were kept and mixed at 4°C, then pipetted into sterile circular glass molds (a #1.5 precision coverslip with a 20 mm diameter raised well of depth 2 mm), and allowed to polymerize for 24 hours at 37 °C, 5% CO2 and 95% humidity before testing.

4.2.2 Microrheology

A custom-built single beam optical tweezer instrumentation[103] was modified for active microrheology. Briefly, the laser was steered via a galvanometer (Thor Labs, Newton, NJ) controlled through LabView. Microbeads were subjected to oscillatory forces Fi(t) in directions i = x, y at a frequency of 2Hz using a moving laser trap whose force on the 2 m beads was described by a spring constant (kOT) of 0.17± 0.026 pN/nm and a sinusoidal displacement with a maximum amplitude () of 500 nm. The positions of the driven bead,

‘A’ and of the non-driven satellite bead ‘B’ were monitored by their fluorescence signal on a CCD which had a framerate of 20-40 Hz. At least 1000 frames were recorded for each measurement, corresponding to a total duration of 25-50s. A correlation method was applied on the individual CCD frames to find the positions of the beads as a function of time. A discrete Fourier transform (FT) of the bead position sequences was employed to determine the 2 Hz components for each driven or satellite bead (Figure 15). These (FT) components, in turn, were used to determine the in-phase (and out-of-phase) displacements of the driven and satellite beads, Ui (U’i) and ui (ui’) respectively.

The signal-to-noise (SNR) was determined (Figure 15) by a comprehensive evaluation of

(a) in-phase displacements of driven and satellite beads at 2Hz, in a direction transverse to

Fi(t) and (b) in-phase displacements of driven and satellite beads at off-resonance (1-1.5 and 2.5-3.46 Hz) in the direction of Fi(t). The uncertainties in measurement of satellite bead displacements were on the order of 0.3±0.2 nm. Our SNR for measurement of satellite bead displacement was >45 for all our samples and was significantly higher than the SNR observed in particle tracking passive microrheology approaches [104].

Figure 15. In-phase power spectrum of (a, c) driven and (b, d) satellite bead displacements along x and y-directions when Fi (the sinusoidal force of frequency 2Hz) was applied along x-direction. Beads were 10.2μm apart along the x-axis. Driven (Ux) and satellite (ux) bead displacements were calculated as the square root of peak amplitude in (a) and (c) respectively. Error in Ux and ux was determined using the signal present outside the 2Hz

60 signal. For comparison a power spectrum with no optical trap force is shown in (e).

(Manuscript submitted).

We verified that our laser trap displacement was linear with respect to both displacement and laser power, by varying Fi(t) (i.e. by changing either and/or kOT) and monitoring the driven-bead displacement, Ui, for individual beads (Figure 16).

Figure 16. (a) Linearity of trap tested at a constant trap strength, k (0.04pN/nm) at two OT

different trap displacements () for three beads in water. All beads yielded the same slope

as expected. (b) Linearity of trap tested using three different values of applied force (Fx)

62 obtained using a combination of different k or  (inset Table). All three beads tested OT exhibited a linear relationship between F and bead displacement (U ). (Manuscript x x submitted).

Figure 17. Schematic representing displacement (Ui: Ux or Uy) of the driven bead, ‘A’, and the induced displacement (ui: ux or uy) of the satellite bead, ‘B’, separated by a vector r.

(Manuscript submitted).

For 2P micro-rheology measurements, the axis joining the centroids from the driven to the non-driven bead was defined as the vector r (Figure 17). The bead-pairs were selected such that r was aligned along the x-axis (± 10o) in the frame of the camera CCD. Each bead was

63 subjected to a sinusoidal displacement, alternatively, along x or y by the oscillatory optical tweezer force, Fi. The maximum force Fi applied to a driven bead (along a irection i) is related to its elastic displacement, Ui in a material by Hooke’s law by using the following equation:

퐹푖 = 푘푂푇(∆ − 푈푖) [1]

For a material that is homogenous and isotropic, a micro-bead tightly coupled to the material will show a displacement U that satisfies: i

퐹푖 5−6푣 g푖 = [2] 푈푖 24휋(1−푣)푎

Where gi is the ‘shear modulus’ obtained using 1P-microrheology by driving a bead of radius a in the i direction. We utilized equation [2] to evaluate gi for collagen gels by taking the Poisson’s ratio, n , as 0.5 (value for an incompressible material) as reported in earlier studies[100]. We also calculated gi using  determined using 2P microrheology

(Figure 19).

For active two particle (2P) microrheology of a bead pair ‘AB’ separated by a distance r , the displacement of the satellite bead u due to a force, F , can be described by a shear i j

64 storage modulus G, by employing the equation describing the displacement field of a driven finite sphere of radius a placed at the origin of a linear elastic material [105]:

푎2 푎2 (퐹푗.푥푖)푥푗 (3−4휈+ 2)퐹푗훿푖푗+(1− 2) 2 푢 = 3푟 푟 푟 [3] 푖 16휋(1−휈)퐺푟

where xi and xj are the vector positions of the satellite bead with respect to the origin and

 is the Kronecker delta. For the (2P) measurements we make two measurements u , F ij ( x x ) and u , F . Thus, for each bead pair, using equation [3], we can extract a value of G, and ( y y ) the material’s Poisson’s ratio, n . The error in measurement of G and  was estimated using partial derivatives with respect to ux and uy such that the uncertainty propagation could be calculated. Also, it should be noted that for separations much larger than the bead radius, equation [3] predicts a fall-off in the displacement ui as ~ 1/ r or, G is independent of r.

Thus a distance dependence of G on r can be taken as evidence of mechanical heterogeneity for the length scale investigated.

At least n=6 bead pairs in each sample type were examined using active microrheology.

The distance, r , between beads in a pair ranged from ~ 5 to 15 m. For each bead pair, we also evaluated a reciprocal response in 2P microrheology by switching the role of the driven ‘A’ and satellite ‘B’ beads and ascertaining G in each case. We could thus compare the moduli ratio GAB:GBA for each bead pair.

4.2.3 Macrorheology

A stress-controlled rheometer (Kinexus ultra+; Malvern Instruments, UK) with titanium or stainless steel 20mm parallel plate geometry was used to measure the macroscopic shear storage (G’) and loss (G”) moduli of collagen gels. Samples with/out NCPs were prepared in an identical manner as described above, except without incorporation of latex beads.

Collagen samples were directly aliquoted on the rheometer plate (separated with a gap of

0.1 mm) and allowed to polymerize at 37°C in a humidified environment for 2 hours before testing. The samples were tested at an oscillatory frequency of 2Hz with strains ranging from 0.1 to 1.0%. Shear storage modulus (G’) and loss modulus (G”) were output directly by the rheometer software using Eq. 4.

2MD G' 4 cos R  [4] 2MD G" sin  R4

Where M is the torque, D is the gap height between the parallel plates, R is the radius of the geometry, δ is the phase angle, and θ is the deflection angle in radians.

4.2.4 Gel contraction assay

NIH3T3 murine fibroblasts were cultured in Dulbecco’s Modified Eagle Medium (DMEM,

Invitrogen) supplemented with 10% fetal bovine serum (Sigma Aldrich) and 1% penicillin- streptomycin solution (Sigma Aldrich) until 70% confluence at which time they were harvested for the gel contraction assay. Collagen solutions (2 mg/ml) were prepared by successive addition of 64.5% vol/vol of 3.1 mg/ml bovine dermal collagen type 1 (PureCol)

10% 10X DMEM, 1% 1N NaOH solution, and 24.5% cell inoculum (NIH3T3 fibroblasts) in the presence and absence of BSA or collagen binding protein in a w/w ratio of 1:3 (ex.

BSA:collagen). Cell-seeded gels (100,000 cells/cm2) were polymerized in 48 well plates for gel contraction assays. Cell-seeded collagen gels were allowed to polymerize for 5 hours at 37ºC in a CO2 incubator before additional supplemented cell-culture medium was added to maintain the cells. The gels were released from the wells 24 hours after onset of polymerization with an identical initial area of the cell-seeded collagen gels. Contraction of the collagen gels was monitored for up to 7 days by measuring the area (A) of the gels from digital images acquired at each time point using ImageJ.

4.3 Results

4.3.1 Viscoelastic nature of collagen gels

The material behavior was primarily elastic, as the out-of-phase component of the driven as well as satellite bead displacement was less than 20% of its in-phase displacement in microrheology (Figure 18a). Macro-rheology measurements also revealed that the loss moduli were less than 20% of the storage moduli for all samples. In addition, upon acquiring consecutive 1P and 2P microrheology measurements on the same bead(s) the resulting storage moduli were nearly constant with a std. dev less than 6% (Figure 18b).

The macrorheology measurements also showed a relatively constant storage (G’) (Figure

19a) and loss moduli (G”) (Figure 19b) at low strains for all samples except the one containing oligomeric DDR2. This sample was the only one that showed a change in moduli before failing at strains above 10%. Frequency sweeps performed at 0.1% strain on the same rheometer using collagen gels with PBS showed that there was a slight increase in storage modulus as the frequency increased, but there was little to no change in the loss modulus (Figure 20).

Figure 18. (a) Estimation of viscous vs. elastic response by the ratio of out-of-phase vs. in-phase displacement of the driven and satellite bead in the driven direction. Macro values were ascertained using ratio of loss vs. storage moduli determined using a parallel plate rheometer. (b) Repeated 1P and 2P micro-rheology measurements on the same bead in a 69

PBS collagen. Trials 2 and 3 were 10 and 20 minutes after trail 1 respectively. G=31.4±1.5 and g =71.8±4.1Pa (Manuscript submitted).

Figure 19. Strain sweeps of different collagen gels performed with parallel plate rheometry at 2Hz showing (a) storage (G’) and (b) loss moduli (G”). (Manuscript submitted).

Figure 20. Storage modulus (G’) and loss modulus (G”) of collagen gel with PBS over different frequencies performed with parallel plate rheometry at 0.1% strain. (Manuscript submitted).

4.3.2 Effect of DDRs on shear modulus of collagen I

We determined the 1P micro-moduli (gi) and 2P micro-moduli (G) using optical tweezer microrheology and equations [2] and [3] respectively. While the 2P moduli, GAB was measured for bead pairs ‘AB’ with inter-bead separation ranging from ~ 5 to 15 m, the

1P moduli, gAx was evaluated for each driven bead ‘A’ in a bead pair. We also ascertained

71 the macro-modulus (Gmacro) of similarly prepared collagen gels using a parallel-plate rheometer.

Figure 21. 1P micro-moduli (gi) calculated using n=0.5 and n determined using 2P microrheology as indicated. No significant differences in gi were observed for the two cases. (Manuscript submitted).

Figure 22. Comparison of 1P and macro rheology data for all samples. Oligomeric DDR2 resulted in the largest change in shear modulus compared to controls. (Manuscript submitted). 73

Figure 22. and Table 3. summarize the effect of DDR2 on the Gmacro and 1P moduli, gAx for collagen gels as compared to the control (PBS) sample. As shown in Figure 22a, the non- collagen-binding NCP, BSA did not significantly alter Gmacro or gAx of collagen gels as compared to PBS samples. In contrast, the collagen-binding NCPs differed in their effects on Gmacro vs. gAx. While DDR2-Fc gels exhibited an increase in Gmacro as compared to PBS

(p<0.05), this effect was not manifested in gAx measurements. Thus, collagen binding NCPs differentially modulated the macro vs. 1P micro-moduli of collagen gels.

In contrast to macrorheology, microrheology can enable estimating of local intra-sample mechanical heterogeneity within a sample. However, 1P microrheology is limited in its ability to characterize variations in local moduli, as bead-matrix coupling effects can significantly affect gi. As can be seen in Table 3, gAx values for all samples exhibited a ~

25 % variation from average and no significant differences in the mechanical heterogeneity across samples could be ascertained using 1P measurements alone. 2P microrheology on the other hand can provide a robust measure of heterogeneity in local moduli by evaluating

G as a function of coarse-grained material of length r. As shown in Figure 23, collagen gels with PBS, BSA, DDR2-Fc dimer and oligomer exhibited that GAB was nearly independent of r (determined using a linear fit, Table 3), indicating that these sample were mechanically homogenous at the length scales examined. 2P microrheology thus helped elucidate how collagen gels could be approximated as mechanically homogenous, linear elastic materials even with certain collagen binding proteins present.

In an effort to examine to what extent G is affected by the selection of driven vs. satellite bead, we also compared the reciprocal response obtained in 2P microrheology by switching the role(s) of driven and satellite beads for a subset (n=5) of bead pairs per sample. As seen in Figure 24, G between bead pairs remained unaffected by selection of the driven bead and the moduli ratio GAB/GBA ≈ 1 for all samples (Table 3).

To evaluate if the oligomeric status of DDR2 impacted the local Poisson’s ratio () of collagen gels, we determined for each bead pair using equation [3] in 2P microrheology.

As shown in Figure 25, majority of values fell in the 0 to 0.5 range for all samples with a few (n=2 for PBS and DDR2-Fc) outliers which exhibited abnormal values (>1) of .

The average value of  (~0.35) did not significantly differ between PBS and dimeric

DDR2-Fc samples (p>0.85) with BSA and oligomerized DDR2-Fc exhibiting a slightly higher  (~0.5) than PBS (p<0.004) (Table 3). For all samples  was independent of r. This experimentally determined  was also used to calculate 1P micro-moduli gi. No significant difference between gi calculated using =0.5 or  from 2P was observed for all samples

(Figure 14).

Figure 23. Coarse-grained micro-moduli (G) determined using 2P microrheology for each bead pair separated by a distance r. Dotted lines indicate average values of Gmacro and their std. dev. Dashed lines represent the average value of G in PBS, BSA and both DDR2-Fc samples. Solid line represents a linear fit of G vs. r. Average values and range for GAB and parameters for linear fit are listed in Table 1. (Manuscript submitted).

Figure 24. Moduli ratio obtained by measuring the reciprocal response in G for each bead pair by switching the role of driven ‘A’ and satellite bead ‘B’ in 2P microrheology. Ideal value of 1 is indicated by solid line. Dotted line indicated a linear fit of moduli ratio vs. r.

Average values of moduli ratio for each sample are listed in Table 1. (Manuscript submitted).

Figure 25. Poisson’s ratio () ascertained using ux/Fx and uy/Fy for each bead pair in 2P microrheology. Outliers falling outside 0≤≤1 are excluded from this plot. Dashed line indicates the average  obtained for each line. For comparison =0.5 for an incompressible material is indicated via dotted line. Solid lines represent a linear fit for  vs. r. Average values of  for each sample are listed in Table 1. (Manuscript submitted).

4.3.3 Effect of DDRs on gel contraction

To examine a manifestation of micro-moduli on cell matrix interaction, we monitored the cell mediated contraction of collagen gels prepared with or without DDR2. Figure 26 shows how gels with BSA or dimeric DDR2-Fc exhibited a nearly identical response as

PBS. In addition, after 2 days the PBS and BSA gels reached the same area. The most unexpected result of this experiment was the effect of oligomeric DDR2-Fc on the contraction of collagen gels. Collagen gels formed with oligomeric DDR2-Fc contracted to a much larger degree and at a faster rate than what was observed for all other samples

(p<0.05).

120% Collagen Gel Contractions PBS 100% BSA

80% DDR2-Fc

60% Olig. DDR2-Fc

40% Percent Percent Original Area 20%

0% 0 10 20 30 40 50 60 70 80 Time (hours)

Figure 26. Gel contraction assay results show that collagen formed in the presence of oligomerized DDR2-Fc contract to a very small size relative to the original area and at a very fast rate. The presence of other proteins in the collagen gel such as dimeric DDR2-Fc and BSA did not appear to alter the contraction of the gels when compared to the PBS control. (Manuscript submitted).

4.4 Discussion

In this study we demonstrate that by employing optical tweezer instrumentation, we could perform both 1P and 2P active microrheology using the same beads embedded in collagen gels. Active 2P microrheology has thus far primarily been accomplished using magnetic

80 tweezers[106], [107] with limited studies using optical tweezers[108]–[110]. 2P microrheology approaches are particularly advantageous as studies using passive microrheology have demonstrated that the 2P technique is insensitive to the bead-matrix coupling effects[111], [112] and provides a true measure of micro-scale moduli of the coarse-grained material. By using a parallel plate rheometer we also obtained macro- moduli of identically prepared samples and could thus compare the 1P, 2P and macro- moduli across and within samples.

Consistent with our observations, earlier studies using 1P approaches have yielded a large

(up to an order of magnitude) scatter [95], [96], [100], [101] and an overestimation of the storage moduli of the collagen gels compared to macroscale measurements[100][92]. An inconsistency in micro and macro moduli has also been reported when using passive

(particle tracking) 1P microrheology for other materials like F-actin and microtubules [112].

Bead-matrix coupling effects[111] and local structural changes in the vicinity of the beads[102] have been postulated as major factors affecting 1P microrheology measurements. These effects could be manifested as changes in the local density of material in the vicinity of a bead, amount of material attached to the beads and affine or non-affine local deformations of the attached fibrils. Based on our observations that 1P moduli, gi, for collagen gels are higher than their corresponding Gmacro we postulate an isostrain model[100] in which forces due to local bead-matrix coupling act in parallel with the elastic force exerted by the gel material such that the effective 1P moduli is a sum of Gmacro and a modulus arising due to bead-matrix coupling.

We elucidate here that for mechanically homogenous collagen networks (PBS, BSA and

DDR2-Fc) the active 2P moduli, G were similar to their corresponding Gmacro. In earlier studies, a similar comparison of 1P and 2P passive microrheology on actin networks has shown that 2P moduli exhibit a closer match with macro-rheology[113], [114]. 2P microrheology further enabled us to determine the Poisson’s ratio of coarse-grained material which is not possible when using 1P microrheology. Our estimates of are consistent with earlier micro[115] and macro [116], [117] measurements, where  ranged from 0 to 0.5 for collagen gels.

DDR2-Fc resulted in an increase in the macro-moduli of collagen gels as compared to PBS samples. This increase in macro-moduli can be accounted to the formation of a collagen network with thin fibrils as revealed by AFM analysis. These results are in accordance with earlier studies, which report that macro-moduli of collagen networks are inversely correlated to fibril diameter[118]. Our results on macro-moduli of DDR2-Fc differ from our earlier study[70] where DDR2-Fc was found to lower the persistence length (PL) (and corresponding Young’s modulus, E) of isolated collagen fibrils. This discrepancy can be explained in part by the methodology employed as a passive estimation of PL was used in our earlier study whereas the current study employs active microrheology. Further, network mechanics are influenced by additional factors such as inter-fibrillar crosslinks and nonaffine transformations, which are not present in isolated fibrils. In another study on collagen gels, the individual fibril stiffness was also not correlated with macro-scale moduli[97]. It is interesting to note that 1P moduli were not different between PBS and

DDR2-Fc, indicating that bead-matrix coupling effects may be lowering the effective moduli in the vicinity of the beads and thus compensating for the increase in Gmacro observed for DDR2-Fc gels.

Our study showed that despite a higher storage modulus obtained for collagen gels formed in the presence of oligomeric DDR2, the cell-mediated contraction of these gels was significantly enhanced. This result can partially be explained by comparing macrorheology and gel-contraction data. Strain-dependent macrorheology shows that oligomeric DDR2 samples show a failure at lower strains as compared to other samples. Since the forces exerted by cells in gel contraction are significantly higher than the low strain experienced in micro-rheology, we postulate that it is the high-strain behavior of gels which dictate contraction of collagen gels. As shown in Figure 19, the collagen samples containing oligomeric DDR2 ECD resulted in a failure at much lower strains as compared to other samples.

Taken together our results demonstrate that 3D collagen networks can be interpreted as a homogenous, linear elastic material and the oligomeric status of proteins like DDR2 ECD can modulate this behavior. We further show that it is important to characterize both the macro and micro-scale mechanical properties as they could be differentially modulated by collagen binding NCPs. Our investigations also suggest that assessment of microrheological properties using 1P approach(es) should be used with caution as they can significantly depart from the macro-scale mechanical behavior of the sample. 2P microrheology provides a more accurate measure of micro-scale moduli and can be 83 particularly useful in characterizing micro-mechanical heterogeneities. A robust mechanical and structural evaluation at the microscale, as presented in our study holds importance to understand the mechanical properties of the complex matrix environment present in-vivo and its modulation by NCPs.

PBS BSA DDR2 O. DDR2 Parameters describing heterogeneity of collagen gels using micro-rheology Linear fit for G vs. r m -0.31 -0.31 -0.09 -1.67 (using 2P micro- b 26.1 32.9 28.9 101.3 rheology) R2 0.04 0.02 0.00 0.08 Parameters describing mechanical properties of collagen gels (average ± SD) G (in Pa): Macro-rheology 20.0±2.1 20.5±1.9 36.6±4.7 71.0±9.1 G (in Pa): 2P micro-rheology 22.8.0±5.7 31.3±7.7 31.5±9.2 82.2±30.0 Range of 2P G (in Pa) 14 – 28 21 – 47 21 – 50 49 – 139 gAx (in Pa): 1P micro-rheology 61.3±14.5 68.6±21.4 60.4±14.6 272.2±85.6 Range of 1P gAx (in Pa) 35 – 78 46 – 139 40 – 90 154 – 399 Moduli ratio (GAB/GBA) from 2P 0.98±0.10 0.93±0.13 1.0±0.12 0.91±0.10 Range of 1P g (in Pa) 35 – 78 46 – 139 40 – 90 154 – 399  from 2P microrheology 0.36±0.17 0.50±0.07 0.36±0.12 0.47±0.08

Table 3. Parameters describing heterogeneity and mechanical properties of collagen gels.

R2: coefficient of determination for the linear fit G=mr+b. (Manuscript submitted).

Chapter 5: Conclusions

5.1 Comparison of oligomeric status of DDRs with other RTKS post ligand binding

DDRs are often considered an anomaly of sorts in the family of RTKs. This is because unlike other RTKs, which bind to soluble growth factors, DDRs bind to collagen(s) an insoluble ECM component. In addition, DDRs exhibit an unusually slow phosphorylation rate upon ligand binding as compared to other RTKs [63], [62]. Further, the inability of chimeric versions of other RTKs such as platelet derived growth factor receptor (PDGFR)

ECDs when combined with DDR1 ICDs to be ligand-activated [119] point to distinct structural differences inherent to the DDRs.

Unlike the commonly accepted paradigm for RTKs, DDRs exist as constitutive dimers instead of undergoing the commonly accepted pathway of ligand mediated dimerization of

RTKs such as such as Tropomyosin receptor kinase A (TrkA) [120], tyrosine protein kinase

Kit (KIT) [121], [122], fibroblast growth factor receptor (FGFR) [44], [123]. However, recent studies indicate that DDRs may have more in common with other receptors than 85 previously thought. For example, the Insulin receptor and IGF1-receptor also exist on the cell surface as dimers. Their binding to ligands induces structural changes which result in tyrosine kinase activity [124]. Additional RTKs like EGF and TrKA and also exist as dimers in an inactive state before binding to their ligands [125]–[127].

Oligomerization of membrane receptors plays an important role in the receptor function and existing literature points toward specific downstream signaling that is unique to multivalent ligands [128], [129] (Table 4). Numerous receptors including both RTKs and other receptors, such as EGFR [130], Tie2 [131], integrins [132] toll-like receptor [133],

ErbB family [134], and Ephrin [135] have been found to assemble into higher-order structures where downstream signaling events are often correlated to cluster formation.

Consistent with our observations with DDR1, a large heterogeneity in oligomer size has been reported for other RTKs, like the epidermal growth factor receptor (EGFR) [136]. A recent study using a nanopatterned supported lipid bilayer technique to control EGFR clustering levels in living cells found that large-scale clustering of EGFR dampens its phosphorylation and that the cell endocytosis machinery contributes to this clustering behavior [130].

Receptor Pre-ligand Post- Receptor Phosphorylation Reference binding ligand endocytosis binding EGFR/ErbB Monomer Dimer Rapid after Dimers become [137]– which ligand tetramers and [139] become binding oligomers after oligomers activation Tie2 Monomer Clusters Rapid after Occurs on [131], ligand clusters [140] binding Integrins Heterodimer Focal Yes Clustering to aid [141]– adhesions affinity may [143] enhance activation Toll-like Heterodimer Cluster Lipid raft Occurs at cell [133] dependent surface after clustering Ephrin Monomer Clusters Terminates Occurs on [144], signaling clusters [145] DDR1 Dimer Clusters Yes Occurs on [22], [79] preformed clusters DDR2 Dimer Filaments Unknown Occurs on Manuscript filamentous submitted structures

Table 4. Comparison of oligomeric status of RTKs before and after ligand binding

Dimerization of RTKs is understood to involve conserved motifs, particularly, in the transmembrane domain [146]. One of the earlier motifs thought to mediate TM dimerization was the P0-P4 motif [147]. This is similar to the GXXXG-like motifs which are important for dimerization of glycophorin A and several receptors of the ErbB family

[148]–[150]. The transmembrane domain of DDR1 also contains the GXXXG motif, however in terms of dimerization, it appears to be overshadowed by the leucine zipper also found in the TM domain [151]. Additionally, recent studies have shown that the TM also

87 plays a role in the clustering of DDR1 dimers together and their subsequent phosphorylation after collagen binding [68].

The ECD of DDRs is another domain which can influence the oligomeric status of the receptor. Crystallization experiments have shown the existence of a conserved patch which exists on an outward facing portion of the DS domain in DDR1 but away from the collagen binding region. It has been shown that a portion of the DS-like domain of one DDR is able to interact with this patch on a separate DDR molecule. While certain mutations in this area did not impact collagen binding or expression of the receptors, they prevented DDR1 function [35]. Therefore, one postulation is that these patches in the extracellular domain are involved in clustering one dimer to another resulting in larger oligomers [37].

Clustering of RTKs can also influence the spatial location of the receptor and this seems to be the case for DDR1. In our previous and current studies, we elucidated how DDR1 clustering occurs on the cell surface and is followed by receptor endocytosis in a dynamin- dependent manner [79]. DDR1b used in our studies consist of NPXY motif which is known to mediate receptor endocytosis. It is interesting to note that this motif is absent in DDR2 and in certain isoforms of DDR1 such as DDR1a and it is likely that these receptors do not undergo endocytosis. Other RTKs which undergo ligand-induced endocytosis include insulin receptor (IR), insulin-like growth factor 1 receptor (IGF-1R), platelet-derived growth factor receptor (PDGFR) β [152]–[155] and consist of the NPXY motif.

Our study elucidates how clustering and internalization of DDR1 occurs prior to its phosphorylation. This is consistent with other studies which indicate that certain RTKs like

EGFR and TrkA, and can activate intracellular signaling even when inside endocytic vesicles [156]–[158]. Whereas it may have been previously thought that receptor internalization was a means to stop receptor activation, this may not be true and is not the case with DDR1. The delayed onset of the phosphorylation of DDRs in our cell based assays is consistent with earlier reports. Recent studies have shown a link between DDR1- collagen binding and the formation of invadosomes which are F-actin based protrusions that degrade the ECM to enable tumor cell invasion [159]. The structure of the invadosomes formed via DDR1 activation in cancer cells is remarkably similar to the filamentous organization of DDR2 which was observed in the live cell assays performed in chapter 2. Further studies need to be performed to identify if these formations are indeed invadosomes. Taken together, the relationship between the temporal-spatial organization of the DDRs and their frequency of phosphorylation continues to raise questions about the mechanisms at play.

5.2 Comparison of oligomeric status of DDRs with other collagen binding proteins

Another important aspect of DDRs in interacting with their ligand collagen(s) is the oligomeric status of DDR ECD pre-ligand binding. A few other collagen receptors, glycoproteins and proteoglycans have also been studies to show how their oligomeric status 89 impacts collagen binding. Among these the collagen binding integrins α1β1 and α2β1 are the most common collagen receptors [160]. The end of the α2 subunit, binds to the collagen triple helix through a metal ion-dependent adhesion site, [160], [161]. It has been shown that conformational changes to integrins and their clustering into oligomers can affect their binding affinity to ligands [162], but the degree to which this impacts the function of integrins is still unclear. What is known is that the clustering of integrins can be caused by several different factors including inside-out signaling [163]–[165], the homodimerization of integrin extracellular domains during ligand binding [166], or the diffusion of integrins in cell membrane after their release from cytoskeletal constraints [167]. This clustering has also been demonstrated to be important for the transmission of signals caused by bound extracellular ligands, the recycling of integrins [168], and intracellular mechanotransduction [169]. Thus, DDRs and integrins appear to share some similarities in the role of their oligomeric status pre- and post- ligand-binding.

Other collagen receptors include the Glycoprotein VI (GPVI), LAIR-1 and OSCAR [170].

The GPVI is comprised of two immunoglobulin-like (IG) domains followed by a mucine- like stalk domain and small intracellular domain [171]. The crystal structure of collagen bound GPVI reveals a back-to-back dimer configuration resulting from the interaction between the IG2 domains [172]. This organizational phenomenon is supported by results showing that GPVI forms dimers on the surface of platelets [173]. Studies using recombinant GPVI-Fc have shown that in its dimeric form, GPVI was able to bind to collagen with much higher affinity than in its monomeric form [174].

Another set of collagen binding proteins are immune inhibitory receptors named leukocyte associated Ig-like receptors (LAIR) 1 and 2. These also show differential binding ability based on their clustering with LAIR-2 showing a higher affinity to collagen. In vitro experiments have shown that dimeric forms of the LAIRs both exhibit much stronger collagen binding than their monomeric forms [175]. Another member of the immunoglobulin family of receptors which binds to collagen is the osteoclast associated receptor (OSCAR). Its involvement in osteoclast differentiation is area of ongoing study.

Several binding sites on the collagen monomer have been identified, and it is this observation that has led others to believe that OSCAR may undergo clustering when binding to an abundant ECM target [176]. It is interesting to note that like collagen receptors, a subset of membrane-bound proteoglycans also tend to form oligomers [177].

Of note are, syndecans, which bind to collagen through their HS domains and their subsequent clustering is vital for their ability to transmit signals influencing cell motility and adhesion [178].

Besides collagen receptors a large number of glycoproteins and proteoglycans bind to collagen. These include SPARC and vWF which share the collagen binding site as DDR1 and DDR2 as well as SLRPs consisting of a core protein with leucine rich repeats (LRRs), and one or more attached GAGs [179]. The best studied among these is the SLRP decorin, which plays a role in regulating not only collagen fibrillogenesis [180], [181], but other related ECM remodeling such as tumor metastasis [182] and angiogenesis [183]. The

SLRPs tend to have a characteristic curvature associated with their core protein, and it was commonly believed that the ability to bind to their ligands, such as collagen, relied on 91 individual monomers contacting their binding sites on the inside of the 'horeshoe' shape

[184]. However, more recent discoveries have suggested that decorin exists as a dimer in solution [185], [186]. Several new configurations of SLRP binding have been posited including dimer to monomer transitions or oligomerization binding patterns [187]. In fact, the work done by Orgel et al. have supported the idea that the decorin monomers must dissociate from their dimeric form to yield the most energetically favorable binding interactions with collagen [188]. Thus, the role of oligomeric status of decorin in collagen binding may be acting in a manner very different from that of DDRs.

5.3 Role of DDRs in matrix mechanics and cell behavior

It has become increasingly understood that cells derive cues from the mechanical properties of the surrounding environment. The stiffness of the extracellular matrix, as well as the microarchitecture of the matrix have been shown to influence cell migration [189], [190], proliferation [191]–[193], cell spreading [194]–[197], and even the differentiation of some stem cells [198], [199]. A primary challenge in the understanding of matrix properties on cell behavior is the ability to separate possible confounding factors from each other such as matrix stiffness, density, and binding locations. Understanding how cells interact with an ECM modified by DDRs is particularly challenging as the modified collagen fibril structure does not only impact the network mechanics but may also differentially expose

DDR (as well as other) binding sites on its surface.

To this end, it has been shown that smooth muscle cells isolated from the arteries of DDR1- null mice behaved differently in culture compared to their wild type counterparts. The results of this study suggest that DDR1 plays a role in cells’ ability to attach to collagen, migrate, proliferate and produce MMPs [24]. Other work has shown the impact of DDRs on matrix stiffness. For example, DDR2-/- mice displayed wounded skin with lower tensile strength than wild types [200]. Corsa et al. also suggest that since tumors in DDR2-/- mice lack the same amount and thickness of collagen compared to DDR2+/+ they may have decreased rigidity [201]. Our studies show that recombinant DDR2 showed a strong propensity to inhibit normal collagen fibrillogenesis as evidenced by the smaller fibrils seen in AFM and confocal reflectance. This was phenomenon was even more strongly observed with the oligomerized form of DDR2, resulting in a stiffer matrix n. This is also noteworthy due to the lack of other matrix constituents and in the absence of cross-linking.

However, a puzzling yet informative result from this study was the ability of cells to contract the collagen gels containing oligomerized DDR2 to a much greater extent than any other treatment of collagen tested. This poses questions for future studies about how cells interact with their matrix during contraction beyond just exerting force on their surroundings. A differential ability to modulate the collagen fibrils formed in the presence of oligomerized DDR2 may be occurring.

5.4 Future Directions and translational potential

The use of collagen scaffolds has been traditionally approached by either the decellularization of existing ECM or by the polymerization of solubilized collagen [202].

The latter may provide more flexibility due to the many parameters that can be varied such as concentration or incorporation of other materials. However, the downside of this approach is that these structures are often mechanically more fragile unless very high concentrations of collagen are used. The use of recombinant proteins to control the mechanical properties of collagen gels has been emerging as a unique way to modulate the collagen fibril formation and adjust the network structure of the matrix. Studies using recombinant proteins have shown how the matrix can be stiffened using decorin [203].

In vivo, both DDR1 and DDR2 being implicated in a variety of pathological conditions from developmental diseases to cancer and fibrosis [20],. One commonality that may link all of these pathologies together is that they involve some form of extracellular matrix remodeling. This is not surprising since DDRs bind to collagen and have been shown to influence MMP production in different cell types [63], [23]. In conjunction with this, results from this study as well as previous work have shown how the binding of DDRs to collagen fibrils alone have been able to alter the fibrillogenesis and matrix architecture [69],

[70], [204]. Thus, expression and/or phosphorylation profile of proteins like DDRs can be investigated as potential biomarkers for altering the matrix architecture as well as mechanical properties. Therefore, understanding the expression, phosphorylation and

ECM remodeling by DDRs has widespread relevance in interpreting cell behavior as well as for designing collagen scaffolds for regenerative and tissue engineering applications. 94

References

[1] B. Leitinger, “Transmembrane Collagen Receptors,” Annu. Rev. Cell Dev. Biol.,

vol. 27, no. 1, pp. 265–290, 2011.

[2] C. Mihai, M. Chotani, T. Elton, and G. Agarwal, “Mapping of DDR1 distribution

and oligomerization on the cell surface by FRET microscopy,” J. Mol. Biol., vol.

385, no. 2, pp. 432–445, 2009.

[3] B. Leitinger, “Molecular analysis of collagen binding by the human discoidin

domain receptors, DDR1 and DDR2. Identification of collagen binding sites in

DDR2.,” J. Biol. Chem., vol. 278, no. 19, pp. 16761–9, May 2003.

[4] N. a Noordeen, F. Carafoli, E. Hohenester, M. a Horton, and B. Leitinger, “A

transmembrane leucine zipper is required for activation of the dimeric receptor

tyrosine kinase DDR1.,” J. Biol. Chem., vol. 281, no. 32, pp. 22744–51, Aug.

2006.

[5] C. Lai and G. Lemke, “Structure and expression of the Tyro 10 receptor tyrosine

kinase.,” Oncogene, vol. 9, no. 3, pp. 877–83, Mar. 1994.

[6] M. P. Sánchez, P. Tapley, S. S. Saini, B. He, D. Pulido, and M. Barbacid,

“Multiple tyrosine protein kinases in rat hippocampal neurons: isolation of Ptk-3, a

receptor expressed in proliferative zones of the developing brain.,” Proc. Natl. 95

Acad. Sci. U. S. A., vol. 91, no. 5, pp. 1819–23, Mar. 1994.

[7] E. Di Marco, N. Cutuli, L. Guerra, R. Cancedda, and M. De Luca, “Molecular

Cloning of trlzE, a Novel trk-related Putative Mosine Kinase Receptor Isolated

from Normal Human Keratinocytes and Widely Expressed by Normal Human

Tissues*,” vol. 268, no. 32, pp. 24290–24295, 1993.

[8] J. D. Johnson, J. C. Edmant, and W. J. Rutter, “A receptor tyrosine kinase found in

breast carcinoma cells has an extracellular discoidin I-like domain,” Biochemistry,

vol. 90, pp. 5677–5681, 1993.

[9] S. Laval, R. Butler, A. N. Shelling, A. M. Hanby, R. Poulsom, and T. S. Ganesan2,

“Isolation and Characterization of an Epithelial-specific Receptor Tyrosine Kinase

from an Ovarian Cancer Cell Line1,” Cell Growth Differ., vol. 5, pp. 1173–1183,

1994.

[10] J. L. Perez, S. Q. Jing, and T. W. Wong, “Identification of two isoforms of the Cak

receptor kinase that are coexpressed in breast tumor cell lines.,” Oncogene, vol.

12, no. 7, pp. 1469–77, Apr. 1996.

[11] J. L. Perez, X. Shen, S. Finkernagel, L. Sciorra, N. A. Jenkins, D. J. Gilbert, N. G.

Copeland, and T. W. Wong, “Identification and chromosomal mapping of a

receptor tyrosine kinase with a putative phospholipid binding sequence in its

ectodomain.,” Oncogene, vol. 9, no. 1, pp. 211–9, Jan. 1994.

[12] F. Alves, W. Vogel, K. Mossie, B. Millauer, H. Höfler, and A. Ullrich, “Distinct

structural characteristics of discoidin I subfamily receptor tyrosine kinases and

complementary expression in human cancer.,” Oncogene, vol. 10, no. 3, pp. 609–

18, Feb. 1995. 96

[13] B. Roig, N. Franco-Pons, L. Martorell, J. Tomàs, W. F. Vogel, and E. Vilella,

“Expression of the tyrosine kinase discoidin domain receptor 1 (DDR1) in human

central nervous system myelin,” vol. 1, 2018.

[14] R. Ram, G. Lorente, K. Nikolich, R. Urfer, E. Foehr, and U. Nagavarapu,

“Discoidin Domain Receptor-1a (DDR1a) Promotes Glioma Cell Invasion and

Adhesion in Association with Matrix Metalloproteinase-2,” J. Neurooncol., vol.

76, no. 3, pp. 239–248, Feb. 2006.

[15] V. Abbonante, C. Gruppi, D. Rubel, O. Gross, R. Moratti, and A. Balduini,

“Discoidin domain receptor 1 protein is a novel modulator of megakaryocyte-

collagen interactions.,” J. Biol. Chem., vol. 288, no. 23, pp. 16738–46, Jun. 2013.

[16] H. Kamohara, S. Yamashiro, C. Galligan, and T. Yoshimura, “Discoidin domain

receptor 1 isoform-a (DDR1alpha) promotes migration of leukocytes in three-

dimensional collagen lattices.,” FASEB J., vol. 15, no. 14, pp. 2724–6, Dec. 2001.

[17] J. Márquez and E. Olaso, “Role of discoidin domain receptor 2 in wound healing.,”

Histol. Histopathol., vol. 29, no. 11, pp. 1355–64, Nov. 2014.

[18] J.-E. Lee, C.-S. Kang, X.-Y. Guan, B.-T. Kim, S.-H. Kim, Y.-M. Lee, W.-S.

Moon, and D.-K. Kim, “Discoidin domain receptor 2 is involved in the activation

of bone marrow-derived dendritic cells caused by type I collagen,” Biochem.

Biophys. Res. Commun., vol. 352, no. 1, pp. 244–250, Jan. 2007.

[19] W. F. Vogel, R. Abdulhussein, and C. E. Ford, “Sensing extracellular matrix: an

update on discoidin domain receptor function.,” Cell. Signal., vol. 18, no. 8, pp.

1108–16, Aug. 2006.

[20] C. M. Borza and A. Pozzi, “Discoidin domain receptors in disease.,” Matrix Biol., 97

vol. 34, pp. 185–92, Feb. 2014.

[21] B. Leitinger, “Discoidin domain receptor functions in physiological and

pathological conditions,” Int. Rev. Cell Mol. Biol., vol. 310, pp. 39–87, 2014.

[22] A. R. Blissett, D. Garbellini, E. P. Calomeni, C. Mihai, T. S. Elton, and G.

Agarwal, “Regulation of Collagen Fibrillogenesis by Cell-surface Expression of

Kinase Dead DDR2,” J. Mol. Biol., vol. 385, no. 3, pp. 902–911, 2009.

[23] N. Ferri, N. O. Carragher, and E. W. Raines, “Role of discoidin domain receptors 1

and 2 in human smooth muscle cell-mediated collagen remodeling: potential

implications in atherosclerosis and lymphangioleiomyomatosis.,” Am. J. Pathol.,

vol. 164, no. 5, pp. 1575–85, May 2004.

[24] G. Hou, W. Vogel, and M. P. Bendeck, “The discoidin domain receptor tyrosine

kinase DDR1 in arterial wound repair.,” J. Clin. Invest., vol. 107, no. 6, pp. 727–

35, Mar. 2001.

[25] G. Hou, W. F. Vogel, and M. P. Bendeck, “Tyrosine Kinase Activity of Discoidin

Domain Receptor 1 Is Necessary for Smooth Muscle Cell Migration and Matrix

Metalloproteinase Expression,” Circ. Res., vol. 90, no. 11, pp. 1147–1149, Jun.

2002.

[26] M. E. Roberts, L. Magowan, I. P. Hall, and S. R. Johnson, “Discoidin domain

receptor 1 regulates bronchial epithelial repair and matrix metalloproteinase

production.,” Eur. Respir. J. Off. J. Eur. Soc. Clin. Respir. Physiol., vol. 37, no. 6,

pp. 1482–93, Jul. 2011.

[27] E. Olaso, K. Ikeda, F. J. Eng, L. Xu, L. Wang, H. C. Lin, and S. L. Friedman,

“DDR2 receptor promotes MMP-2 – mediated proliferation and invasion by 98

hepatic stellate cells,” vol. 108, no. 9, pp. 1369–1378, 2001.

[28] P. V Afonso, C. P. McCann, S. M. Kapnick, and C. A. Parent, “Discoidin domain

receptor 2 regulates neutrophil chemotaxis in 3D collagen matrices.,” Blood, vol.

121, no. 9, pp. 1644–50, Feb. 2013.

[29] J. Su, J. Yu, T. Ren, W. Zhang, Y. Zhang, X. Liu, T. Sun, H. Lu, K. Miyazawa,

and L. Yao, “Discoidin domain receptor 2 is associated with the increased

expression of matrix metalloproteinase-13 in synovial fibroblasts of rheumatoid

arthritis,” Mol. Cell. Biochem., vol. 330, no. 1–2, pp. 141–152, Oct. 2009.

[30] R. R. Valiathan, M. Marco, B. Leitinger, C. G. Kleer, and R. Fridman, “Discoidin

domain receptor tyrosine kinases: new players in cancer progression.,” Cancer

Metastasis Rev., vol. 31, no. 1–2, pp. 295–321, Jun. 2012.

[31] C. Avivi-Green, M. Singal, and W. F. Vogel, “Discoidin domain receptor 1-

deficient mice are resistant to bleomycin-induced lung fibrosis.,” Am. J. Respir.

Crit. Care Med., vol. 174, no. 4, pp. 420–7, Aug. 2006.

[32] S. Development, “6 . Jiapeng Xu ( 2007-2009 ) graduated with a M . S . in

Chemical and Biomolecular Eng . ( PhD from Boston University ) Present

employer : SolidEnergy Systems , Greater Boston Area Position : Senior Coating

Scientist,” pp. 2007–2008, 2014.

[33] L. Xu, H. Peng, S. Glasson, P. L. Lee, K. Hu, K. Ijiri, B. R. Olsen, M. B. Goldring,

and Y. Li, “Increased expression of the collagen receptor discoidin domain

receptor 2 in articular cartilage as a key event in the pathogenesis of

osteoarthritis,” Arthritis Rheum., vol. 56, no. 8, pp. 2663–2673, Aug. 2007.

[34] F. Carafoli, D. Bihan, S. Stathopoulos, A. D. Konitsiotis, M. Kvansakul, R. W. 99

Farndale, B. Leitinger, and E. Hohenester, “Crystallographic insight into collagen

recognition by discoidin domain receptor 2.,” Structure, vol. 17, no. 12, pp. 1573–

81, Dec. 2009.

[35] F. Carafoli, M. C. Mayer, K. Shiraishi, M. A. Pecheva, L. Y. Chan, R. Nan, B.

Leitinger, and E. Hohenester, “Structure of the discoidin domain receptor 1

extracellular region bound to an inhibitory Fab fragment reveals features important

for signaling.,” Structure, vol. 20, no. 4, pp. 688–97, Apr. 2012.

[36] O. Ichikawa, M. Osawa, N. Nishida, N. Goshima, N. Nomura, and I. Shimada,

“Structural basis of the collagen-binding mode of discoidin domain receptor 2,”

EMBO J., vol. 26, pp. 4168–4176, 2007.

[37] F. Carafoli and E. Hohenester, “Collagen recognition and transmembrane

signalling by discoidin domain receptors.,” Biochim. Biophys. Acta, vol. 1834, no.

10, pp. 2187–94, Oct. 2013.

[38] T. N. Phan, E. L. Wong, S. Y. Park, H. J. Kim, and B. S. Yang, “Defective Ca2+

binding in a conserved binding site causes incomplete N-glycan processing and

endoplasmic reticulum trapping of discoidin domain receptors,” Biosci.

Biotechnol. Biochem., vol. 79, no. 4, pp. 574–580, 2015.

[39] B. R. Ali, H. Xu, N. A. Akawi, A. John, N. S. Karuvantevida, R. Langer, L. Al-

Gazali, and B. Leitinger, “Trafficking defects and loss of ligand binding are the

underlying causes of all reported DDR2 missense mutations found in SMED-SL

patients,” Hum. Mol. Genet., vol. 19, no. 11, pp. 2239–2250, 2010.

[40] C. A. Curat, M. Eck, X. Dervillez, and W. F. Vogel, “Mapping of Epitopes in

Discoidin Domain Receptor 1 Critical for Collagen Binding*,” 2001. 100

[41] T. N. Phan, E. L. Wong, X. Sun, G. Kim, S. H. Jung, C. N. Yoon, and B.-S. Yang,

“Low Stability and a Conserved N-Glycosylation Site Are Associated with

Regulation of the Discoidin Domain Receptor Family by Glucose via Post-

Translational N-Glycosylation,” Biosci. Biotechnol. Biochem., vol. 77, no. 9, pp.

1907–1916, 2013.

[42] H.-L. Fu, R. R. Valiathan, R. Arkwright, A. Sohail, C. Mihai, M. Kumarasiri, K. V

Mahasenan, S. Mobashery, P. Huang, G. Agarwal, and R. Fridman, “Discoidin

domain receptors: unique receptor tyrosine kinases in collagen-mediated

signaling.,” J. Biol. Chem., vol. 288, no. 11, pp. 7430–7, Mar. 2013.

[43] R. Abdulhussein, C. McFadden, P. Fuentes-Prior, and W. F. Vogel, “Exploring the

collagen-binding site of the DDR1 tyrosine kinase receptor.,” J. Biol. Chem., vol.

279, no. 30, pp. 31462–70, Jul. 2004.

[44] J. Schlessinger, “Cell Signaling by Receptor Tyrosine Kinases A large group of

genes in all eukaryotes encode for,” October, vol. 103, no. 2, pp. 211–225, 2000.

[45] M. A. Lemmon and J. Schlessinger, “Cell signaling by receptor tyrosine kinases,”

Cell, vol. 141, no. 7, pp. 1117–1134, 2010.

[46] R. Abdulhussein, D. H. H. Koo, and W. F. Vogel, “Identification of disulfide-

linked dimers of the receptor tyrosine kinase DDR1.,” J. Biol. Chem., vol. 283, no.

18, pp. 12026–33, May 2008.

[47] C. Finger, C. Escher, and D. Schneider, “The Single Transmembrane Domains of

Human Receptor Tyrosine Kinases Encode Self-Interactions,” Sci. Signal., vol. 2,

no. 89, p. ra56-ra56, Sep. 2009.

[48] G. N. RAMACHANDRAN and G. KARTHA, “Structure of collagen.,” Nature, 101

vol. 176, no. 4482, pp. 593–5, Apr. 1955.

[49] A. Rich, U. S. A. And, and F. H. C. Crick, “The Molecular Structure of Collagen,”

J. Mol. Biol, vol. 3, pp. 483–506, 1961.

[50] A. RICH and F. H. C. CRICK, “The Structure of Collagen,” Nature, vol. 176, no.

4489, pp. 915–916, Nov. 1955.

[51] B. Leitinger and A. P. L. Kwan, “The discoidin domain receptor DDR2 is a

receptor for type X collagen,” Matrix Biol., vol. 25, no. 6, pp. 355–364, Aug.

2006.

[52] G. Agarwal, L. Kovac, C. Radziejewski, and S. J. Samuelsson, “Binding of

discoidin domain receptor 2 to collagen I: an atomic force microscopy

investigation.,” Biochemistry, vol. 41, no. 37, pp. 11091–8, Sep. 2002.

[53] G. Agarwal, C. Mihai, and D. F. Iscru, “Interaction of discoidin domain receptor 1

with collagen type 1.,” J. Mol. Biol., vol. 367, no. 2, pp. 443–55, Mar. 2007.

[54] A. D. Konitsiotis, N. Raynal, D. Bihan, E. Hohenester, R. W. Farndale, and B.

Leitinger, “Characterization of high affinity binding motifs for the discoidin

domain receptor DDR2 in collagen.,” J. Biol. Chem., vol. 283, no. 11, pp. 6861–8,

Mar. 2008.

[55] H. Xu, N. Raynal, S. Stathopoulos, J. Myllyharju, R. W. Farndale, and B.

Leitinger, “Collagen binding specificity of the discoidin domain receptors: binding

sites on collagens II and III and molecular determinants for collagen IV

recognition by DDR1.,” Matrix Biol., vol. 30, no. 1, pp. 16–26, Jan. 2011.

[56] D. J. S. Hulmes and A. Miller, “Quasi-hexagonal molecular packing in collagen

fibrils,” Nature, vol. 282, no. 5741, pp. 878–880, 1979. 102

[57] J. P. R. O. Orgel, O. Antipova, I. Sagi, a Bitler, D. Qiu, R. Wang, Y. Xu, and J. D.

San Antonio, “Collagen fibril surface displays a constellation of sites capable of

promoting fibril assembly, stability, and hemostasis.,” Connect. Tissue Res., vol.

52, no. 1, pp. 18–24, Feb. 2011.

[58] J. P. R. O. Orgel, T. C. Irving, A. Miller, and T. J. Wess, “Microfibrillar structure

of type I collagen in situ,” Proc. Natl. Acad. Sci., vol. 103, no. 24, pp. 9001–9005,

2006.

[59] A. B. Herr and R. W. Farndale, “Structural insights into the interactions between

platelet receptors and fibrillar collagen.,” J. Biol. Chem., vol. 284, no. 30, pp.

19781–5, Jul. 2009.

[60] S. Perumal, O. Antipova, and J. P. R. O. Orgel, “Collagen fibril architecture,

domain organization, and triple-helical conformation govern its proteolysis.,”

Proc. Natl. Acad. Sci. U. S. A., vol. 105, no. 8, pp. 2824–9, Feb. 2008.

[61] C. L. Hoop, J. Zhu, A. M. Nunes, D. A. Case, and J. Baum, “Revealing

accessibility of cryptic protein binding sites within the functional collagen fibril,”

Biomolecules, vol. 7, no. 4. 2017.

[62] a Shrivastava, C. Radziejewski, E. Campbell, L. Kovac, M. McGlynn, T. E. Ryan,

S. Davis, M. P. Goldfarb, D. J. Glass, G. Lemke, and G. D. Yancopoulos, “An

orphan receptor tyrosine kinase family whose members serve as nonintegrin

collagen receptors.,” Mol. Cell, vol. 1, no. 1, pp. 25–34, Dec. 1997.

[63] W. Vogel, G. D. Gish, F. Alves, and T. Pawson, “The Discoidin Domain Receptor

Tyrosine Kinases Are Activated by Collagen,” Mol. Cell, vol. 1, pp. 13–23, 1997.

[64] C. Mihai, D. F. Iscru, L. J. Druhan, T. S. Elton, and G. Agarwal, “Discoidin 103

domain receptor 2 inhibits fibrillogenesis of collagen type 1.,” J. Mol. Biol., vol.

361, no. 5, pp. 864–76, Sep. 2006.

[65] H.-L. Fu, R. R. Valiathan, L. Payne, M. Kumarasiri, K. V Mahasenan, S.

Mobashery, P. Huang, and R. Fridman, “Glycosylation at ASN211 regulates the

activation state of the discoidin domain receptor 1 (DDR1).,” J. Biol. Chem., vol.

1, Mar. 2014.

[66] H. Xu, T. Abe, J. K. H. Liu, I. Zalivina, E. Hohenester, and B. Leitinger, “Normal

activation of discoidin domain receptor 1 mutants with disulphide cross-links,

insertions or deletions in the extracellular juxtamembrane region: mechanistic

implications.,” J. Biol. Chem., pp. 0–22, Mar. 2014.

[67] N. M. Coelho, P. D. Arora, S. van Putten, S. Boo, P. Petrovic, A. X. Lin, B. Hinz,

and C. A. McCulloch, “Discoidin Domain Receptor 1 Mediates Myosin-

Dependent Collagen Contraction,” Cell Rep., vol. 18, no. 7, pp. 1774–1790, 2017.

[68] V. Juskaite, D. S. Corcoran, and B. Leitinger, “Collagen induces activation of

DDR1 through lateral dimer association and phosphorylation between dimers,”

Elife, vol. 6, p. e25716, Jun. 2017.

[69] L. A. Flynn, A. R. Blissett, E. P. Calomeni, and G. Agarwal, “Inhibition of

collagen fibrillogenesis by cells expressing soluble extracellular domains of DDR1

and DDR2,” J. Mol. Biol., vol. 395, no. 3, pp. 533–543, 2009.

[70] L. Sivakumar and G. Agarwal, “The influence of discoidin domain receptor 2 on

the persistence length of collagen type I fibers.,” Biomaterials, vol. 31, no. 18, pp.

4802–8, Jun. 2010.

[71] J. R. Tonniges, B. Albert, E. P. Calomeni, S. Roy, J. Lee, X. Mo, S. E. Cole, and 104

G. Agarwal, “Collagen Fibril Ultrastructure in Mice Lacking Discoidin Domain

Receptor 1,” Microsc. Microanal., vol. 22, no. 3, pp. 599–611, Jun. 2016.

[72] G. Agarwal, C. Mihai, and D. F. Iscru, “Interaction of discoidin domain receptor 1

with collagen type 1.,” J. Mol. Biol., vol. 367, no. 2, pp. 443–455, 2007.

[73] D. Kim, P. Ko, E. You, and S. Rhee, “The intracellular juxtamembrane domain of

discoidin domain receptor 2 (DDR2) is essential for receptor activation and

DDR2-mediated cancer progression,” Int. J. Cancer, vol. 135, no. 11, pp. 2547–

2557, 2014.

[74] C. L. Jia, Z. J. Zhou, R. C. Liu, S. D. Chen, and R. H. Xia, “EGF receptor

clustering is induced by a 0.4 mT power frequency magnetic field and blocked by

the EGF receptor tyrosine kinase inhibitor PD153035,” Bioelectromagnetics, vol.

28, no. 3, pp. 197–207, 2007.

[75] A. Sebollela, G. M. Mustata, K. Luo, P. T. Velasco, K. L. Viola, E. N. Cline, G. S.

Shekhawat, K. C. Wilcox, V. P. Dravid, and W. L. Klein, “Elucidating molecular

mass and shape of a neurotoxic a?? oligomer,” ACS Chem. Neurosci., vol. 5, no.

12, pp. 1238–1245, 2014.

[76] M. Hilsch, B. Goldenbogen, C. Sieben, C. T. Höfer, J. P. Rabe, E. Klipp, A.

Herrmann, and S. Chiantia, “Influenza a matrix protein m1 multimerizes upon

binding to lipid membranes,” Biophys. J., vol. 107, no. 4, pp. 912–923, 2014.

[77] D. Pogoryelov, C. Reichen, A. L. Klyszejko, R. Brunisholz, D. J. Muller, P.

Dimroth, and T. Meier, “The oligomeric state of c rings from cyanobacterial F-

ATP synthases varies from 13 to 15,” J. Bacteriol., vol. 189, no. 16, pp. 5895–

5902, 2007. 105

[78] D. N. Petsev, B. R. Thomas, S.-T. Yau, and P. G. Vekilov, “Interactions and

Aggregation of Apoferritin Molecules in Solution: Effects of Added Electrolytes,”

Biophys. J., vol. 78, no. 4, pp. 2060–2069, 2000.

[79] D. Yeung, D. Chmielewski, C. Mihai, and G. Agarwal, “Oligomerization of DDR1

ECD affects receptor-ligand binding.,” J. Struct. Biol., vol. 183, no. 3, pp. 495–

500, Sep. 2013.

[80] K. Ikeda, L.-H. Wang, R. Torres, H. Zhao, E. Olaso, F. J. Eng, P. Labradorʈ, R.

Kleinʈ, D. Lovett, G. D. Yancopoulos, S. L. Friedman, and H. C. Lin, “Discoidin

Domain Receptor 2 Interacts with Src and Shc following Its Activation by Type I

Collagen*,” 2002.

[81] K. Yang, J. H. Kim, H. J. Kim, I.-S. Park, I. Y. Kim, and B.-S. Yang, “Tyrosine

740 Phosphorylation of Discoidin Domain Receptor 2 by Src Stimulates

Intramolecular Autophosphorylation and Shc Signaling Complex Formation *,”

2005.

[82] J. Dejmek, K. Dib, M. Jönsson, and T. Andersson, “Wnt-5a and G-protein

signaling are required for collagen-induced DDR1 receptor activation and normal

mammary cell adhesion,” Int. J. Cancer, vol. 103, no. 3, pp. 344–351, Jan. 2003.

[83] K. K. Lu, D. Trcka, and M. P. Bendeck, “Collagen stimulates discoidin domain

receptor 1-mediated migration of smooth muscle cells through Src.,” Cardiovasc.

Pathol., vol. 20, no. 2, pp. 71–6, 2011.

[84] X. Li, A. G. Garrity, H. Xu, and H. Xu, “Regulation of membrane trafficking by

signalling on endosomal and lysosomal membranes,” J Physiol, vol. 0, pp. 1–13,

2013. 106

[85] T. Stasyk and L. A. Huber, “Spatio-Temporal Parameters of Endosomal Signaling

in Cancer: Implications for New Treatment Options,” J. Cell. Biochem., vol. 117,

no. 4, pp. 836–843, Apr. 2016.

[86] Q.-F. Li, A. M. Spinelli, R. Wang, Y. Anfinogenova, H. A. Singer, and D. D.

Tang, “Critical Role of Vimentin Phosphorylation at Ser-56 by p21-activated

Kinase in Vimentin Cytoskeleton Signaling *,” 2006.

[87] P. Canning, L. Tan, K. Chu, S. W. Lee, N. S. Gray, and A. N. Bullock, “Structural

Mechanisms Determining Inhibition of the Collagen Receptor DDR1 by Selective

and Multi-Targeted Type II Kinase Inhibitors,” 2014.

[88] M. Doube, M. M. Kłosowski, I. Arganda-Carreras, F. P. Cordelières, R. P.

Dougherty, J. S. Jackson, B. Schmid, J. R. Hutchinson, and S. J. Shefelbine,

“BoneJ: Free and extensible bone image analysis in ImageJ,” Bone, vol. 47, pp.

1076–1079, 2010.

[89] J. C. Smith, “Experimental values for the elastic constants of a particulate-filled

glassy polymer,” J. Res. Natl. Bur. Stand. Sect. A Phys. Chem., vol. 80A, no. 1, p.

45, 1976.

[90] R. Fridman and P. H. Huang, Discoidin domain receptors in health and disease.

2016.

[91] C. A. R. Jones, M. Cibula, J. Feng, E. A. Krnacik, D. H. Mcintyre, H. Levine, B.

Sun, F. C. Mackintosh, and J. L. Ross, “Micromechanics of cellularized

biopolymer networks.”

[92] L. Y. Leung, D. Tian, C. P. Brangwynne, D. A. Weitz, and D. J. Tschumperlin, “A

new microrheometric approach reveals individual and cooperative roles for TGF- 107

β1 and IL-β1 in fibroblast-mediated stiffening of collagen gels.,” FASEB J., vol.

21, no. 9, pp. 2064–2073, 2007.

[93] M. Keating, A. Kurup, M. Alvarez-Elizondo, A. J. Levine, and E. Botvinick,

“Spatial distributions of pericellular stiffness in natural extracellular matrices are

dependent on cell-mediated proteolysis and contractility,” Acta Biomater., vol. 57,

pp. 304–312, 2017.

[94] D. P. Jones, W. Hanna, H. El-Hamidi, and J. P. Celli, “Longitudinal measurement

of extracellular matrix rigidity in 3D tumor models using particle-tracking

microrheology.,” J. Vis. Exp., no. 88, Jan. 2014.

[95] J. R. Staunton, W. Vieira, K. L. Fung, R. Lake, A. Devine, and K. Tanner,

“Mechanical Properties of the Tumor Stromal Microenvironment Probed In Vitro

and Ex Vivo by In Situ-Calibrated Optical Trap-Based Active Microrheology,”

Cell. Mol. Bioeng., vol. 9.

[96] C. A. R. Jones, M. Cibula, J. Feng, E. A. Krnacik, D. H. McIntyre, H. Levine, and

B. Sun, “Micromechanics of cellularized biopolymer networks.,” Proc. Natl. Acad.

Sci. U. S. A., vol. 112, no. 37, pp. E5117-22, Sep. 2015.

[97] J. Xie, M. Bao, S. M. C. Bruekers, and W. T. S. Huck, “Collagen Gels with

Different Fibrillar Microarchitectures Elicit Different Cellular Responses,” ACS

Appl. Mater. Interfaces, vol. 9, no. 23, pp. 19630–19637, 2017.

[98] C. A. Mullen, T. J. Vaughan, K. L. Billiar, and L. M. Mcnamara, “The Effect of

Substrate Stiffness, Thickness, and Cross-Linking Density on Osteogenic Cell

Behavior,” Biophysj, vol. 108, pp. 1604–1612, 2015.

[99] M. D. Stevenson, H. Piristine, N. J. Hogrebe, T. M. Nocera, M. W. Boehm, R. K. 108

Reen, K. W. Koelling, G. Agarwal, A. L. Sarang-Sieminski, and K. J. Gooch, “A

self-assembling peptide matrix used to control stiffness and binding site density

supports the formation of microvascular networks in three dimensions,” Acta

Biomater., vol. 9, no. 8, pp. 7651–7661, 2013.

[100] D. Velegol and F. Lanni, “Cell traction forces on soft biomaterials. I.

Microrheology of type I collagen gels.,” Biophys. J., vol. 81, no. 3, pp. 1786–92,

Sep. 2001.

[101] M. Shayegan and N. R. Forde, “Microrheological characterization of collagen

systems: from molecular solutions to fibrillar gels.,” PLoS One, vol. 8, no. 8, p.

e70590, Jan. 2013.

[102] O. Latinovic, L. a Hough, and H. Daniel Ou-Yang, “Structural and

micromechanical characterization of type I collagen gels.,” J. Biomech., vol. 43,

no. 3, pp. 500–5, Feb. 2010.

[103] B. E. Henslee, A. Morss, X. Hu, G. P. Lafyatis, and L. J. Lee, “Electroporation

Dependence on Cell Size: Optical Tweezers Study,” | Anal. Chem, vol. 83, pp.

3998–4003, 2011.

[104] J. C. Crocker and B. D. Hoffman, “Multiple Particle Tracking and Two-Point

Microrheology in Cells,” Part Cell Biol. Commons Postprint version. Publ.

Methods Cell Biol. Methods Cell Biol., vol. 83, no. 83, pp. 141–178, 2007.

[105] N. Phan-Thien and S. Kim, Microstructures in elastic media : principles and

computational methods, 1st ed. Oxford: Oxford University Press, 1994.

[106] F. G. Schmidt, F. Ziemann, and E. Sackmann, “Shear field mapping in actin

networks by using magnetic tweezers.,” Eur. Biophys. J., vol. 24, no. 5, pp. 348– 109

353, 1996.

[107] A. R. Bausch, W. Möller, and E. Sackmann, “Measurement of local viscoelasticity

and forces in living cells by magnetic tweezers.,” Biophys. J., vol. 76, no. 1 Pt 1,

pp. 573–9, Jan. 1999.

[108] M. H. Lee and E. M. Furst, “Response of a colloidal gel to a microscopic

oscillatory strain,” Phys. Rev. E, vol. 77, no. 4, p. 41408, Apr. 2008.

[109] L. A. Hough and H. D. Ou-Yang, “Correlated motions of two hydrodynamically

coupled particles confined in separate quadratic potential wells,” Phys. Rev. E -

Stat. Physics, Plasmas, Fluids, Relat. Interdiscip. Top., vol. 65, no. 2, 2002.

[110] T. Paust, T. Neckernuss, L. K. Mertens, I. Martin, M. Beil, P. Walther, T.

Schimmel, and O. Marti, “Active multi-point microrheology of cytoskeletal

networks,” Beilstein J. Nanotechnol., vol. 7, no. 1, pp. 484–491, 2016.

[111] M. T. Valentine, Z. E. Perlman, M. L. Gardel, J. H. Shin, P. Matsudaira, T. J.

Mitchison, and D. A. Weitz, “Colloid surface chemistry critically affects multiple

particle tracking measurements of biomaterials,” Biophys. J., vol. 86, no. 6, pp.

4004–4014, 2004.

[112] J. C. Crocker, M. T. Valentine, E. R. Weeks, T. Gisler, P. D. Kaplan, A. G. Yodh,

and D. A. Weitz, “Two-point microrheology of inhomogeneous soft materials,”

Phys. Rev. Lett., vol. 85, no. 4, pp. 888–891, 2000.

[113] A. J. Levine and T. C. Lubensky, “One- and two-particle microrheology,” Phys.

Rev. Lett., vol. 85, no. 8, pp. 1774–1777, 2000.

[114] M. Atakhorrami, G. H. Koenderink, J. F. Palierne, F. C. MacKintosh, and C. F.

Schmidt, “Scale-Dependent Nonaffine Elasticity of Semiflexible Polymer 110

Networks,” Phys. Rev. Lett., vol. 112, no. 8, p. 88101, Feb. 2014.

[115] D. Vader, A. Kabla, D. Weitz, and L. Mahadevan, “Strain-Induced Alignment in

Collagen Gels,” PLoS One, vol. 4, no. 6, 2009.

[116] A. A. Tomei, F. Boschetti, F. Gervaso, and M. A. Swartz, “3D collagen cultures

under well-defined dynamic strain: A novel strain device with a porous

elastomeric support,” Biotechnol. Bioeng., vol. 103, no. 1, pp. 217–225, 2009.

[117] C. B. Raub, A. J. Putnam, B. J. Tromberg, and S. C. George, “Predicting bulk

mechanical properties of cellularized collagen gels using multiphoton

microscopy,” Acta Biomater., vol. 6, no. 12, pp. 4657–4665, 2010.

[118] C. B. Raub, J. Unruh, V. Suresh, T. Krasieva, T. Lindmo, E. Gratton, B. J.

Tromberg, and S. C. George, “Image Correlation Spectroscopy of Multiphoton

Images Correlates with Collagen Mechanical Properties,” Biophys. J., vol. 94, pp.

2361–2373.

[119] E. D. Foehr, A. Tatavos, E. Tanabe, S. Raffioni, S. Goetz, E. Dimarco, M. De

Luca, and R. a Bradshaw, “Discoidin domain receptor 1 (DDR1) signaling in

PC12 cells: activation of juxtamembrane domains in PDGFR/DDR/TrkA chimeric

receptors.,” FASEB J., vol. 14, no. 7, pp. 973–81, May 2000.

[120] T. Wehrman, X. He, B. Raab, A. Dukipatti, H. Blau, and K. C. Garcia, “Structural

and Mechanistic Insights into Nerve Growth Factor Interactions with the TrkA and

p75 Receptors,” Neuron, vol. 53, no. 1, pp. 25–38, 2007.

[121] H. Liu, X. Chen, P. J. Focia, and X. He, “Structural basis for stem cell factor–KIT

signaling and activation of class III receptor tyrosine kinases,” EMBO J., vol. 26,

pp. 891–901, 2007. 111

[122] S. Yuzawa, Y. Opatowsky, Z. Zhang, V. Mandiyan, I. Lax, and J. Schlessinger,

“Structural Basis for Activation of the Receptor Tyrosine Kinase KIT by Stem Cell

Factor.”

[123] D. J. Stauber, A. D. DiGabriele, and W. A. Hendrickson, “Structural interactions

of fibroblast growth factor receptor with its ligands.,” Proc. Natl. Acad. Sci. U. S.

A., vol. 97, no. 1, pp. 49–54, 2000.

[124] C. Ward, M. Lawrence, V. Streltsov, T. Garrett, N. McKern, M. Z. Lou, G.

Lovrecz, and T. Adams, “Structural insights into ligand-induced activation of the

insulin receptor,” in Acta Physiologica, 2008, vol. 192, no. 1, pp. 3–9.

[125] T. Moriki, H. Maruyama, and I. N. Maruyama, “Activation of preformed EGF

receptor dimers by ligand-induced rotation of the transmembrane domain,” J. Mol.

Biol., vol. 311, no. 5, pp. 1011–1026, Aug. 2001.

[126] R.-H. Tao and I. N. Maruyama, “All EGF(ErbB) receptors have preformed homo-

and heterodimeric structures in living cells.,” J. Cell Sci., vol. 121, no. Pt 19, pp.

3207–17, Oct. 2008.

[127] J. Shen and I. N. Maruyama, “Nerve growth factor receptor TrkA exists as a

preformed, yet inactive, dimer in living cells,” FEBS Lett., vol. 585, no. 2, pp.

295–299, Jan. 2011.

[128] J. R. Cochran, T. O. Cameron, and L. J. Stern, “The Relationship of MHC-Peptide

Binding and T Cell Activation Probed Using Chemically Defined MHC Class II

Oligomers,” Immunity, vol. 12, no. 3, pp. 241–250, Mar. 2000.

[129] L. L. Kiessling, J. E. Gestwicki, and L. E. Strong, “Synthetic Multivalent Ligands

as Probes of Signal Transduction,” Angew. Chemie Int. Ed., vol. 45, no. 15, pp. 112

2348–2368, Apr. 2006.

[130] D. Stabley, S. Retterer, S. Marshall, and K. Salaita, “Manipulating the lateral

diffusion of surface-anchored EGF demonstrates that receptor clustering modulates

phosphorylation levels,” Integr. Biol., vol. 5, no. 4, p. 659, 2013.

[131] W. A. Barton, D. Tzvetkova-Robev, E. P. Miranda, M. V Kolev, K. R.

Rajashankar, J. P. Himanen, and D. B. Nikolov, “Crystal structures of the Tie2

receptor ectodomain and the angiopoietin-2–Tie2 complex,” Nat. Struct. Mol.

Biol., vol. 13, no. 6, pp. 524–532, Jun. 2006.

[132] D. Boettiger, “Mechanical control of integrin-mediated adhesion and signaling,”

Curr. Opin. Cell Biol., vol. 24, no. 5, pp. 592–599, Oct. 2012.

[133] M. Triantafilou, F. G. J. Gamper, R. M. Haston, M. A. Mouratis, S. Morath, T.

Hartung, and K. Triantafilou, “Membrane sorting of toll-like receptor (TLR)-2/6

and TLR2/1 heterodimers at the cell surface determines heterotypic associations

with CD36 and intracellular targeting,” J. Biol. Chem., vol. 281, no. 41, pp.

31002–31011, 2006.

[134] Y. Yarden and M. X. Sliwkowski, “Untangling the ErbB signalling network,” Nat.

Rev. Mol. Cell Biol., vol. 2, no. 2, pp. 127–137, Feb. 2001.

[135] K. Salaita, P. M. Nair, R. S. Petit, R. M. Neve, D. Das, J. W. Gray, and J. T.

Groves, “Restriction of receptor movement alters cellular response: Physical force

sensing by EphA2,” Science (80-. )., vol. 327, no. 5971, pp. 1380–1385, 2010.

[136] A. Abulrob, Z. Lu, E. Baumann, D. Vobornik, R. Taylor, D. Stanimirovic, and L.

J. Johnston, “Nanoscale imaging of epidermal growth factor receptor clustering:

Effects of inhibitors,” J. Biol. Chem., vol. 285, no. 5, pp. 3145–3156, 2010. 113

[137] G. Carpenter and S. Cohen, “125I-labeled human epidermal growth factor.

Binding, internalization, and degradation in human fibroblasts.,” J. Cell Biol., vol.

71, no. 1, pp. 159–71, 1976.

[138] K. M. Ferguson, M. B. Berger, J. M. Mendrola, H.-S. Cho, D. J. Leahy, and M. A.

Lemmon, “EGF Activates Its Receptor by Removing Interactions that Autoinhibit

Ectodomain Dimerization,” Mol. Cell, vol. 11, pp. 507–517, 2003.

[139] E. R. Purba, E.-I. Saita, and I. N. Maruyama, “Activation of the EGF Receptor by

Ligand Binding and Oncogenic Mutations: The " Rotation Model "”

[140] E. Bogdanovic, V. P. K. H. Nguyen, and D. J. Dumont, “Activation of Tie2 by

angiopoietin-1 and angiopoietin-2 results in their release and receptor

internalization,” J. Cell Sci., vol. 119, no. 17, pp. 3551–3560, 2006.

[141] D. Lepzelter, O. Bates, and M. Zaman, “Integrin clustering in two and three

dimensions,” Langmuir, vol. 28, no. 12, pp. 5379–5386, 2012.

[142] R. E. Bridgewater, J. C. Norman, and P. T. Caswell, “Integrin trafficking at a

glance,” J. Cell Sci., vol. 125, pp. 3695–3701.

[143] C. G. Gahmberg, S. C. Fagerholm, S. M. Nurmi, T. Chavakis, S. Marchesan, and

M. Grönholm, “Regulation of integrin activity and signalling,” BBA - Gen. Subj.,

vol. 1790, pp. 431–444, 2009.

[144] W. A. Barton, A. C. Dalton, T. C. M. Seegar, J. P. Himanen, and D. B. Nikolov,

“Tie2 and Eph Receptor Tyrosine Kinase Activation and Signaling,” Cold Spring

Harb. Perspect. Biol., vol. 6, no. 3, 2014.

[145] J. P. Himanen, L. Yermekbayeva, P. W. Janes, J. R. Walker, K. Xu, L. Atapattu,

K. R. Rajashankar, A. Mensinga, M. Lackmann, D. B. Nikolov, and S. Dhe- 114

Paganon, “Architecture of Eph receptor clusters,” Proc. Natl. Acad. Sci., vol. 107,

no. 24, pp. 10860–10865, 2010.

[146] C. A. Bell, J. A. Tynan, K. C. Hart, A. N. Meyer, S. C. Robertson, and D. J.

Donoghue, “Rotational Coupling of the Transmembrane and Kinase Domains of

the Neu Receptor Tyrosine Kinase,” Mol. Biol. Cell, vol. 11, pp. 3589–3599, 2000.

[147] M. J. Sternberg and W. J. Gullick, “A sequence motif in the transmembrane region

of growth factor receptors with tyrosine kinase activity mediates dimerization.,”

Protein Eng., vol. 3, no. 4, pp. 245–8, Mar. 1990.

[148] W. P. Russ and D. M. Engelman, “The GxxxG motif: A framework for

transmembrane helix-helix association,” J. Mol. Biol., vol. 296, no. 3, pp. 911–

919, Feb. 2000.

[149] A. Senes, M. Gerstein, and D. M. Engelman, “Statistical analysis of amino acid

patterns in transmembrane helices: the GxxxG motif occurs frequently and in

association with β-branched residues at neighboring positions,” J. Mol. Biol., vol.

296, no. 3, pp. 921–936, Feb. 2000.

[150] J. M. Mendrola, M. B. Berger, M. C. King, and M. A. Lemmon, “The single

transmembrane domains of ErbB receptors self-associate in cell membranes,” J.

Biol. Chem., vol. 277, no. 7, pp. 4704–4712, 2002.

[151] N. a Noordeen, F. Carafoli, E. Hohenester, M. a Horton, and B. Leitinger, “A

transmembrane leucine zipper is required for activation of the dimeric receptor

tyrosine kinase DDR1.,” J. Biol. Chem., vol. 281, no. 32, pp. 22744–51, Aug.

2006.

[152] W.-J. Chen, J. L. Goldstein, and M. S. Brown, “NPXY, a Sequence Often Found in 115

Cytoplasmic Tails, Is Required for Coated Pit-mediated Internalization of the Low

Density Lipoprotein Receptor*,” J. Biol. Chem., vol. 265, no. 6, 1990.

[153] J. M. Backer, S. E. Shoelson, M. A. Weiss, Q. X. Hua, R. B. Cheatham, E. Haring,

D. C. Cahill, and M. F. White, “The insulin receptor juxtamembrane region

contains two independent tyrosine/beta-turn internalization signals.,” J. Cell Biol.,

vol. 118, no. 4, pp. 831–9, Aug. 1992.

[154] D. Prager, H.-L. Li, H. Yamasaki, and S. Melmeds, “Human Insulin-like Growth

Factor I Receptor Internalization ROLE OF THE JUXTAMEMBRANE

DOMAIN*,” vol. 269, no. 16, pp. 11934–11937, 1994.

[155] H. Wu, D. A. Windmiller, L. Wang, and J. M. Backer, “Y XX M Motifs in the

PDGF-β Receptor Serve Dual Roles as Phosphoinositide 3-Kinase Binding Motifs

and Tyrosine-based Endocytic Sorting Signals,” J. Biol. Chem., vol. 278, no. 42,

pp. 40425–40428, Oct. 2003.

[156] G. M. Di Guglielmo, P. C. Baass, W. J. Ou, B. I. Posner, and J. J. Bergeron,

“Compartmentalization of SHC, GRB2 and mSOS, and hyperphosphorylation of

Raf-1 by EGF but not insulin in liver parenchyma.,” EMBO J., vol. 13, no. 18, pp.

4269–77, 1994.

[157] M. Miaczynska, L. Pelkmans, and M. Zerial, “Not just a sink: Endosomes in

control of signal transduction,” Current Opinion in Cell Biology, vol. 16, no. 4.

Elsevier Current Trends, pp. 400–406, 01-Aug-2004.

[158] M. von Zastrow and A. Sorkin, “Signaling on the endocytic pathway,” Current

Opinion in Cell Biology, vol. 19, no. 4. pp. 436–445, 2007.

[159] A. Juin, J. Di Martino, B. Leitinger, E. Henriet, A.-S. Gary, L. Paysan, J. Bomo, G. 116

Baffet, C. Gauthier-Rouvière, J. Rosenbaum, V. Moreau, and F. Saltel, “Discoidin

domain receptor 1 controls linear invadosome formation via a Cdc42–Tuba

pathway,” J. Cell Biol., vol. 207, no. 4, pp. 517–533, Nov. 2014.

[160] J. Jokinen, E. Dadu, P. Nykvist, J. Käpylä, D. J. White, J. Ivaska, P. Vehviläinen,

H. Reunanen, H. Larjava, L. Häkkinen, and J. Heino, “Integrin-mediated cell

adhesion to type I collagen fibrils.,” J. Biol. Chem., vol. 279, no. 30, pp. 31956–63,

Jul. 2004.

[161] J. Emsley, C. G. Knight, R. W. Farndale, M. J. Barnes, and R. C. Liddington,

“Structural Basis of Collagen Recognition by Integrin α2β1,” Cell, vol. 101, no. 1,

pp. 47–56, 2000.

[162] C. V Carman and T. A. Springer, “Integrin avidity regulation: are changes in

affinity and conformation underemphasized?,” Curr. Opin. Cell Biol., vol. 15, no.

5, pp. 547–556, Oct. 2003.

[163] D. R. Critchley and A. R. Gingras, “Talin at a glance,” J. Cell Sci., vol. 121, no. 9,

pp. 1345–1347, May 2008.

[164] C. Wu, “The PINCH-ILK-parvin complexes: Assembly, functions and regulation,”

Biochim. Biophys. Acta - Mol. Cell Res., vol. 1692, no. 2–3, pp. 55–62, 2004.

[165] C. Wu, “PINCH, N(i)ck and the ILK: Network wiring at cell-matrix adhesions,”

Trends in Cell Biology, vol. 15, no. 9. Elsevier, pp. 460–466, 01-Sep-2005.

[166] R. Li, N. Mitra, H. Gratkowski, G. Vilaire, R. Litvinov, C. Nagasami, J. W.

Weisel, J. D. Lear, W. F. DeGrado, and J. S. Bennett, “Activation of integrin

alphaIIbbeta3 by modulation of transmembrane helix associations.,” Science, vol.

300, no. 5620, pp. 795–8, May 2003. 117

[167] D. F. Kucik, “Rearrangement of Integrins in Avidity Regulation by Leukocytes,”

Immunol. Res., vol. 263, no. 1, pp. 199–206, 2002.

[168] P. T. Caswell and J. C. Norman, “Integrin trafficking and the control of cell

migration,” Traffic, vol. 7, no. 1, pp. 14–21, 2006.

[169] E. Puklin-Faucher and M. P. Sheetz, “The mechanical integrin cycle,” J. Cell Sci.,

vol. 122, no. 4, pp. 575–575, 2009.

[170] B. An and B. Brodsky, “Collagen binding to OSCAR: The odd couple,” Blood,

vol. 127, no. 5. American Society of Hematology, pp. 521–522, 04-Feb-2016.

[171] M. Moroi and S. M. Jung, “Platelet glycoprotein VI: its structure and function.,”

Thromb. Res., vol. 114, no. 4, pp. 221–33, Jan. 2004.

[172] K. Horii, M. L. Kahn, and A. B. Herr, “Structural basis for platelet collagen

responses by the immune-type receptor glycoprotein VI.,” Blood, vol. 108, no. 3,

pp. 936–42, Aug. 2006.

[173] J. F. Arthur, S. Dunkley, and R. K. Andrews, “Platelet glycoprotein VI-related

clinical defects,” Br. J. Haematol., vol. 139, no. 3, pp. 363–372, Oct. 2007.

[174] Y. Miura, T. Takahashi, S. M. Jung, and M. Moroi, “Analysis of the Interaction of

Platelet Collagen Receptor Glycoprotein VI (GPVI) with Collagen A DIMERIC

FORM OF GPVI, BUT NOT THE MONOMERIC FORM, SHOWS AFFINITY

TO FIBROUS COLLAGEN*,” 2002.

[175] M. J. M. O. Nordkamp, J. A. G. Van Roon, M. Douwes, T. De Ruiter, R. T.

Urbanus, and L. Meyaard, “Enhanced secretion of leukocyte-associated

immunoglobulin-like receptor 2 (LAIR-2) and soluble LAIR-1 in rheumatoid

arthritis: LAIR-2 is a more efficient antagonist of the LAIR-1-collagen inhibitory 118

interaction than is soluble LAIR-1,” Arthritis Rheum., vol. 63, no. 12, pp. 3749–

3757, 2011.

[176] A. D. Barrow, N. Raynal, T. L. Andersen, D. A. Slatter, D. Bihan, N. Pugh, M.

Cella, T. Kim, J. Rho, T. Negishi-Koga, J. M. Delaisse, H. Takayanagi, J. Lorenzo,

M. Colonna, R. W. Farndale, Y. Choi, and J. Trowsdale, “OSCAR is a collagen

receptor that costimulates osteoclastogenesis in DAP12-deficient humans and

mice,” J. Clin. Invest., vol. 121, no. 9, pp. 3505–3516, 2011.

[177] J. D. Klemm, S. L. Schreiber, and G. R. Crabtree, “DIMERIZATION AS A

REGULATORY MECHANISM IN SIGNAL TRANSDUCTION,” Annu. Rev.

Immunol, vol. 16, pp. 569–92, 1998.

[178] M. R. Morgan, M. J. Humphries, and M. D. Bass, “Synergistic control of cell

adhesion by integrins and syndecans,” Nat. Rev. Mol. Cell Biol., vol. 8, no. 12, pp.

957–969, 2007.

[179] J. Huxley - Jones, D. L. Robertson, and R. P. Boot-Handford, “On the origins of

the extracellular matrix in vertebrates,” Matrix Biol., vol. 26, no. 1, pp. 2–11, Jan.

2007.

[180] K. G. Danielson, H. Baribault, D. F. Holmes, H. Graham, K. E. Kadler, and R. V

Iozzo, “Targeted disruption of decorin leads to abnormal collagen fibril

morphology and skin fragility.,” J. Cell Biol., vol. 136, no. 3, pp. 729–43, Feb.

1997.

[181] Z. Ferdous, L. D. Lazaro, R. V Iozzo, M. Höök, and K. J. Grande-Allen,

“Influence of cyclic strain and decorin deficiency on 3D cellularized collagen

matrices,” Biomaterials, vol. 29, no. 18, pp. 2740–2748, 2008. 119

[182] S. Goldoni and R. V. Iozzo, “Tumor microenvironment: Modulation by decorin

and related molecules harboring leucine-rich tandem motifs,” Int. J. Cancer, vol.

123, no. 11, pp. 2473–2479, Dec. 2008.

[183] H. Järveläinen, P. Puolakkainen, S. Pakkanen, E. L. Brown, M. Höök, R. V. Iozzo,

E. H. Sage, and T. N. Wight, “A role for decorin in cutaneous wound healing and

angiogenesis,” Wound Repair Regen., vol. 14, no. 4, pp. 443–452, Jul. 2006.

[184] I. T. Weber, R. W. Harrison, and R. V Iozzo, “Model structure of decorin and

implications for collagen fibrillogenesis.,” J. Biol. Chem., vol. 271, no. 50, pp.

31767–70, Dec. 1996.

[185] P. G. Scott, J. G. Grossmann, C. M. Dodd, J. K. Sheehan, and P. N. Bishop, “Light

and X-ray scattering show decorin to be a dimer in solution.,” J. Biol. Chem., vol.

278, no. 20, pp. 18353–9, May 2003.

[186] P. G. Scott, P. A. McEwan, C. M. Dodd, E. M. Bergmann, P. N. Bishop, and J.

Bella, “Crystal structure of the dimeric protein core of decorin, the archetypal

small leucine-rich repeat proteoglycan.,” Proc. Natl. Acad. Sci. U. S. A., vol. 101,

no. 44, pp. 15633–8, Nov. 2004.

[187] P. A. McEwan, P. G. Scott, P. N. Bishop, and J. Bella, “Structural correlations in

the family of small leucine-rich repeat proteins and proteoglycans,” J. Struct. Biol.,

vol. 155, no. 2, pp. 294–305, Aug. 2006.

[188] J. P. R. O. Orgel, A. Eid, O. Antipova, J. Bella, and J. E. Scott, “Decorin core

protein (decoron) shape complements collagen fibril surface structure and

mediates its binding,” PLoS One, vol. 4, no. 9, 2009.

[189] C.-M. Lo, H.-B. Wang, M. Dembo, and Y. Wang, “Cell Movement Is Guided by 120

the Rigidity of the Substrate,” Biophys. J., vol. 79, no. 1, pp. 144–152, Jul. 2000.

[190] S. R. Peyton and A. J. Putnam, “Extracellular matrix rigidity governs smooth

muscle cell motility in a biphasic fashion,” J. Cell. Physiol., vol. 204, no. 1, pp.

198–209, Jul. 2005.

[191] C. S. Chen, M. Mrksich, S. Huang, G. M. Whitesides, and D. E. Ingber,

“Geometric control of cell life and death.,” Science, vol. 276, no. 5317, pp. 1425–

8, May 1997.

[192] J. D. Mih, A. Marinkovic, F. Liu, A. S. Sharif, and D. J. Tschumperlin, “Matrix

stiffness reverses the effect of actomyosin tension on cell proliferation.,” J. Cell

Sci., vol. 125, no. Pt 24, pp. 5974–83, Dec. 2012.

[193] E. Hadjipanayi, V. Mudera, and R. A. Brown, “Close dependence of fibroblast

proliferation on collagen scaffold matrix stiffness,” J. Tissue Eng. Regen. Med.,

vol. 3, no. 2, pp. 77–84, Feb. 2009.

[194] R. J. Pelham and Y.-L. Wang, “Cell locomotion and focal adhesions are regulated

by substrate flexibility,” Cell Biol., vol. 94, pp. 13661–13665, 1997.

[195] T. Yeung, P. C. Georges, L. A. Flanagan, B. Marg, M. Ortiz, M. Funaki, N. Zahir,

W. Ming, V. Weaver, and P. A. Janmey, “Effects of substrate stiffness on cell

morphology, cytoskeletal structure, and adhesion,” Cell Motil. Cytoskeleton, vol.

60, no. 1, pp. 24–34, Jan. 2005.

[196] A. Engler, L. Bacakova, C. Newman, A. Hategan, M. Griffin, and D. Discher,

“Substrate Compliance versus Ligand Density in Cell on Gel Responses,” Biophys.

J., vol. 86, pp. 617–628, 2004.

[197] A. S. Rowlands, P. A. George, and J. J. Cooper-White, “Directing osteogenic and 121

myogenic differentiation of MSCs: interplay of stiffness and adhesive ligand

presentation,” Am. J. Physiol. Physiol., vol. 295, no. 4, pp. C1037–C1044, Oct.

2008.

[198] Y. Sun, C. S. Chen, and J. Fu, “Forcing Stem Cells to Behave: A Biophysical

Perspective of the Cellular Microenvironment,” Annu. Rev. Biophys., vol. 41, no.

1, pp. 519–542, 2012.

[199] W. L. Murphy, T. C. McDevitt, and A. J. Engler, “Materials as stem cell

regulators,” Nat. Mater., vol. 13, no. 6, pp. 547–557, Jun. 2014.

[200] E. Olaso, K. Ikeda, and F. Eng, “DDR2 receptor promotes MMP-2–mediated

proliferation and invasion by hepatic stellate cells,” J. Clin. …, vol. 108, no. 9, pp.

1369–1378, 2001.

[201] C. A. S. Corsa, A. Brenot, W. R. Grither, S. Van Hove, A. J. Loza, K. Zhang, S.

M. Ponik, Y. Liu, D. G. Denardo, K. W. Eliceiri, P. J. Keely, and G. D. Longmore,

“The action of Discoidin Domain Receptor 2 in basal tumor cells and stromal

Cancer Associated Fibroblasts is critical for breast cancer metastasis.”

[202] R. Parenteau-Bareil, R. Gauvin, and F. Berthod, “Collagen-based biomaterials for

tissue engineering applications,” Materials (Basel)., vol. 3, no. 3, pp. 1863–1887,

2010.

[203] S. P. Reese, C. J. Underwood, and J. a Weiss, “Effects of decorin proteoglycan on

fibrillogenesis, ultrastructure, and mechanics of type I collagen gels.,” Matrix

Biol., vol. 32, no. 7–8, pp. 414–23, 2013.

[204] J. R. Tonniges, B. Albert, E. P. Calomeni, S. Roy, J. Lee, X. Mo, S. E. Cole, and

G. Agarwal, “Collagen Fibril Ultrastructure in Mice Lacking Discoidin Domain 122

Receptor 1,” Microsc. Microanal, vol. 22, pp. 599–611, 2016.

123