ON THE MECHANOBIOLOGY OF GROWTH AND REMODELLING

A Dissertation Presented

By

Seyed Mohammad Siadat

to

The Department of Bioengineering

in partial fulfillment of the requirements for the degree of

Doctor of Philosophy

in the field of

Bioengineering

Northeastern University Boston, Massachusetts

August 2020 ii

ABSTRACT

How organized collagenous structure can arise and grow from a cluster of cells remains one of the most important basic science questions associated with research. Despite more than a century of research, there are currently no widely accepted mechanistic models of formation, growth, and remodeling of collagen fibrils. It has been hypothesized in our research group that collagen monomers and enzymes are in a dynamic

equilibrium with existing fibrils. Tensile forces on fibrils can shift this equilibrium and

change the balance between molecular association and dissociation. Here, we sought to

answer this question: Does fibril strain promote the molecular assembly of collagen?

To investigate this question, individual collagen fibrils were stretched to 0%, 4%, and 6% strain between two microneedles and exposed to a subthreshold concentration of fluorescently labeled collagen molecules to quantify molecular association onto the stretched fibrils. It was shown that labeled monomers rapidly incorporate onto all tested fibrils and reach a plateau. The time to reach plateau was significantly faster for the stretched fibrils (15.6, 7.0, and 6.0 minutes for fibrils under 0%, 4%, and 6% strain, respectively). Analysis of the fibril intensity and photobleaching data indicated that the

association rate was significantly higher for fibrils under 6% strain compared to fibrils under 0% and 4% strain, increasing the association rate by 100%. It was concluded that mechanical stresses and strains could increase fibril growth by decreasing the activation energy required for reaction between monomers and fibrils and also by setting fibrils in a lower state of energy, increasing the association rate of monomers and fibrils. iii

ACKNOWLEDGEMENTS

I would like to thank my advisor Dr. Jeffrey Ruberti who taught me how to pursue science properly. I learned to develop a hypothesis and try my best to prove it wrong. Thank you for giving me the opportunity to work in your research group even though I was a mechanical engineer and didn’t know much about biology when I started. I would like to thank my parents who have always supported and encouraged me to keep learning. I would like to thank my friends and coworkers - JJ, Ramin, Monica, Ebraheim, Alex, Isabel and everyone else - which made my life as a graduate student fun and exciting. I learned so much from you and had lots of adventures. I would like to especially thank Dr. Paten and

Dr. Susilo. I always benefited and developed ideas from our lunchtime discussions. I would like to thank the Bioengineering department and all the amazing people who work there, especially Susan Wilcox which who was always helpful and her love for soccer was also pleasing. I would like to thank Dr. Michael Jaeggli and Dr. Timothy Lannin who I had the opportunity to work with as a teaching assistance during which I discovered a passion for teaching. Last, but not least, I would like to thank my committee, Dr. Charles Dimarzio and Dr. Chiara Bellini who I benefitted from their expertise and knowledge. Thank you all.

iv

TABLE OF CONTENTS

1 INTRODUCTION ...... 1

1.1 COLLAGEN ...... 3

1.2 SELF-ASSEMBLY ...... 7

1.3 FIBRILLOGENESIS, GROWTH, AND REMODELLING ...... 10

1.3.1 The Site and Mechanism of Initial Fibril Formation ...... 10

1.3.2 Fibril Growth Mechanism ...... 43

1.3.3 Role of Mechanics in Fibrillogenesis, Growth, and Remodeling of Collagenous

Tissue ...... 71

2 DYNAMIC TRACKING OF FLUORESCENTLY LABELED TYPE I COLLAGEN

MOLECULES; DIRECT QUANTIFICATION OF MOLECULAR ASSOCIATION

WITH NATIVE FIBRILS ...... 75

2.1 INTRODUCTION ...... 75

2.1.1 Radioactive and Nonradioactive Isotopically-Labeled Amino Acids ...... 76

2.1.2 Non-collagen Based Probes ...... 77

2.1.3 Endogenous Labeling with GFP ...... 79

2.1.4 Exogenous Labeling ...... 81

2.2 OBJECTIVES AND APPROACHES ...... 84 v

2.3 EXPERIMENTAL METHODS ...... 88

2.3.1 Collagen Labeling ...... 88

2.3.2 Purifying the Labeled Collagen ...... 89

2.3.3 Determining Collagen Concentration and Degree of Labeling ...... 90

2.3.4 Spectrophotometry ...... 91

2.3.5 Scleral Fibril Extraction ...... 91

2.3.6 Quantification ...... 93

2.3.7 Imaging ...... 96

2.3.8 Fibril Diameter Measurement using TEM and DIC ...... 97

2.3.9 Molecular Association of Labeled Monomers with Native Fibrils Experiment

...... 98

2.3.10 Image Analysis for Fluorescence ...... 99

2.3.11 Arrhenius Plot and Activation Energy Measurement ...... 100

2.3.12 Orientation of Collagen Molecules ...... 101

2.3.13 Statistical Information ...... 102

2.4 RESULTS ...... 102

2.4.1 The Degree of Labeling (DOL) ...... 102

2.4.2 De Novo Fibrillogenesis Experiments with Labeled Collagen...... 104 vi

2.4.3 Single Molecule, Multi-label Fluorescence Orientation Microscopy (SMO

Microscopy) ...... 109

2.4.4 Labeled Collagen/Native Fibril Association Experiments ...... 114

2.5 DISCUSSION ...... 118

2.6 SUMMARY ...... 121

3 A CORRELATIVE METHOD TO MEASURE COLLAGEN FIBRIL DIAMETER IN

DIFFERENTIAL INTERFERENCE CONTRAST MICROSCOPY ...... 123

3.1 INTRODUCTION ...... 123

3.2 CURRENT METHODS FOR MEASURING COLLAGEN FIBRIL DIAMETER ...... 124

3.2.1 SAXS ...... 124

3.2.2 TEM ...... 125

3.2.3 SEM ...... 127

3.2.4 AFM ...... 128

3.2.5 SHG ...... 130

3.3 DIC MICROSCOPY ...... 131

3.4 EXPERIMENTAL METHODS ...... 135

3.4.1 Fibril Preparation for TEM-DIC imaging ...... 135

3.4.2 DIC Imaging ...... 136 vii

3.4.3 TEM Processing ...... 136

3.4.4 Microfabrication of Trenches in PDMS Sheets ...... 136

3.4.5 Fibril Preparation for SEM-DIC imaging ...... 137

3.4.6 SEM Processing ...... 139

3.4.7 Image Processing ...... 139

3.4.8 Statistical Information ...... 140

3.5 RESULTS ...... 140

3.5.1 DIC-EIS is sensitive to collagen fibril diameter change ...... 140

3.5.2 DIC-EIS is unaffected by small changes in fibril direction ...... 142

3.5.3 DIC-EIS is linearly correlated to collagen fibril diameter ...... 144

3.5.4 DIC-EIS is capable of live measurements of hydrated fibrils ...... 147

3.5.5 DIC-EIS shows a reversible diameter change due to dehydration ...... 148

3.6 DISCUSSION...... 149

4 EFFECT OF FIBRIL STRAIN ON FIBRIL/MONOMER ASSOCIATION ...... 153

4.1 INTRODUCTION ...... 153

4.1.1 Tensile forces on fibrils can regulate the dynamic equilibrium between

molecular accretion and enzymatic degradation ...... 153 viii

4.1.2 Tensile forces on fibril can regulate the thermodynamics of fibril growth

reaction ...... 155

4.2 EXPERIMENTAL METHODS ...... 157

4.2.1 Fibril Attachment Between Microneedles ...... 157

4.2.2 Fibril Transfer into PBS, Preconditioning, and stretching ...... 160

4.2.3 Label monomer addition and recording ...... 163

4.2.4 Image Processing and Assessment of Incorporation Rate ...... 163

4.2.5 Control Test and Long-Term Experiments ...... 165

4.3 RESULTS ...... 166

4.3.1 Labeled Monomers Incorporate onto the Fibrils ...... 166

4.3.2 Microfibrils Form on the Fibrils between Microneedles ...... 169

4.3.3 Incorporation rate onto 6% strained fibrils remained significantly higher than

0% and 4% strained fibrils ...... 169

4.4 DISCUSSIONS ...... 172

4.5 SUMMARY AND CONCLUSIONS ...... 175

ix

LIST OF FIGURES

FIGURE 1.1. BIOSYNTHESIS OF COLLAGEN INCLUDING HYDROXYLATION, GLYCOSYLATION,

AND DISULFIDE-BOND FORMATION OF THE PROCOLLAGEN IN THE ENDOPLASMIC

RETICULUM. REPRINTED WITH PERMISSION FROM CHEN AND RAGHUNATH (2009) [57].

...... 4

FIGURE 1.2. SCHEMATIC REPRESENTATION OF A CROSS-SECTION THROUGH A COLLAGEN

TRIPLE HELIX. REPRINTED WITH PERMISSION FROM VAN DER REST AND GARRONE

(1991) [63]...... 5

FIGURE 1.3. STRUCTURE OF A TYPE I PROCOLLAGEN MOLECULE. REPRINTED WITH

PERMISSION FROM HOLMES ET AL. (2018) [18]...... 6

FIGURE 1.4. SCHEMATIC OF HIERARCHICAL COLLAGEN STRUCTURE AND ORGANIZATION.

REPRINTED WITH PERMISSION FROM CANELON AND WALLACE (2016) [77]...... 7

FIGURE 1.5. ASYMMETRIC PATTERN OF TYPE I COLLAGEN FIBRIL SHOWING D-BANDING AND

SUB-BANDS LABELED C1, B2, B1, A4-A1, E2, E1, C3, AND C2 FROM LEFT TO RIGHT AS SHOWN

IN PANEL B. A) NEGATIVE AND B) POSITIVE STAINING PATTERN OF A TYPE I COLLAGEN

FIBRIL. C) SCHEMATIC OF THE ORIENTATION OF COLLAGEN MOLECULES SHOWN IN

PANELS A AND B. REPRINTED WITH PERMISSION FROM KADLER ET AL. (2000) [89]. ... 8

FIGURE 1.6. COLLAGEN STRUCTURE IN NATIVE TISSUES SUCH AS A) CORNEAL

(REPRINTED WITH PERMISSION FROM MEEK (2009) [91]) AND B) (REPRINTED

WITH PERMISSION FROM SAWADKAR ET AL. (2013) [92]) IN COMPARISON WITH C) x

RECONSTITUTED FIBRILS IN VITRO (COURTESY OF E. ISMAIL NORTHEASTERN

UNIVERSITY)...... 9

FIGURE 1.7. DIAGRAM SUMMARIZING THE SYNTHESIS OF THE COLLAGEN OF THE

MATRIX. THE BLACK ARROWS INDICATE THE PROGRESSION OF THE H3-PROLINE AS IT

ENTERS THE CELL, PACKAGED IN SECRETORY VACUOLES IN THE GOLGI ZONE, AND THEN

RELEASED FROM THE CELL. THE NEWLY FORMED COLLAGEN DIFFUSES SOME DISTANCE

FROM THE CELL BEFORE POLYMERIZING WITH STRIATED FIBRILS. REPRINTED WITH

PERMISSION FROM REVEL AND HAY (1963) [141]...... 27

FIGURE 1.8. RESIDUAL BODIES FOUND IN A GUINEA-PIG FIBROCYTE (LEFT) AND A

DEVELOPING MOUSE (RIGHT) CONTAINING COILED BANDED

FIBRILS. FROM TEN CATE (1972) [146]...... 29

FIGURE 1.9. DIAGRAM ILLUSTRATING THE FIBROBLAST PHAGOCYTOSES EXTRACELLULAR

COLLAGEN AND REPLACEMENT OF COLLAGENASE ACTIVITY BY LYSOSOMAL ENZYME

ACTIVITY. 1) PHAGOCYTOSIS OF EXTRACELLULAR COLLAGEN FIBRILS (THE BLACK DOTS

REPRESENT ACTIVITY OF COLLAGENASE). 2) THE COLLAGEN FIBRIL IS IN THE

FIBROBLAST IN A PHAGOSOME. 3) THE COLLAGEN FIBRIL IS NOW WITHIN AN ELECTRON-

DENSE PHAGOLYSOSOME DERIVED FROM FUSION OF LYSOSOMES WITH THE PHAGOSOME.

FROM TEN CATE AND SYRBU (1974)...... 31

FIGURE 1.10. MODE OF COLLAGEN FIBRILLOGENESIS IN EMBRYONIC CHICK AND MOUSE

TENDONS BY TRELSTAD AND HAYASHI (1979). FIBRILS ASSEMBLY OCCUR AT RECESSES

OF THE FIBROBLAST CELL SURFACE. ENDOPLASMIC RETICULUM (ER); GOLGI xi

APPARATUS (GA); CONDENSATION VACUOLES (CV). REPRINTED WITH PERMISSION FROM

TRELSTAD AND HAYASHI (1979) [161]...... 33

FIGURE 1.11. PROPOSED MODEL OF THE COMPARTMENTALIZATION OF THE EXTRACELLULAR

SPACE BY THE CORNEAL FIBROBLAST. REPRINTED WITH PERMISSION FROM BIRK AND

TRELSTAD (1984) [164]...... 35

FIGURE 1.12. PROPOSED MODEL OF THE COMPARTMENTALIZATION OF THE EXTRACELLULAR

SPACE BY THE TENDON FIBROBLAST. REPRINTED WITH PERMISSION FROM BIRK AND

TRELSTAD (1986) [165]...... 36

FIGURE 1.13. PROPOSED MODEL OF THE COMPARTMENTALIZATION OF THE EXTRACELLULAR

SPACE BY THE TENDON FIBROBLAST BASED ON SEM IMAGES. REPRINTED WITH

PERMISSION FROM YANG AND BIRK (1986) [166]...... 36

FIGURE 1.14. FIBRIPOSITOR MODEL PROPOSED BY CANTY ET AL. (2004) [12]. REPRINTED

WITH PERMISSION...... 37

FIGURE 1.15. THE PROCESSES OF COLLAGEN FIBRIL NUCLEATION AND MOVEMENT IN THE

FIBRIPOSITOR MODEL PROPOSED BY KALSON ET AL. (2013) [168]. REPRINTED WITH

PERMISSION...... 38

FIGURE 1.16. FLOW-INDUCED CRYSTALLIZATION MODEL BY PATEN ET AL. (2016) [42].

REPRINTED WITH PERMISSION...... 40

FIGURE 1.17. CHAPMAN MODEL. A) SCHEMATIC REPRESENTATION OF ASSEMBLING

COLLAGEN MOLECULES. B) SCHEMATIC TRANSVERSE SECTIONS THROUGH A FIBRIL xii

ASSEMBLING FROM PN-COLLAGEN MOLECULES. SECTIONS LABELED 5* REPRESENT THE

N-PROPEPTIDE SECTION OF THE MOLECULE WHICH ARE LARGER THAN OTHER SECTIONS

OF THE MOLECULE LABELED 1-5. C) AT CRITICAL DIAMETER, THE GROWTH INHIBITING

DOMAINS FORM A CONTINUES LAYER ON THE SURFACE OF THE FIBRIL, INHIBITING

FURTHER ACCRETION OF MOLECULES. REPRINTED WITH PERMISSION FROM CHAPMAN

(1989) [27]...... 50

FIGURE 1.18. SCHEMATIC OF THE STRUCTURES FORMED FROM THE CO-POLYMERIZATION OF

PN-COLLAGEN III OR PN-COLLAGEN I WITH COLLAGEN I. PN-COLLAGEN III AND

COLLAGEN I CO-POLYMERIZE TO FORM LONG, THIN, AND CYLINDRICAL FIBRILS WITH 67

NM PERIODICITY (B) AS WELL AS THIN AND NON-STRIATED STRUCTURES (A).

COLLAGEN I POLYMERIZES TO GENERATE CYLINDRICAL FIBRILS THAT ARE SHORTER

AND THICKER (C) THAN HYBRID FIBRILS IN (B). PN-COLLAGEN I CO-POLYMERIZES WITH

COLLAGEN I TO FORM RIBBON-LIKE STRUCTURES THAT HAVE A 67 NM PATTERN (D). PN-

COLLAGEN I POLYMERIZES BY ITSELF TO GENERATE LARGE, THIN SHEET-LIKE

STRUCTURES THAT ARE BANDED (E). FROM ROMANIC ET AL. (1992) [257]...... 54

FIGURE 1.19. REGULATION OF COLLAGEN FIBRIL DIAMETER AND GROWTH BY TYPE V

COLLAGEN MOLECULES. TYPE V AMINO-TERMINAL DOMAINS PROJECT ONTO THE FIBRIL

SURFACE THROUGH THE GAP ZONES OF THE FIBRIL, AND WHEN SUFFICIENT NUMBERS

HAVE ACCUMULATED, THEY BLOCK FURTHER ACCRETION OF COLLAGEN MONOMERS

THRU ELECTROSTATIC HINDRANCE OR BY INTERACTIONS WITH OTHER CHARGED

MOLECULES SUCH AS THE LEUCINE-RICH . REPRINTED WITH

PERMISSION FROM BIRK (2001) [269] AND MARCHANT ET AL. (1996) [270]...... 55 xiii

FIGURE 1.20. UNIPOLAR AND BIPOLAR COLLAGEN FIBRILS FROM EMBRYONIC CHICK TENDON.

THE MOLECULAR SWITCH REGION OF A BIPOLAR FIBRIL IS SHOWN IN THE LOWER PANEL.

REPRINTED WITH PERMISSION FROM KADLER ET AL. (2000) [89]...... 58

FIGURE 1.21. GROWTH OF A PARABOLOIDAL TIP. DOTTED CURVE SHOWS THE TIP AFTER A

SHORT INTERVAL OF TIME. THE GROWING TIP REMAINS UNCHANGED IN SHAPE. FROM

HOLMES ET AL. (1992) [282]...... 60

FIGURE 1.22. A MODEL FOR LINEAR AND LATERAL GROWTH OF FIBRIL SEGMENTS DURING

DEVELOPMENT. REPRINTED WITH PERMISSION FROM BIRK ET AL. (1995) [287]...... 61

FIGURE 1.23. POSSIBLE MODELS OF FIBRIL FUSION BASED ON FIBRILS POLARITY. ARROWS

INDICATE MOLECULAR POLARITY WITHIN A FIBRIL AND PINK BOXES INDICATE REGIONS

OF POLARITY REVERSAL. FROM KADLER ET AL. (1996) [76]...... 63

FIGURE 1.24. A MODEL FOR GROWTH OF TENDON FIBRIL SEGMENTS BY BIRK ET AL. (1997)

[288]. REPRINTED WITH PERMISSION...... 64

FIGURE 1.25. MODEL OF FIBRIL GROWTH REGULATION DURING TENDON DEVELOPMENT BY

NURMINSKAYA AND BIRK (1998) [291]. REPRINTED WITH PERMISSION...... 66

FIGURE 1.26. A MULTI-STEP MODEL FOR REGULATION OF FIBRILLOGENESIS BY LUMICAN AND

FIBROMODULIN IN THE MOUSE TENDON. FIBRIL INTERMEDIATES FORM BY MOLECULAR

ACCRETION AND STABILIZED THROUGH THEIR INTERACTIONS WITH LEUCINE-RICH

REPEAT PROTEOGLYCANS. THE CHANGE IN COMPOSITION OF THE MATRIX

PROTEOGLYCANS LEADS TO A MULTI-STEP FUSION GROWTH PROCESS. REPRINTED WITH

PERMISSION FROM EZURA ET AL. (2000) [222]...... 67 xiv

FIGURE 1.27. GROWTH OF FIBRIL ENDS. A) EXTRACTED FIBRIL FROM 13-DAY CHICK

EMBRYONIC TENDON. GROWTH OF FIBRIL END AFTER 30 MIN (B) AND AFTER 2 H (C)

INCUBATION IN COLLAGEN SOLUTION. D) A SCHEMATIC OF NUCLEATION AND

PROPAGATION GROWTH MODEL. FROM HOLMES ET AL. (2010) [294]...... 69

FIGURE 1.28. MODEL OF FIBRIL NUCLEATION AND GROWTH DURING TENDON DEVELOPMENT

BY KALSON ET AL. (2015) [13]...... 70

FIGURE 2.1. AMINO ACID LYSINE. THE FREE AMINE GROUP IN ITS SIDE CHAIN IS A POTENTIAL

BINDING SIDE FOR AMINE REACTIVE FLUOROPHORES...... 84

FIGURE 2.2. COLLAGEN ALPHA-1(I) CHAIN AMINO ACID SEQUENCE (BOVINE). THE TRIPLE-

HELICAL REGION IS HIGHLIGHTED AND LYSINE IS SHOWN WITH LETTER K. THERE ARE

36 LYSINES IN THE TRIPLE-HELICAL REGION AND 1 LYSINE IN EACH TERMINAL

TELOPEPTIDES. SIGNAL PEPTIDE (1-22), N-TERMINAL PROPEPTIDE (23-161), N-

TERMINAL TELOPEPTIDES (162-177), TRIPLE-HELICAL REGION (178-1191), C-

TERMINAL TELOPEPTIDES (1192-1215), C-TERMINAL PROPEPTIDE (1218-1463).

REPRINTED WITH PERMISSION FROM UNIPROT: THE UNIVERSAL PROTEIN

KNOWLEDGEBASE [417]...... 85

FIGURE 2.3. A) ALEXA FLUOR 488 CARBOXYLIC ACID, TFP ESTER (MOLECULAR FORMULA:

C39H44F4N4O11S2, MOLECULAR WEIGHT: 884.91). B) FLUORESCENCE EXCITATION (—

) AND EMISSION (- - -) OF AF488 (EX/EM: 494/519 NM). IMAGE PROVIDED BY SUPPLIER,

THERMO SCIENTIFIC...... 86 xv

FIGURE 2.4. A) REACTION OF ALEXA FLUOR 488 WITH A FREE AMINE GROUP. FROM

WWW.THERMOFISHER.COM AND THE MOLECULAR PROBES HANDBOOK [425]...... 87

FIGURE 2.5. A) ALEXA FLUOR 488 PHOTOSTABILITY IN COMPARISON WITH OREGON GREEN

488 AND FLUORESCEIN. SAMPLES WERE SCANNED 10 TIMES WITH A 25 MW LASER

POWER AND APPROXIMATELY 5 MINUTES EACH TIME. B) RELATIVE FLUORESCENCE OF

ALEXA FLUOR 488 AND FITC AT MULTIPLE MOLE OF DYE PER MOLE OF PROTEIN. FROM

WWW.THERMOFISHER.COM AND THE MOLECULAR PROBES HANDBOOK [425]...... 88

FIGURE 2.6. SCHEMATIC OF COLLAGEN MOLECULE LABELED WITH 2 FLUOROPHORES. THERE

ARE 38 POSITIVELY CHARGED AND HYDROPHILIC LYSINES AND ONE N-TERMINAL Α-

AMINE AS POTENTIAL BINDING SITE ON EACH Α-CHAIN FOR THE LABEL [417].

DIMENSIONS ARE NOT TO SCALE...... 89

FIGURE 2.7. A) BOVINE EYE DISSECTION. B) CLEANING THE SCLERA. C) CLEANED SCLERA

WAS CUT INTO SMALLER PIECES...... 92

FIGURE 2.8. SCHEMATIC OF FIBRIL EXTRACTION PROCESS. SCLERAL TISSUE WAS DISSECTED

FROM 1-10 DAY OLD BOVINE EYEBALLS. TISSUE WAS CUT INTO 3-4 PIECES AND SOME

SHALLOW CUTS WERE MADE ON EACH PIECE. THEN THEY WERE PLACED IN 20 ML OF THE

TRYPSIN EXTRACTION SOLUTION. SAMPLES WERE SHAKEN FOR ~5 MINUTES UNTIL THE

SCLERA SWELLED AND SOME FIBRILS WERE EXTRACTED. PIECES OF SCLERA WERE THEN

REMOVED FROM SOLUTION AND FIBRIL SUSPENSION WAS CENTRIFUGED AT 5000×G FOR

30 MINUTES. THE SUPERNATANT WAS REMOVED, AND THE FIBRIL PELLET WAS

RESUSPENDED AND STORED IN PBS AT 4 °C...... 93 xvi

FIGURE 2.9. SCHEMATIC OF SAMPLE PREPARATION STEPS FOR PROTEOGLYCAN ANALYSIS.

300 ΜL OF FIBRIL SUSPENSION WAS USED FOR EACH SAMPLE. SAMPLES WERE

CENTRIFUGED FOR 30 MIN AT 14,500X G TO ISOLATE THE SUPERNATANT AND PELLET.

SUPERNATANT WAS STORED UNTIL 50 ΜL 8M GUHCL WAS ADDED 24 HOURS PRIOR TO

PROTEOGLYCAN STAINING. THE PELLET WAS RESUSPENDED FOR 24 HOURS IN 300 ΜL

OF 10MM HCL AND THEN 50 ΜL 8M GUHCL WAS ADDED 24 HOURS PRIOR TO

PROTEOGLYCAN STAINING...... 94

FIGURE 2.10. SCHEMATIC OF PROTEOGLYCANS ANALYSIS PROCESS...... 95

FIGURE 2.11. NIKON INVERTED MICROSCOPE (ECLIPSE TE2000-E) EQUIPPED WITH A

COOLSNAP EZ CCD CAMERA AND A HIGH SPEED EMCCD CAMERA...... 96

FIGURE 2.12. DIAMETER MEASUREMENT OF EXTRACTED SCLERA FIBRILS. A) TYPICAL TEM

IMAGE OF TRYPSIN EXTRACTED SCLERA FIBRILS. IMAGES SHOWED NATIVE D-BANDING

PERIODICITY AND UNIFORM DIAMETER ALONG THE LENGTH OF A SINGLE FIBRIL. B) THE

DIC IMAGE SHOWS A TYPICAL FIBRIL (ORIENTED IN NORTHWEST-SOUTHEAST

DIRECTION) FROM THE SAME SCLERA THAT WAS USED FOR TEM IMAGING. C) THE

INTENSITY PROFILE ACROSS THE FIBRIL (ALONG THE BLACK ARROW) SHOWN IN B.

FIBRIL DIAMETERS WITH A MEAN AND STANDARD DEVIATION OF 116.7 ± 38.6 NM WERE

CORRELATED TO THE INTENSITY SHIFT ACROSS FIBRILS WITH MEAN AND STANDARD

DEVIATION OF 2687 ± 524 (ARBITRARY UNITS)...... 98

FIGURE 2.13. ARRHENIUS PLOT. BY MEASURING THE REACTION RATE CONSTANT AT LEAST

AT 2 DIFFERENT TEMPERATURES, THE ACTIVATION ENERGY (EA) AND THE PRE- xvii

EXPONENTIAL FACTOR (A) CAN BE CALCULATED FROM THE EQUATION OF THE LINE.

...... 101

FIGURE 2.14. DOL ACHIEVED AT PH 7.5, 8.0, AND 8.5 BY ADDING 3, 9, AND 15X EXCESS

MOLES OF ALEXA FLUOR 488 TO MOLES OF COLLAGEN (AF488:COLL). ANOVA TEST

WITH A SIGNIFICANCE LEVEL OF 0.05 SHOWED THAT LABELING PERFORMANCE IS NOT

SIGNIFICANTLY DIFFERENT AT PH 7.5, 8.0, AND 8.5 WITHIN EACH INITIAL MOLAR RATIO

OF AF488:COLL GROUPS. HOWEVER, LABELING PERFORMANCE WAS SIGNIFICANTLY

DIFFERENT BETWEEN INITIAL MOLAR RATIO OF AF488:COLL GROUPS. DATA ARE

EXPRESSED AS MEAN ± STANDARD DEVIATION (N=3 REPLICATES)...... 103

FIGURE 2.15. TURBIDITY OF 50 µG/ML LABELED COLLAGEN AT 37 °C. IN THIS EXPERIMENT,

100% OF THE COLLAGEN MOLECULES WERE LABELED. EACH LABELED MOLECULE HAD

AN AVERAGE DOL OF 0, 2, 4 OR 9. LAG AND PLATEAU TIMES WERE CALCULATED AS THE

TIME REQUIRED TO REACH 10% AND 90% OF THE MAXIMUM TURBIDITY, RESPECTIVELY.

DATA ARE PRESENTED AS MEAN ± STANDARD DEVIATION, N=3 REPLICATES PER GROUP.

...... 105

FIGURE 2.16. COLLAGEN FIBRILS FORMED IN NEUTRALIZED SOLUTIONS OF 50 µG/ML

COLLAGEN AT 37°C. A) DIC AND C), E), AND D) FLUORESCENT IMAGES OF FIBRILS

FORMED BY MONOMERS WITH DOL OF 0, 2, 4, AND 9 RESPECTIVELY. WE USED DIC

IMAGING FOR THE UNLABELED FIBRILS (DOL =0) BECAUSE THERE WAS NO

FLUORESCENCE SIGNAL. B), D), F), AND H) TEM IMAGES OF FIBRILS FORMED BY

MONOMERS WITH DOL OF 0, 2, 4, AND 9 RESPECTIVELY...... 107 xviii

FIGURE 2.17. SMO MICROSCOPY SHOWING ORIENTATION OF TYPE I BOVINE COLLAGEN

MOLECULES ASSOCIATED WITH FIBRILS. WHEN COLLAGEN IS LABELED WITH TWO

FLUOROPHORES SEPARATED BY A LARGE ENOUGH DISTANCE (A), IT IS POSSIBLE TO FIT

AN ELLIPSE TO THE PIXEL REPRESENTATION (B). THUS, WE CAN DETERMINE IF THE

SINGLE MOLECULES ARE CO-ALIGNED WITH THE FIBRIL AXIS OR JUST ASSOCIATING (C,

D, AND E). COURTESY OF DR. MONICA SUSILO...... 111

FIGURE 2.18. SYNTHETIC IMAGES OF LABELED COLLAGEN MONOMER WITH THE MAJOR AND

MINOR AXES PLOTTED. THE COLOUR IS REPRESENTATIVE OF THE NUMBER OF PHOTONS

PER PIXEL. A) SYNTHETIC IMAGE OF DOUBLY-LABELED MONOMER FOR 25 PHOTONS PER

FLUOROPHORE AT A SPACING OF 300 NM. B) SYNTHETIC IMAGE OF SINGLY-LABELED

MONOMER FOR 25 PHOTONS PER FLUOROPHORE FOR COMPARISON. C) SYNTHETIC

IMAGE OF DOUBLY-LABELED MONOMER FOR 2500 PHOTONS PER FLUOROPHORE AT A

SPACING OF 68 NM. COURTESY OF DR. CHARLES A. DIMARZIO...... 113

FIGURE 2.19. ROC CURVES FOR A) 68 NM AND B) 300 NM SPACINGS. THE CURVES IN EACH

CASE WERE COMPUTED WITH, FROM TOP TO BOTTOM 2500, 500, 250, 100, 50, AND 25

PHOTONS PER FLUOROPHORE. FOR THE 300 NM SPACING, THE DETECTION STATISTICS

ARE NEARLY PERFECT AT ALL NUMBERS OF PHOTONS. COURTESY OF DR. CHARLES A.

DIMARZIO...... 114

FIGURE 2.20. QUANTIFICATION OF PROTEOGLYCANS WITH ALCIAN BLUE (N=6). THE

CONCENTRATION OF PROTEOGLYCANS WAS MEASURED USING THE STANDARD

CONCENTRATION OF . “PELLET” REFERS TO THE SAMPLE

CONTAINING SEPARATED FIBRILS AFTER EXTRACTION AND “SUPERNATANT” REFERS TO xix

THE SAMPLE CONTAINING THE FIBRIL EXTRACTION SUSPENSION WITHOUT THE FIBRILS.

NOTE THAT SOME ERROR BARS ARE TOO SMALL TO BE SEEN. COURTESY OF ALEXANDRA

A. SILVERMAN...... 115

FIGURE 2.21. INCORPORATION OF LABELED COLLAGEN INTO FIBRILS. SINGLE, NATIVE

SCLERA COLLAGEN FIBRILS INCORPORATE EXOGENOUS LABELED MONOMER WITH

KINETICS SIMILAR TO FIBRIL ASSEMBLY. (A), (B), AND (C) FLUORESCENT IMAGES SHOW

INCORPORATION OF LABELED MONOMERS ON A TYPICAL FIBRIL OVER TIME AT 25 °C.

(D) THE GRAPH SHOWS TYPICAL ACCUMULATION OF LABELED MONOMERS INTO SCLERA

FIBRILS AT 25 AND 30 °C. LAG AND PLATEAU TIMES IN MINUTES (THE TIMES WHEN THE

FIBRIL INTENSITY REACHED 10% AND 90% OF ITS MAXIMUM INTENSITY) WERE

MEASURED AS 59.9 ± 19.4 AND 111.8 ± 37.4 AT 25 °C (N=21 REPLICATES) AND 43.5 ±

11.7 AND 97.8 ± 20.7 AT 30 °C (N=23 REPLICATES), RESPECTIVELY...... 116

FIGURE 2.22. PHOTOBLEACHING OF FIBRILS AFTER REACHING EQUILIBRIUM WITH 2 µG/ML

LABELED MONOMERS. FIBRILS WERE IMAGED CONTINUALLY WITH 1 SECOND EXPOSURE

TIME...... 117

FIGURE 3.1. SMALL-ANGLE X-RAY DIFFRACTION FROM CORNEAL COLLAGEN FIBRILS. FROM

BOOTE ET AL. (2003) [455]...... 125

FIGURE 3.2. ELECTRON MICROGRAPH OF TYPE I COLLAGEN FIBRILS IN BOTH TRANSVERSE

AND LONGITUDINAL ORIENTATIONS. SCALE BAR IS 300 NM. REPRINTED WITH

PERMISSION FROM CASSELLA ET AL. [469]...... 126 xx

FIGURE 3.3. HIGH-RESOLUTION SEM VIEW OF COLLAGEN FIBRILS FROM THE RAT TAIL

TENDON. REPRINTED FROM OHTANI (1992) [479]...... 128

FIGURE 3.4. AFM IMAGES OF RECONSTITUTED COLLAGEN FIBRILS. REPRINTED WITH

PERMISSION FROM REVENKO ET AL. (1994) [482]...... 129

FIGURE 3.5. CORRELATIVE SHG-TEM IMAGING. A) SHG AND B) TEM IMAGES OF

UNSTAINED COLLAGEN FIBRILS (SCALE BAR, 10 MM). C AND D) ZOOMED REGIONS OF

INTEREST FROM A AND B (SCALE BAR, 5 MM). E) SHG PHOTON NUMBER AS A FUNCTION

OF THE FIBRIL DIAMETER MEASURED ON THE TEM IMAGE. F) ALL SAMPLES USING

DIAMETER BINS OF 50 NM. REPRINTED WITH PERMISSION FROM BANCELIN ET AL. (2014)

[500]...... 130

FIGURE 3.6. SCHEMATIC DIAGRAM OF THE DIC OPTICAL PATH SHOWING THE OPTICAL

COMPONENTS (LEFT) AND THEIR EFFECTS ON THE ORTHOGONALLY-POLARISED LIGHT

(RIGHT). FROM COGSWELL ET AL. (1997) [502]...... 132

FIGURE 3.7. BEHAVIOUR OF A) THE SIGNAL STRENGTH, B) THE CONTRAST FROM A PHASE

CHANGE, C) THE LINEARITY OF THE RESPONSE, AND D) THE SIGNAL-TO-NOISE RATIO OF

A DIC SYSTEM FOR DIFFERENT VALUES OF BIAS RETARDATION. REPRINTED WITH

PERMISSION FROM COGSWELL AND SHEPPARD (1992) [503]...... 133

FIGURE 3.8. DIC-EIS FOR SIMULATED DIC IMAGES WITH DIFFERENT DIAMETER FIBRILS (FOR

A 60X OBJECTIVE AND 1.45 NA). NOTE THE LINEAR REGION BETWEEN 60 NM AND 240

NM DIAMETER FIBRILS. FROM BRENDAN FLYNN DISSERTATION, NORTHEASTERN

UNIVERSITY (2012) [508]...... 134 xxi

FIGURE 3.9. THE EXPERIMENTAL SETUP WHERE COLLAGEN FIBRILS WERE AIR-DRIED ON A

REFERENCED TEM GRID AND IMAGED WITH DIC MICROSCOPY. THE COLLAGEN FIBRIL

IS NOT IN SCALE...... 135

FIGURE 3.10. SEM IMAGE OF 25 µM WIDE AND 20 µM DEEP TRENCHES ON 100 µM THICK

PDMS SHEET. SCALE BAR IS 20 µM. COURTESY OF DR. POOYAN TIRANDAZI...... 137

FIGURE 3.11. A) THE MICROSCOPE STAGE EQUIPPED WITH MICROMANIPULATORS. B)

SCHEMATIC OF PULLING COLLAGEN FIBRILS OUT OF FIBRIL SUSPENSION USING

MICRONEEDLES AND PLACEMENT OF FIBRILS OVER TRENCHES OF PDMS...... 138

FIGURE 3.12. A) DIC IMAGE OF A REPRESENTATIVE FIBRIL AT 40X MAGNIFICATION. THE

FIBRIL WAS ORIENTED IN NORTHWEST-SOUTHEAST DIRECTION FOR DIC IMAGING.

HOWEVER, THE IMAGE IS ROTATED HERE FOR COMPARISON WITH TEM IMAGE. B) TEM

IMAGE OF THE SAME FIBRIL. C) HIGH MAGNIFICATION OF A SELECTED REGION OF THE

FIBRIL WITH A RAPID DIAMETER CHANGE. D) CORRELATION OF DIC-EIS AND DIAMETER

ALONG THE SELECTED REGION. THE X AXIS REPRESENTS POSITION ALONG THE FIBRIL

STARTING FROM THE LEFT SIDE OF THE FIBRIL...... 141

FIGURE 3.13. SHEARING DIRECTION IN NIKON ECLIPSE TE2000 DIC INVERTED

MICROSCOPE. FROM NIKON’S INSTRUCTIONS MANUAL...... 142

FIGURE 3.14. CHANGE OF DIC-EIS AS A FUNCTION OF FIBRIL DIRECTION (N = 35 FIBRILS).

ASTERISKS SHOW STATISTICAL DIFFERENCE (P < 0.05)...... 143 xxii

FIGURE 3.15. DIC IMAGE OF A FIBRIL OVER THE 25 ΜM WIDE GAP. THE MIDDLE SECTION OF

THE FIBRIL WAS USED TO CALCULATE DIC-EIS ACROSS THE FIBRIL. THE SCALE BAR IS

10 µM...... 144

FIGURE 3.16. SEM IMAGE OF A FIBRIL OVER THE 25 ΜM WIDE GAP. THE MIDDLE SECTION OF

THE FIBRIL WHICH WAS USED TO MEASURE FIBRIL DIAMETER IS SHOWN IN HIGH

MAGNIFICATION...... 145

FIGURE 3.17. DIC-EIS AS A FUNCTION OF FIBRIL DIAMETER (N = 30 FIBRILS). A LINEAR

REGION CAN BE SEEN FOR FIBRILS WITH DIAMETER BETWEEN ~100 NM TO ~300 NM (N =

23 FIBRILS). NOTE THAT DIAMETER VALUES ARE MEASUREMENTS OF DEHYDRATED

FIBRILS WITH 5 NM PLATINUM COATING...... 146

FIGURE 3.18. SEM IMAGE OF A BROKEN FIBRIL DUE TO HIGH ENERGY OF FOCUSED BEAM OF

ELECTRONS. THE BROKEN FIBRIL FORMED A SPRING-LIKE STRUCTURE. THE SCALE BAR

IS 1 µM...... 146

FIGURE 3.19. CORRELATION OF DIC-EIS FOR WET AND DRY FIBRILS. DIC-EIS OF FIBRILS IN

PBS WERE ALSO LINEARLY CORRELATED TO FIBRIL DIAMETERS. Y-AXIS ERROR BARS

WERE SMALLER THAN 53 AU LEADING TO LESS THAN ±2 NM UNCERTAINTY IN

PREDICTED DIAMETER (TOO SMALL TO SHOW ON THE FIGURE). AN ASTERISK INDICATES

A SIGNIFICANT DIFFERENCE, P < 0.05, BETWEEN CONDITIONS...... 147

FIGURE 3.20. DIC IMAGES OF A TYPICAL FIBRIL IN A) DRY AND B) WET CONDITIONS ON

GLASS. C) DIC-EIS OF THE FIBRIL SHOWN IN A AND B...... 148

FIGURE 3.21. REVERSIBLE DIC-EIS OF FIBRILS AFTER AIR DRYING ON GLASS...... 149 xxiii

FIGURE 3.22. MERGED FIBRILS IN A) DIC, B) SEM, AND C) HIGHER MAGNIFICATION OF

REGION SHOWN B. SCALE BARS ARE 10 µM IN A AND B AND 2 µM IN C...... 152

FIGURE 4.1. (A) A DIRECT INTERMOLECULAR HYDROGEN BOND. (B) A BRIDGING WATER

MOLECULE BETWEEN POLAR GROUPS OF TWO TROPOCOLLAGENS. (C) WATER

MOLECULE TRAPPED BETWEEN TWO HYDROPHOBIC SIDE CHAINS OF NEIGHBOURING

TROPOCOLLAGENS. REPRINTED WITH PERMISSION FROM STREETER AND LEEUW (2011)

[203]...... 156

FIGURE 4.2. P-97 SUTTER INSTRUMENT MICROPIPETTE PULLER. FROM WWW.SUTTER.COM.

...... 158

FIGURE 4.3. A REPRESENTATIVE MICRONEEDLE THAT WAS USED TO STRETCH FIBRILS. .. 158

FIGURE 4.4. A FIBRIL THAT WAS DRAWN FROM THE FIBRIL SUSPENSION AND WRAPPED

AROUND MICRONEEDLES IN AIR AND HOLD SLIGHTLY TIGHT TO ADHERE TO THE GLASS.

...... 159

FIGURE 4.5. DIC IMAGE OF A FIBRIL OVER TRENCHES OF PDMS. AFTER THE FIBRIL WAS

WRAPPED AROUND MICRONEEDLES, IT WAS MOVED OVER TRENCHES OF PDMS TO

MEASURE DIC-EIS AND ESTIMATE ITS DIAMETER...... 160

FIGURE 4.6. FIBRILS SECURED BETWEEN NEEDLES INSIDE PBS. THE FIBRIL IN THE PANEL A

IS TIGHT, BUT NOT LOADED (0% STRAIN). THE FIBRIL IN THE PANEL B IS STRETCHED TO

6% STRAIN. NOTE THE NEEDLES DEFLECTION IN THE PANEL B. ALSO, THE COILED

SECTION OF THE FIBRIL CAN BE SEEN ON THE TOP NEEDLE IN THE PANEL A...... 162 xxiv

FIGURE 4.7. ALEXA FLUOR 488 PHOTOBLEACHING. THE DATA IS EXTRAPOLATED FROM THE

MANUFACTURER WEBSITE...... 164

FIGURE 4.8. NORMALIZED FIBRIL INTENSITY. THE FIBRILS’ INTENSITY RAPIDLY INCREASED

AND REACHED A PLATEAU DURING THE FIRST 30 MINUTES. FIBRILS WHICH WERE UNDER

4% AND 6% STRAIN REACHED THEIR MAXIMUM INTENSITY SIGNIFICANTLY FASTER

THAN UNLOADED FIBRILS (T-TEST, P < 0.05)...... 166

FIGURE 4.9. DIC AND FLUORESCENT IMAGES OF A REPRESENTATIVE FIBRIL STRETCHED TO

6% STRAIN BETWEEN NEEDLES. PANEL A (DIC) AND B (FLUORESCENT) SHOW THE

FIBRIL AT TIME 0, RIGHT BEFORE ADDING THE LABELED MONOMERS. PANEL C (DIC)

AND D (FLUORESCENT) SHOW THE FIBRIL AT 30 MINUTES AFTER ADDING THE LABELED

MONOMERS. NOTE THE PUNCTUATED BRIGHT SPOTS ON THE FIBRIL AFTER 30 MINUTES

IN PANEL D...... 167

FIGURE 4.10. EXPONENTIAL FITTED CURVES ON THE NORMALIZED FIBRIL INTENSITY DATA.

THE DASH LINES REPRESENT THE 95% PREDICTION INTERVALS...... 168

FIGURE 4.11. FORMATION OF MICROFIBRILS AROUND THE TESTED FIBRILS BETWEEN

MICRONEEDLES...... 169

FIGURE 4.12. LABELED MONOMERS INCORPORATION ONTO FIBRILS. WHILE ALL FIBRILS

STARTED WITH THEIR HIGHEST MONOMER INCORPORATION, THE NORMALIZED

INCORPORATION ONTO 6% STRAINED FIBRILS REMAINED SIGNIFICANTLY HIGHER THAN

0% AND 4% STRAINED FIBRILS...... 170 xxv

FIGURE 4.13. EXPONENTIAL FITTED CURVES ON THE NORMALIZED LABELED MONOMER

INCORPORATION DATA. THE DASH LINES REPRESENT THE 95% PREDICTION INTERVALS.

...... 171

xxvi

LIST OF TABLES

TABLE PAGE

TABLE 2.1. ABSORBANCE, LAG TIME, AND PLATEAU TIME OF SIGMOIDAL TURBIDITY CURVE

OF 50 µG/ML LABELED COLLAGEN AT 37 °C. IN THIS EXPERIMENT, 100% OF THE

COLLAGEN MOLECULES WERE LABELED. LAG AND PLATEAU TIMES WERE CALCULATED

AS THE TIME REQUIRED TO REACH 10% AND 90% OF THE MAXIMUM TURBIDITY,

RESPECTIVELY. DATA ARE PRESENTED AS MEAN ± STANDARD DEVIATION, N=3

REPLICATES PER GROUP. THE HIGHLIGHTED ROW SHOWS LAG TIME, PLATEAU TIME, AND

TOTAL ABSORBANCE CHANGE OF UNLABELED MONOMERS WHICH WERE SIGNIFICANTLY

DIFFERENT (TWO-TAILED T-TEST, Α=0.05) FROM LABELED MONOMERS...... 106

TABLE 2.2. FIBRILLOGENESIS OF LABELED (DOL≈2) AND UNLABELED MONOMERS MIXTURE.

CHANGES IN KINETICS OF FIBRILLOGENESIS IS NEGLIGIBLE (LESS THAN 10% ERROR) IN

MIXTURES CONTAINING 5% OR LESS LABELED MONOMERS (SHOWN IN HIGHLIGHTED

ROWS). DATA ARE PRESENTED AS MEAN ± STANDARD DEVIATION (N=3 REPLICATES PER

GROUP)...... 109

TABLE 4.1. NORMALIZED INCORPORATION VALUES OBTAINED FROM THE FITTED CURVES AT

5, 10, AND 30 MINUTES...... 172

1

1 INTRODUCTION

Collagen related disorders in several tissues like tendon, ligament, skin, lung, eye, and

are of the main complaints for which patients seek medical care [1-8]. However, the current

therapy methods are mostly ineffective, expensive, and cannot entirely restore the tissue to

its healthy condition. Our understanding of the underlying mechanism of collagen fibril

formation, growth, and remodeling can help as a basis for study of collagen related

disorders which are caused by the imbalance between synthesis, accumulation, and

degradation of collagen as in [9], [10], and tendinopathy [11].

Understanding how an organized collagenous structure can arise and grow from cells

remains one of the most important basic science questions associated with connective

tissue research. There are currently no widely accepted mechanistic models of formation,

growth, and remodeling of collagenous connective tissue. It has been generally accepted

that collagen is produced and organized within cell surface invaginations [12] meaning that

cells directly produce and place collagen fibrils into the complex

(ECM). However, this is unlikely, particularly in the later stages of development when cells 2

lose their ability to directly access the damaged or developing fibrils in the dense and

mature ECM [13, 14]. Further fibril growth has been proposed to be the result of molecular

accretion and/or linear and lateral interfibrillar fusion [15-17] and is regulated by several

intrinsic and extrinsic factors [18] such as other collagen types [19, 20], proteoglycans [21,

22], enzymes activities [23, 24], fibrils structure [25-28].

On the other hand, the role of local tissue micro-mechanical environment, namely stresses and strains acting on collagen fibrils, has been underappreciated in developing these models. It has been shown that placement of collagen is a function of the local tissue micro- mechanical environment [29-32], that fibroblast traction can rearrange collagen fibrils [33-

38], and in some cases, that actin filaments align with fibripositors and intracellular fibrils

[39]. However, still the detailed mechanisms by which the actual collagen molecules

(and/or microfibrils) are added (or subtracted) from the resident collagen fibrils are not known. It is likely, however, that the mechanism will operate independently of the cell surface or at some nominal distance from it [40], guided by spatial cues [41] and mechanical forces [42].

A growth and remodeling theory [43] has been formulated in Dr. Ruberti’s research group that suggests collagen monomers and enzymes are in a dynamic equilibrium with existing fibrils. Tensile forces on fibrils can shift this equilibrium and change the balance between molecular association (kon) and dissociation (koff). Indeed, it has been shown in our

laboratory that 1) tension can directly drive initial fibrillogenesis and structures into the path of force [42], 2) applied mechanical strain preferentially preserves type I collagen against enzymatic degradation in molecule [44], fibril [45, 46], and tissue [47-49] levels, 3

3) that in the absence of cells, unloading of fibers can lead to disassociation of collagen

fibers [42], 4) and cyclic stress leads to strengthening of the collagenous structure [50]. All

together suggest that application of mechanical forces can induce fibril formation, strengthen fibrils, and decrease molecular dissociation rate. The missing piece, for elucidation of load-bearing collagenous tissue evolution, is the role that mechanics can play in the mechanism of collagen fibril’s growth. In this PhD dissertation, we seek to answer this question: Does fibril strain promote the molecular assembly of collagen?

Understanding the of collagen provides new opportunities to develop

treatments for variety of pathological conditions by promoting growth or degradation

when/where needed.

1.1 Collagen

Animal tissues have developed throughout evolution to produce a variety of shapes and

functions in biological systems [51]. Connective tissues, the load-bearing component of these systems, have two co-operating elements: fibrillar element to resist tensile forces and interfibrillar gel to resists compressive forces [52, 53]. Collagen is the predominant fibrillar component and the most abundant protein in vertebrate animals [54]. At least 28 different collagen types have been found in tissues that regularly experience tension, compression, and shear forces [55]. The fibril forming types (I, II, III, V, XI, XXIV, and XXVII) are the principal load bearing molecules, with type I collagen being the most ubiquitous (e.g. the primary protein in blood vessel, bone, tendon, ligament, skin, sclera and cornea) [56]. 4

Figure 1.1. Biosynthesis of collagen including hydroxylation, glycosylation, and disulfide-bond formation of the procollagen in the endoplasmic reticulum. Reprinted with permission from Chen and Raghunath (2009) [57].

The biosynthesis of is a complex multistep process, requiring well-controlled intracellular and extracellular events (Figure 1.1). Collagen genes are transcribed from

DNA into mRNA within the nucleus and post-translationally modified in the rough 5

endoplasmic reticulum. Procollagen molecules (the initial form of the molecule) [58, 59]

are secreted through Golgi vacuoles into the ECM. At this stage, the procollagen molecule

is comprised of three α-chains assembled into a triple-helix with a propeptide domain on either end of the molecule [60]. Each α-chain is comprising of a repeating Gly-X-Y amino

acids, in which X and Y are usually proline and hydroxyproline, respectively [61, 62]. Each

α-chain forms a left-handed helix itself which then assembles into a right-handed triple- helix such that the center of the triple-helix is occupied only by glycyl residues (Figure 1.2)

[63]. Any irregularities in this specific amino acid sequence would disturb the triple-helical

conformation.

Figure 1.2. Schematic representation of a cross-section through a collagen triple helix. Reprinted with permission from van der Rest and Garrone (1991) [63]. 6

The C-propeptide works as a nucleation site and initiates the folding of molecule [64, 65].

Later, the propeptide chains are enzymatically cleaved [66] and the remaining molecule is

called tropocollagen [67] or simply collagen. This final collagen molecule is approximately

300 nm long with a diameter of 1.5 nm and a molecular weight of 300 kDa [68]. The middle

triple-helical region is capped by an amino and carboxyl telopeptide extension [69]. The

structure of a type I procollagen molecule is shown in Figure 1.3.

Figure 1.3. Structure of a type I procollagen molecule. Reprinted with permission from Holmes et al. (2018) [18].

In an adult tissue, collagen molecules are assembled into fibrils in a quarter-staggered arrangement. This specific arrangement causes an axial periodicity of about 67 nm which is called D-banding (Figure 1.4) consisting of an asymmetric pattern labeled a-e (Figure 7

1.5) that is attributed to the groups of hydrophilic or hydrophobic amino acids in adjacent

collagen molecules [70]. Each period has a regularly repeated gap (0.54D) and overlap

(0.46D) region that can be detected by electron microscopy [71, 72] or x-ray diffraction

[73, 74]. The unusual amino acid composition of collagen molecules and the way these asymmetric triple-helixes arrange into fibrils are the hallmark of collagen that also give

rise to diverse patterns such as bundles, weaves, and layers [75, 76].

Figure 1.4. Schematic of hierarchical collagen structure and organization. Reprinted with permission from Canelon and Wallace (2016) [77].

1.2 Self-Assembly

Tissue-extracted collagen molecules in solution polymerize spontaneously in physiological

pH, temperature, and ionic strength [78-81]. Early electron microscopy studies showed that these reconstituted fibrils have the same detailed fine structure of native fibrils [71, 81-83] 8

and slight deviation from physiological conditions leads to formation of abnormal fibrils

[84].

Turbidity measurements of collagen assembly revealed a sigmoidal curve with two different stages for heat-dependent polymerization of collagen molecules in vitro: I) a lag

(nucleation) phase with no detectable turbidity change, and II) a growth phase with linear increase in turbidity. Monomers assemble into intermediate forms smaller than fibrils

(microfibrils) [85] with a prompt decrease in diffusion coefficient [86] during the lag phase and then rapidly form larger aggregates by lateral and longitudinal assembly into fibrils

[78, 80, 87, 88].

Figure 1.5. Asymmetric pattern of type I collagen fibril showing D-banding and sub-

bands labeled c1, b2, b1, a4-a1, e2, e1, c3, and c2 from left to right as shown in panel B. A) Negative and B) positive staining pattern of a type I collagen fibril. C) Schematic of the orientation of collagen molecules shown in panels A and B. Reprinted with permission from Kadler et al. (2000) [89].

Thermodynamically, type I collagen fibrillogenesis in vitro is an entropy-driven self-

assembly process [90] which is driven by the loss of solvent molecules from the collagen

surface. In a spontaneous process, Gibbs free energy change ( ) which is defined as

∆𝐺𝐺 9

= (1.1)

∆𝐺𝐺 ∆𝐻𝐻 − 𝑇𝑇∆𝑆𝑆 is negative. In the equation, is enthalpy change, is entropy change, and is

temperature. Since the process∆ 𝐻𝐻is spontaneous only at ∆high𝑆𝑆 temperatures, the collagen𝑇𝑇

fibrillogenesis is an endothermic reaction (positive ).

∆𝐻𝐻

Figure 1.6. Collagen structure in native tissues such as A) corneal stroma (reprinted with permission from Meek (2009) [91]) and B) tendon (reprinted with permission from Sawadkar et al. (2013) [92]) in comparison with C) reconstituted fibrils in vitro (courtesy of E. Ismail Northeastern University).

Self-assembly however cannot entirely explain the formation of highly organized native

collagenous tissues. Such as, collagen fibrils in cornea are narrow, uniform in diameter and

precisely organised parallel to one another in each lamella but varying angles with those in

neighbouring lamellae (Figure 1.6.A) [93]. This arrangement is vital to maintain transparency of cornea [94, 95]. Alternatively, collagen in tendon forms a multi- hierarchical structure of molecules, microfibrils, fibrils, fibers, and fascicles. All bundles are parallel to the long axis of the tendon to transmit forces and withstand tension (Figure

1.6.B) [96]. However, reconstitution of fibrils in vitro from soluble tissue-extracted 10

collagen through self-assembly leads to formation of an unorganized network of fibrils

with varying diameter [87, 97] (Figure 1.6.C). In this regard, the role of cells in providing

the suitable environment and interactions with non-collagen molecules, seem to be critical.

1.3 Fibrillogenesis, Growth, and Remodelling

Although connective tissue formation has been the subject of investigation for more than

100 years, the cellular and molecular mechanisms that drive initial tissue formation and

growth have yet to be clearly identified [98]. Part of the difficulty is the highly dynamic nature of fibrillogenesis and growth of fibrils in the complex environment of intracellular

and extracellular matrix. It has been challenging to precisely separate the events that cause

conversion of soluble collagen to an insoluble fibril. This section summarizes the main

studies that attempt to reveal the underlying mechanism of collagen fibril formation and

growth.

1.3.1 The Site and Mechanism of Initial Fibril Formation

The literature contains contradictory explanations of the origin and mechanism of

formation of connective tissue fibrils (as named by Mall [99]) or collagenous fibrils (as named by Bell [100]). The question whether the collagen fibrils of the connective tissues arise within the cytoplasm of collagen-secreting cells, on the surface of the cell, or in the intercellular spaces has been the subject of studies at least since Schwann (1839) [101,

102]. One group proposed an intracellular origin of fibrils where fibrils are formed within 11

the cell by fusion of intracellular granules; another group supported an epicellular origin

of fibrils and suggested that fibrils arise at the surface of the cell. In both of these models,

fibrils are excreted into extracellular space through a direct transformation of the cytoplasm of the cell. A third group proposed an intercellular origin of fibrils where fibrils are formed

outside of and at some distance from the cells through accumulation of intercellular

substances secreted by cells. In this section, some of the prominent studies are revisited to

elucidate the source of this never ending contradiction.

In 1902, Mall [99] described stained serial sections of a pig’s skin. He found irregular

shaped cells with processes from which fine threads of protoplasm were radiated. Since the

cell processes contained wavy fibrils, he concluded that the fibrils form in the protoplasm

at the periphery of the cell and then are gradually thrown off of the cell.

In 1903, Mallory [103] described a fibrillar substance named “fibroglia” produced by

connective tissue cells. The fibroglia fibrils were chemically and morphologically different

from ordinary intercellular fibrils. They were stained red with the same method that stained

the connective tissue fibrils blue. The fibroglia fibrils were found in intimate relation to the

protoplasm of the actively growing connective tissue cells, both of inflammatory and of

tumour origin and were hard to find when the cells were in an inactive state. Mallory

concluded that the blue-staining fibrils are intercellular in origin and not a cast-off product

of the cytoplasm.

Hertzler (1910) [104] studied the formation of fibrous tissue in wound-healing of the

peritoneum. He reported that fibril formation started before the advent of cells. Early fibrils

were characterized as fibrin bundles, but were gradually changed into adult connective 12

tissue fibrils. The early fibril formation was before the advent of cells and the appearance of the adult connective tissue fibrils took place without immediate contact with cells.

Therefore, it was suggested that the initial fibril formation processes are chemical like those of blood coagulation, and that cells are playing an entirely secondary role.

Ferguson (1912) [105] studied the living connective tissue cells in the fins of fish embryos.

He described three stages in the histogenesis of connective tissue which were characterized predominantly by round, stellate, and spindle shape cells, respectively. Fine fibrils were found after appearance of cellular processes coincidently with the stellate cells. The stellate cells underwent rapid morphological transformation and corresponded to a more transient duration than either a preceding or succeeding stage. The spindle cells took their position alongside connective tissue fibers or bundles, elongated along them, and then became indistinguishable from the larger bodies. Frequently, these spindle cells thickened to a normal spindle cell and sometimes even threw out other processes. The stellate cells were observed more to lie in relation with the finer fibrils, and the spindle cells with the coarser fibers or fiber bundles. Since fibrils were found both at the surface and within the cells, it was proposed that fibrils are originally embedded in a “plastic cytoplasm” of the cells.

Baitsell (1915) [106] studied living cultures of adult frog tissues from which he concluded that the white fibers of connective tissue are not formed by an intracellular action, but arise directly by a transformation of the fibrin network. While transformation of fibrin is an outdated theory, more interestingly, he showed that exerting tension with needles or movements of living isolated spindle cells hasten the formation of the fibers. He also quoted from Adami, J. G. in a work by Leo Loeb that “When a drop of uncoagulated lymph 13

is placed between two glass slides, the mere act of pulling one slide over the other leads to

the appearance of fibrils, which grow in length and bulk; which, like those of connective

tissue, are not only intracellular, but actually traverse cell bodies situated in their path;

which show themselves first in immediate connection with these cells, the cells, as we now

hold, liberating an enzyme that determines the modification of the more soluble protein into a precipitated or coagulated modification. But the lines of the precipitation are evidently along the lines of strain”. In a following study [107] on prepared sections of skin

wounds in frogs he showed that the formation of new fibrous tissue takes place before the

invasion of cells into the coagulating tissue and therefore cannot be due to an intracellular action.

Baitsell continued his research on the origin of connective tissue through study of both the prepared and living material of the connective tissues in the amphibian [108] and chick

[109] embryos. In both studies, he showed that the forerunner of the connective tissues is a transparent, gelatinous, and cell-free ground-substance. It may be worthwhile to quote a couple of paragraphs from him: “From the morphological standpoint the results of the present study indicate that the formation of connective tissue in the amphibian embryo is similar to the process which takes place in transformation of the plasma clot. The intercellular ground substance of developing connective tissue may therefore be compared in its morphology to the plasma clot. This ground substance when first formed appears homogeneous or with a fine fibrillation. The process of transformation into a fibrous tissue is a progressive one. The fibrillation increases, bundles of fibres are formed, and in time the entire ground-substance, which at first showed such a high degree of homogeneity, 14

becomes transformed into a fibrous tissue. It is indicated that this transformation occurs as

the results of the introduction of mechanical factors in the embryo. These factors may be

due to certain lines of tension in the embryo corresponding to the inherent polarity of the organism or, just as in the plasma clot, the movements of the cells through the ground substance may introduce mechanical factors which aid in the transformation of the ground- substance into a fibrous tissue. The cells, however, are to be regarded primarily as assimilative and secretory agents, chiefly concerned in the formation of the undifferentiated ground-substance” [108]. His results were so convincing that he wrote: “I am of the opinion that considering the results which are now available the question of the origin of connective tissue must be regarded as settled in favour of the intercellular theory.”

[109].

Isaacs (1916) [110] in a short report mentioned that the connective tissue fibrils, that were previously described as “exoplasmic fibrillae”, do not appear in living intercellular connective tissue. Instead, he stated that the fibrils can be produced by any action that cause shrinkage of the intercellular substance. Furthermore, he reported a progressive growth of fibrils (stronger and denser) by increase of substance deposition. He proposed that the movement of cells probably effect the distribution of the material and cause the fibrillar structures of the adult fibers. He stated that “forces acting on the jelly-like embryonic connective tissue affect the cells and colloid alike, but the cells have the power to adjust themselves to the new condition, thus producing permanent structures in reaction to pulls and pushes”. 15

In his following paper [14], he studied the effect of various fixation and staining procedures

on the structure and mechanism of developing connective tissue of chicks, pigs, and

humans. For comparison, living tadpoles and embryos of chicks, pigs and adult frogs were

used for the study of fresh tissue. He showed that the intercellular connective-tissue spaces

are filled with a clear, homogeneous jelly-like substance and fibrils are formed by the

dehydrating or coagulating action of fixatives from the homogeneous jelly. He showed that

in some cases, fibrils could not be demonstrated in the fresh intercellular connective tissue

space, until dehydration of tissue led to fibril formation. Furthermore. he showed that as

the tissue grows older, it becomes denser and cells have less freedom in the thicker

intercellular space to influence the direction of fibers. These observations supported his view of intercellular origin of fibrils and showed the possibility of formation of artificial fibrils due to fixation or dehydration in prepared samples.

Lewis (1917) [111] described the connective tissue fibril formation based on observation of living cells as well as fixed and stained preparations of tissue cultures of chick embryos.

Fibrils appeared in cultures after 24 hours of growth. He showed that most of the cell cytoplasm was drawn out into long processes. These processes were continued back as a

“delicate thread” passing as many as twelve cells and always extended in the direction of cell migration. They contained mitochondria and other granules and extended through a bundle of fibrils. It was suggested that these fibrils were originally in the exoplasm of the

cell. However, they were never observed to change into connective tissue fibrils. In the

fixed and stained preparations, these delicate fibrils appeared first as more refractive lines

within the cytoplasm of individual cells and later continued from one cell to another 16

through the exoplasm of the cell processes. Fibrils eventually appeared more independent of the cytoplasm and were extended across several cells. A few mitochondria were usually found within bundles of fibrils and this was considered as the most convincing argument in favour of the intracellular origin of fibrils. Later, during the cell migration, the long processes were drawn back into the cell. Most remarkably, it was mentioned that “during the mitosis of one of these cells all the delicate network connected with the cell was partly drawn into it, and the space around the cell became free from network. It thus became clear that the protoplasmic network between these cells was not extracellular in origin”.

Furthermore, it was shown that fibril growth was sensitive to changes in its environment by cell contraction. It was mentioned that “Often, coincident with the contraction, there occurred a rolling-back of the edge of the growth, and in this case when the cells migrated out again many of them became changed in their relative positions. Thus it is evident that a decided strain is present during the development of the fibrils”. However, he did not detect any suddenly formed fibrils and stated that existing fibrils were still in the same state of development as that in which they were before the contraction took place.

Wolbach and Howe (1926) [112] and later Wolbach (1933) [113] followed histologic sequences in development of connective tissue of guinea pigs during scorbutic condition and immediate reparative processes following administration of antiscorbutic. While no collagen was found in scorbutic condition, fibroglia fibrils were present and stained sharply. It was shown that cytoplasmic vacuoles of in scorbutic condition discharged an extracellular liquid. After recovery from scorbutic condition, a large volume of intercellular collagen fibrils appeared after 24 hours. In case of isolated cells, the 17

collagen fibrils were near the cell body and parallel to its processes; in cell clusters, fibrils

occurred irregularly between cells without processes. Therefore, it was suggested that rapid

appearance and large volume of intercellular collagen fibrils after recovery from scorbutic

condition is due to presence of a liquid precursor of collagen in the extracellular space; and

that the collagen fibril formation is influenced by forces acting on this homogeneous

collagen. It was also suggested that collagen alignment and distribution is determined by

the shape of the cell and its processes.

Maximow (1928) [114] studied development of collagenous fibrils in cultures of adult

connective tissue of the rabbit in the living condition as well as after fixation and staining

for a period of 2 to 3 weeks. In the first stages, delicate loose networks of thin fibrillae appeared around star shaped fibroblasts (“The fibrillae are argyrophile, i.e., they are electively impregnated with silver”). Nothing was observed supporting the idea of transformation of cellular protoplasm or exoplasm into collagen. It was suggested that fibrillae form as the result of precipitation or transformation of some colloidal under the influence of chemical factors provided by fibroblasts. In later stages, fibrils became arranged in parallel, lost the argyrophilia and stained in the fashion of mature collagen.

Stearns (1940) [115, 116], in an impressive study, microscopically followed the progress

of a healing wound in the connective tissue of a living rabbit’s ear from the earliest stage

of fibroblastic invasion to the final stage of completion of a dense fibrous tissue. Fibroblasts

invaded the healing area after 6 days, and delicate fibrils appeared 4-5 days later. While it was difficult to see the entire fibroblast cell boundary in regions where fiber formation was about to occur, she reported several refractile threads running between the cells through 18

their processes. She described the unusual activity of fibroblast, possibly as a result of

tension produced by contraction of epithelium, as follows: “the cells frequently shot out

long blunt processes or retracted them, simultaneously breaking their connections and

forming new ones, so that the same cell appeared at one time elongated with long fiber-

like processes, and later shortened with rounded bulbous ends.” The earlier fibrils never

separated from the fibroblasts surface to become intercellular connective tissue fibrils.

Although the vacuoles became numerous and active in the fibroblasts about to form the

intercellular connective tissue fibril, they were never observed to participate directly in this

process. The intercellular connective tissue fibrils formed extracellularly as a result of fibroblastic activity. The fibroblasts participated directly in the process by the projection of cytoplasmic material from their surface. Since this cytoplasmic material disappeared as the fibrils formed, it was suggested that it was utilized in the production of fibrils. It was proposed that the development and arrangement of fibrils are influenced by tension and orientation of fibroblast cells. Stearns proposed that the exoplasmic fibrils, described by

authors who support an intracellular origin of connective tissue fibrils, are intracellular

fibroglia fibrils described by Mallory (1903) [103].

By this time, it was generally accepted that fibrils are formed in the intercellular space

[117]. While previous observations were mostly limited by the resolving power of the light microscope, the higher resolution power of electron microscope made it possible to study the fine structural detail of the interrelationship between the cells and collagen fibrils and investigate the origin of fibrils more rigorously. However, the advent of the electron 19

microscope only prolonged the controversy due to the static nature of images and

destructive sample preparation procedures.

Before we review further studies, it is important to mention that early studies with the

electron microscope [71] showed that adult collagen fibrils from a variety of sources have

the same characteristic periodic structure of ~64 nm, which was previously predicted by x-

ray diffraction analysis of native air dried collagenous tissues [118]. However, some fibrils

showed different periodicities, deviating from the accepted ~64 nm. Native fibrils with 17-

25 nm periodicity were found in embryonic rat tail tendon [119]. Smaller fibrils in cultured cells also showed a periodicity of 21-27 nm [117]. In vitro studies of acid soluble collagen revealed that the fibril periodicity depends on conditions such as pH, duration of reaction

[81], salt concentration [84], or presence of other ECM molecules [59, 119]. Furthermore, it was shown that fibril periodicity can be altered during sample preparation such as mechanical extraction of fibrils [71] or pH of tissue environment during the preparation process [120]. Remarkably, studies of replicas of dry and moist fibrils showed that even though moist fibrils adhered to surface and flattened on drying, the axial periodicity was persevered [121].

Jackson (1954) [122] studied the mechanism of formation and early development of connective tissue with the electron microscope. Filamentous structures were demonstrated within the cytoplasm of fibrogenic cells, but their nature and chemical composition were not confirmed. It was suggested that these filaments might 1) be the immature embryonic collagen fibrils, 2) be precursors of collagen fibrils (assuming different molecular units for these filaments and collagen fibrils), 3) form a core material for further deposition of 20

collagen, 4) be associated with the ground substance, or 5) be associated with the

biochemical system that produces collagen fibrils.

Wassermann (1954) [123] studied the fibril formation in ultrathin sections of fixed rat

tendon material. Primary fibrils, 10-20 nm in diameter and immediate product of fibroblasts, were found at the surface of the fibroblasts and also within the cytoplasm. More mature fibrils, with ~65 nm periodicity and 40-50 nm diameter, were found after one week,

but not as closely related to the cells as the primary fibrils. Observation of isolated primary

fibrils by ultrasonic treatment, raised the probability of fusion of primary fibrils. Therefore,

it was suggested that mature collagen fibrils form by aggregation of primary fibrils within

the ground substance with no intimate developmental relationship to the cells, and that

primary fibrils are not merely precursors of the mature fibrils but rather building-blocks of them.

Gross et al. (1955) [124] showed that collagen can be extracted from connective tissues by natural salt solutions. The tissue residues after salt extraction revealed normal and not swollen collagen fibrils, while after acid extraction the fibrils structure were badly disturbed. The salt extracted collagen precipitated irreversibly as typical cross-striated fibrils by increasing the temperature to 30°-37° C. Therefore, they suggested that the salt extractable collagen is derived from extrafibrillar sources, exist in a dispersed state in the tissue, had not yet been incorporated into the formed fibrils, and is a precursor of fibrils.

They speculated that fibrils form by precipitation of these precursors, which are secreted by cells and dispersed in the ground substance, under the influence of environmental factors. 21

Bradbury and Meek (1958) [125] studied fibrillogenesis of collagen in leech. They reported vesicular components just below the cell surface and small fibrils, 25 nm in diameter with

30 nm periodicity, close to long processes of cells. They concluded that the material comprising the fibrils is secreted just below the cell surface and fibrils shed from the cell surface.

Porter and Pappas (1959) [126] studied the relation between collagen fibril and fibroblast of the chick embryo during fibrogenesis with the electron microscope. Collagen fibrils were never observed within the cell. However, it was indicated that they form in close association with the cell surface. Since, I) fibrils were arranged parallel to the line of pull,

II) fibrils remained in close association with the cell surface even after vigorous preparation

process, and III) the fibril concentration was directly proportional to the cell concentration,

it was suggested that fibrils are formed by polymerization of the material available at the

cell surface, coinciding with the stress fibers (fibroglial) within the cell cortex, and then

shed into the intercellular spaces and there grow to limited diameters by accretion of

materials from the general milieu. They proposed that the mechanism for differentiation of

the original unit fibril is distinct from the process of gradual growth and maturation of

fibrils, that the original unit fibrils represent the core upon which subsequent layers of

monomeric collagen are deposited and grow through surface accretion. Still, it was

mentioned that formation of fibrils outside of and some distance from the cells in vivo is

not deniable since these experiments were done in cultures.

Godman and Porter (1960) [127] studied formation of connective tissue matrix in the

developing cartilage of fetal rat. Microfibrils were observed within the ectoplasm subjacent 22

to the cell membrane in the electron microscope images. Therefore, it was supposed that

fibrillogenesis takes place within the cell membrane. It was suggested that the primary

fibrils shed into the matrix and serve as cores of mature fibrils which may grow in the

matrix by incorporation of soluble collagen (tropocollagen units). However, it was also mentioned that the observed fibrils may represent the fibroglia or stress fibers of classical . The authors also considered that the apparently ectoplasmic or cortical fibrils might actually be extracellular or surface-adherent fibrils which are projected onto the ectoplasm.

Chapman (1961) [128] studied the manner in which the collagen protein is released from

the cell in the developing connective tissue of guinea pigs. Fine filaments, about 5 nm in

diameter, were found in the outer regions of the cytoplasm of fibroblast cells. These

filaments tended to lie parallel and closely adjacent to the cell boundary which was usually

disrupted near regions where filaments occurred. These filaments were also exhibited in

the ECM space and showed a periodicity of about 50 nm. Mature extracellular collagen

fibrils also showed a 50 nm periodicity which was suggested to be due to shrinkage during

preparation and to compression of the sections on cutting. Therefore, it was suggested that

the fine fibrils are connective tissue fibril precursors and are released from the fibroblasts

by disintegration or dissolution of the cytoplasmic membrane near elongated cell

processes. Nonetheless, it was discussed that the presence of filaments in fibroblasts may

be an artefact of fixation and made by aggregation of material in the cytoplasm during

sample preparation. 23

Peach et al. (1961) [129] studied tendon regeneration in guinea pigs using light and electron

microscopy. Long and “migratory” fibroblasts were found in the advancing edge of the

tissue. After 4 days, fine filaments (~10 nm in diameter) were found mostly in close

proximity to round and synthesizing fibroblasts. Striated collagen fibrils were never found

in the intracellular spaces. The early collagen fibrils were randomly oriented but later

assumed a more orderly array.

Sheldon and Kimball (1962) [130] described a pattern of banded structures in Golgi

vacuoles. The electron micrographs did not have sufficient resolution to permit complete

characterization of the periodic structure. However, they were reminiscent of the fibrous

long-spacing (FLS) collagen [84]. They proposed several explanations for this observation.

Precipitation of collagen could be an accident that occurred during fixation; it could be due to presence of other substances such as chondroitin sulfate; or it could be due to tendency of liquid crystals to orient themselves on solid surfaces such as membrane of the Golgi vacuole. The role of mechanical forces was also considered as a possible mechanism for orientation of long molecules such as tropocollagen: “A tendency to orientation and

alignment of long molecules is facilitated by various stresses, such as a mechanical force

applied externally to any liquid in any capillary system”.

Goldberg and Green (1964) [131] studied collagen synthesis of cultured mouse fibroblast

by electron microscopy. They found some fibrils, approximately 5 nm in diameter,

throughout the cytoplasmic matrix before day 4. However, hydroxyproline data did not

show evidence of any collagen production. After day 4, 5-10 nm collagen fibrils were

found in ECM. Initial extracellular fibrils were non-periodic and 4-16 nm in diameter. 24

Thicker fibrils, 16-25 nm in diameter, with asymmetric periodicity were observed after day

7. Proportion of periodic fibrils were increased over time. However, non-periodic fibrils were found at all-time mostly near the cell surface. In horizontal sections, fibrils with and without periodicity were found near lower density cell membrane. They argued that the unusual cell surface is due to horizontal sectioning of flattened cells and is not related to collagen fibril shedding into ECM. Also, they demonstrated that non-collagen synthesizing cells before day 4 show the same phenomenon in horizontal sections. They suggested that extracellular elements can be projected onto the cytoplasmic plane and appear to be intracellular due to the depth of field of the electron microscope. Therefore, they concluded that soluble collagen is delivered to the cell surface in secretory vesicles and discharged in molecular form from the cell through fusion of secretory vesicles membranes with the cell membrane.

Autoradiography and electron microscopy studies gave their impressive evidence of the route by which collagen are contributed to the ECM. Previously it was shown that the hydroxyproline cannot incorporate into the proteins, but rather is derived from the oxidation of proline which is already bound [132-135]. Similar results were obtainable for hydroxylysine; it was shown that collagen hydroxylysine forms from lysine [136], but not from free hydroxylysine [137]. Therefore, labeled amino acids, mostly proline and hydroxyproline, became great indicators of collagen.

Jackson and Smith (1957) [138] studied the biosynthesis and formation of collagen in fibrin-free tissue cultures of osteoblasts. During the growth of the culture and before appearance of any fibrils or filaments between cells, high resolution electron microscopy 25

demonstrated a particular fine substance in the medium. At a later stage, a close network

of long cytoplasmic processes and long fine fibrils appeared between adjacent cells. These

fine fibrils were less than 10 nm wide and had a 7-8 nm periodicity. After 45 hours, fine

fibrils with a periodicity of 21 nm and a diameter of 20-30 nm stained like collagen. Later,

the electron microscope demonstrated the presence of collagen fibrils with a periodicity of

64 nm. On the other hand, a high initial protein-bound hydroxyproline content was

observed during the first 24 hours of growth, before appearance of fibrils. Formation of

banded fibrils was not associated with a significant rise in the mean hydroxyproline content

of the tissue and it was concluded that the appearance of a hydroxyproline-rich collagen

precursor precedes that of banded fibrils. Therefore, it was suggested that cells synthesise

and secrete a hydroxyproline-rich precursor of protein which becomes directly transformed

into collagen fibrils.

Ross and Benditt (1961) [139] studied collagen formation in healing guinea pig skin wounds. Fibroblasts first appeared at 24 hours and had enlarged mitochondria.

Intracytoplasmic filamentous material, 5-7 nm in width, were observed near the cell surface, but no banded fibrils were present within the cytoplasm. Collagen fibrils were first seen at 3 days extracellularly near the cell surfaces. These collagen fibrils with 60-70 nm

periodicity were narrower immediately adjacent to the cell than those which were farther

displaced. They indicated that the role of the intracytoplasmic filaments in collagen

formation is unclear for these reasons: 1) the intracytoplasmic filaments were not banded,

2) many other cell types contain intracytoplasmic filaments with similar appearance, and

3) there is no morphologic evidence of the manner in which the cell re-establishes 26

continuity of its plasma membrane after fibril release. Therefore, it was suggested that a

soluble collagen precursor is secreted and polymerized, both free of and dependent upon

the fibroblast.

In their following work, Ross and Benditt (1962) [140] studied the sequence of

incorporation and utilization of tritium-labeled proline in healing wounds from normal and

scorbutic guinea pigs. The results confirmed the utilization of proline by the fibroblasts

and its incorporation into collagen in normal animals and into non-collagenous material

(filamentous non-banded substance) in scorbutic animals. In both groups the proline reached a maximum over the cytoplasm within 4 hours. The proline disappeared from the cell and reached a maximum over the collagen in normal animals and extracellular fibrillar

material in scorbutic animals by 24 hours. It was concluded that the fibroblast synthesizes

the “collagen unit” and secretes it into the extracellular space where fibril aggregation takes

place.

Revel and Hay (1963) [141] used radioactive amino acids, H3-proline, as a precursor for

localization of collagen synthesis and secretion in developing limbs of Amblystoma

maculatum larvae. It was shown that collagen synthesized in the ergastoplasm, transported

to the Golgi complex, and left the cell 1-2 hours after exposure of the limb to H3-proline.

Two hours after the administration of the labeled amino acid, the labeled protein was

moved to extracellular space by fusion of secretory vacuoles with the cell membrane

(Figure 1.7). The matrix appeared to be homogeneously labeled at 2 hours which indicated that the new secreted collagen into the matrix had diffused away from the cell surface and did not accumulate adjacent to the secretory cells. Furthermore, they proposed that the 27

small filaments in the matrix and those seen in Golgi vacuoles may be monomeric collagen

and the larger fibrils in the Golgi vacuoles are formed during fixation and embedding of

the tissue or that they represent a reversible aggregation of tropocollagen molecules.

Figure 1.7. Diagram summarizing the synthesis of the collagen of the cartilage matrix. The black arrows indicate the progression of the H3-proline as it enters the cell, packaged in secretory vacuoles in the Golgi zone, and then released from the cell. The newly formed collagen diffuses some distance from the cell before polymerizing with striated fibrils. Reprinted with permission from Revel and Hay (1963) [141].

Voelz (1964) [142] found intracellular rod-like fibers consisting of a bundle of approximately 50 single fibrils, about 5 nm in diameter, in cultured of chicken fibroblasts.

The fibers were enclosed by a membrane of ~60 nm in width and undetermined length.

The nature of these fibers were not confirmed since no cross-banding was detected. It was

suggested that they may be a phagocytized or secretion of the fibroblast.

Porter (1964) [143] in a review of these early electron microscopy and autoradiography

studies suggested that fibroblast probably contributes to the collagen formation by more

than one type of secretion. He wrote that “A Golgi-derived vesicle migrates to the cell 28

surface, fuses with it, forms an opening or stoma, and discharges its contents. These diffuse away to be incorporated into the growing fibrils of collagen in the surroundings. This story seems well established … But is this the only manner in which a fibroblast contributes of itself to its environment?”. He supported the existence of primary fibril structures associated with cell processes or immediately adjacent to them: “The striking diameter differences in mature collagen fibrils found in different tissues argue for differences in the

"core" structure of the fibrils, differences which could limit the amount of tropocollagen incorporated in the growth of the mature fibril”.

Welsh (1966) [144] for the first time demonstrated intracytoplasmic banded collagen fibrils in high resolution electron micrographs of desmoid fibroma fibroblasts. Collagen fibrils were found within narrow tubules of Golgi complexes. However, no evidence of fibrils exiting the cells was found. It was suggested that the intracellular collagen fibrils are due to pathological conditions involving rapid synthesis of collagen.

Trelstad (1971) [145] reported two distinct types of vacuoles in the embryonic chick corneal epithelium. One type of vacuoles, only infrequently observed, contained cross- striated aggregates. They were located near the basal cell surface of the basal cells and were

300-700 nm in diameter. The aggregates were 20-60 nm in width and 300-600 nm in length and located in the center of the vacuole. Another type of vacuoles had an elongated shape with 100 nm in diameter and 600 nm in length. They contained a dense, slightly fibrillar material which were found near the vacuoles containing collagen-like aggregates. It was proposed that synthesized collagen in the endoplasmic reticulum, is transported to the

Golgi apparatus by small vesicles, packaged and condensed in vacuoles, and excreted by 29

fusion of the vacuole with the cell surface membrane. It was suggested that intracellular collagen, in the process of concentrating and destined for excretion, precipitated to form aggregates during fixation for electron microscopy.

Figure 1.8. Residual bodies found in a guinea-pig fibrocyte (left) and a developing mouse ligament fibroblast (right) containing coiled banded fibrils. From Ten Cate (1972) [146].

Ten Cate and co-workers in a number of studies demonstrated that intracytoplasmic fibrils

are evidence for the ability of fibroblast to phagocytose its own product [147]. Ten Cate

(1972) [146] studied the fibroblast cells associated with rapid connective tissue breakdown

in the guinea-pig periodontium and with rapid connective tissue remodelling in the

developing mouse periodontal ligament and monkey gingiva. Intracytoplasmic,

membrane-bound, and banded collagen fibrils were observed. Frequently a loss of banding

of the collagen fibril was observed and in some instances, membrane-bound structures

contained coiled banded fibrils (Figure 1.8). It was suggested that fibroblasts phagocytose

collagen fibrils, and that the final degradation of collagen occurs intracellularly when tissue

is rapidly remodelling. It was also suggested that excessive collagen synthesis during rapid 30

remodeling can lead to intracellular aggregation of collagen which might become subsequently degraded intracellularly as well. Similar intracytoplasmic fibrils were observed in mouse periodontal ligament (a connective tissue structure with a high rate of remodelling) [148] and the junctional zone between old and new connective tissue of skin wounds (a site of active remodelling) [149].

In a following study Deporter and Ten Cate (1973) [150] demonstrated the presence of acid phosphatase and alkaline phosphatase activity within collagen containing profiles of mouse ligament fibroblasts which indicated collagenase activity. Some collagen fibrils were observed situated partly within the fibroblast and partly outside of it, suggesting that the intracellular collagens were once extracellular. Later, Ten Cate and Syrbu (1974) [151] demonstrated the activities of alkaline and acid phosphatase in fibroblasts of the connective tissue of erupting teeth. Sections stained for acid and alkaline phosphatases activity confirmed alkaline phosphatase activity on the cell membrane and in the clear phagosome, but not acid phosphatase activity in the phagolysosome. This pattern of enzyme activity supported the idea of phagocytosis of collagen by fibroblasts (Figure 1.9).

Listgarten (1973) [152] also found banded intracytoplasmic collagen fibrils in fibroblasts of hamster periodontal ligament. From the nature of tissues in which intracellular fibrils have been detected, it was suggested that these fibrils are most likely ingested collagen during high turnover rate of connective tissue.

Dyer and Peppler (1977) [153] demonstrated intracellular collagen fibrils within membrane-bound cytoplasmic vacuoles of rat uterus cells. The fibrils were sometimes tightly packed and linear and sometimes randomly arranged. Since cytoplasmic vacuoles 31

contained ill-defined debris and poorly-visualized structures with a periodicity, it was

suggested that they represent a final phase of fibril breakdown.

Figure 1.9. Diagram illustrating the fibroblast phagocytoses extracellular collagen and replacement of collagenase activity by lysosomal enzyme activity. 1) Phagocytosis of extracellular collagen fibrils (the black dots represent activity of collagenase). 2) The collagen fibril is in the fibroblast in a phagosome. 3) The collagen fibril is now within an electron-dense phagolysosome derived from fusion of lysosomes with the phagosome. From Ten Cate and Syrbu (1974).

Gotjamanos (1979) [154] showed intracellular collagen fibrils in fibroblasts of a ameloblastic fibroma. Fibrils were located either entirely within or totally outside of cells; no cytoplasmic extensions were observed towards collagen fibrils. The intracellular collagen fibrils observed within narrow tubules and closely resembled extracellular fibrils with 64 nm periodicity. Collagen phagocytosis and degradation by fibroblasts were considered to explain the intracellular fibrils. 32

Michna (1988) [155] from experiments with tendon fibroblasts and fibroblastic tumour cells found intracellular collagen fibrils (with 55 nm periodicity) in hormone-treated . These fibrils were not always oriented parallel to the extracellular collagen fibrils.

Due to the distinctive banding pattern of these fibrils and their intravascular location, it was suggested that intracellular fibrils may be attributed to a specific procollagenase acting on newly synthesized procollagen. Therefore, the highly metabolising fibroblasts accumulate collagen precursors exceeding the secretory transport capacity of the cell and inducing compensative autophagocytic reactions.

Studies of Beertsen, Everts, and colleagues also indicated that collagen fibrils enclosed in lysosomal vacuoles of fibroblasts represent collagen that has been phagocytosed from the extracellular space [156]. Everts et al. (1989) [157] showed that metalloproteinase enzymes do not play a crucial role in the phagocytosis and intracellular digestion of collagen fibrils by fibroblasts. Therefore, it was suggested that collagen digestion can be an extracellular collagenase-dependent pathway or an intracellular collagenase-independent pathway. It was shown that intracellular cross-banded collagen fibrils appear even when collagen synthesis is blocked [158-160] and that cytoplasmic actin filament systems are involved in the phagocytosis of collagen [157, 158].

Trelstad and Hayashi (1979) [161] found deep recesses in the fibroblast cell surface which contained collagen fibrils. The recessed fibrils were surrounded by a plasma membrane and resembled the extracellular fibrils in diameter, staining pattern, and electron density.

Autoradiographic analysis showed that [3H]-proline progressed from the cisterns of the endoplasmic reticulum (5 and 15 min) to the vacuoles of the Golgi apparatus (30 and 45 33

min) to the extracellular space (1 and 4 hr). It was concluded that the materials detected

from 45 to 60 minutes in the recesses were recently synthesized. Therefore, recessed fibrils were supposed to be in the process of deposition in the extracellular space. A model for intracellular collagen processing was proposed in which procollagen forms in the endoplasmic reticulum, moves to the Golgi apparatus for packaging and condensation, and fibril assembly occurs in close association with deep recesses of the fibroblast cell surface by the addition of collagen aggregates to the end of the fibril (Figure 1.10).

Figure 1.10. Mode of collagen fibrillogenesis in embryonic chick and mouse tendons by Trelstad and Hayashi (1979). Fibrils assembly occur at recesses of the fibroblast cell surface. Endoplasmic reticulum (er); Golgi apparatus (ga); condensation vacuoles (cv). Reprinted with permission from Trelstad and Hayashi (1979) [161].

Marchi and Leblond (1983 and 1984) [162, 163] examined whether collagen is assembled into fibrils within or outside of fibroblasts. The collagen biogenesis and assembly into fibrils of the rat foot pad connective tissue was investigated by electron microscopy and by 34

radioautography. Quantitative radioautography after injection of 3H-proline revealed that collagen precursors arise in the rough endoplasmic reticulum, transfer to Golgi by intermediate tubules or vesicles, straighten out into cylindrical rods (procollagen), and freed from its saccule to become a secretory granule, respectively. The procollagen content of the granule is then released outside of the cell. While intracytoplasmic collagen fibrils were observed in serial sections of fibroblasts, it was emphasized that these intracellular fibrils did not contain the new labeled proline. However, some lysosomal elements were labeled later. It was shown that the intracellular fibrils were associated with lysosomes and

digestive vacuoles since they had lost their banding and were at various stages of

degeneration. Thus, it was concluded that the long controversy is settled in support of the

views that the assembly of collagen precursors into fibrils does not occur within cells but

is entirely extracellular.

Birk and Trelstad (1984) [164] used high voltage electron microscopy to study collagenous

architecture in developing chick corneas. A model for regulation of collagen fibril, bundle,

and lamellar formation was proposed in which the corneal fibroblast cell surface defined

three major extracellular compartments: 1) Small recesses, containing 1-12 collagen fibrils,

in association with the fibroblast cell surface; 2) larger surface folding containing small

fibril bundles that contained 50-100 collagen fibrils; 3) large surface compartment in which

bundles joined into larger bundles and lamellae (Figure 1.11). It was suggested that the fibroblast formed bundle will be separated from the cell surface by migration of the fibroblast away from the bundle or the fibroblast “spin” the fibril bundle from its surface.

It was concluded that collagenous matrix does not simply "crystallize" onto the epithelial 35

template, instead the transfer of the epithelial template to the stroma occurs under “unique

spatial organizing capacity” of the fibroblast.

Figure 1.11. Proposed model of the compartmentalization of the extracellular space by the corneal fibroblast. Reprinted with permission from Birk and Trelstad (1984) [164].

Birk and Trelstad (1986) [165] also proposed a model for formation of extracellular

compartments during tendon morphogenesis. In this model, through a process of

compound exocytosis, secretory vacuoles containing procollagen molecules fuse with the

cell surface and adjacent vacuoles to produces the fibril-forming compartments (recesses).

Recesses fuse with the cell surface and one another to form the bundle-forming

compartment. Finally, with retraction of cytoplasm the bundles laterally associate (Figure

1.12). Yang and Birk (1986) [166] used freeze fractured chick embryo tendons and scanning electron microscopy (SEM) to show the topographies of these extracytoplasmic compartments in developing chick tendon fibroblasts (Figure 1.13). 36

Figure 1.12. Proposed model of the compartmentalization of the extracellular space by the tendon fibroblast. Reprinted with permission from Birk and Trelstad (1986) [165].

Figure 1.13. Proposed model of the compartmentalization of the extracellular space by the tendon fibroblast based on SEM images. Reprinted with permission from Yang and Birk (1986) [166]. 37

Ploetz et al. (1991) [167] showed extracytoplasmic compartmentalization of chick embryo using transmission electron microscopy (TEM). Three levels of extracellular compartmentalization were shown: 1) narrow channels containing single or small groups of collagen fibrils; 2) fibrils grouped as small bundles in close association with the cell surface; and 3) laterally associated bundles (only in more mature dermises).

Figure 1.14. Fibripositor model proposed by Canty et al. (2004) [12]. Reprinted with permission.

Canty et al. (2004) [12] using serial section and 3-D reconstructions of chick embryonic tendon fibroblasts showed that fibril formation can occur in closed intracellular Golgi to plasma membrane carriers (GPCs). These GPCs were shown to be on the way to plasma 38

membrane protrusions, which were named “fibripositors” (fibril depositors; Figure 1.14).

Also, using pulse-chase experiments it was shown that procollagen can be converted to collagen within the GPCs. It was shown that fibripositors were always oriented along the tendon axis, and they were absent at postnatal stages and in cultured cells. Therefore, it was suggested that fibripositors maintain the parallelism of collagen fibrils in tendon when seeding of the ECM occurs.

Figure 1.15. The processes of collagen fibril nucleation and movement in the fibripositor model proposed by Kalson et al. (2013) [168]. Reprinted with permission. 39

The fibripositor model was revised by Kalson et al. (2013) [168] in a study of serial block

face-scanning electron microscopy images of the cell-matrix interface (Figure 1.15). It was shown that fibricarriers can be generated by out-to-in fibril transport and that nonmuscle myosin II (NMII) powers the transport of newly formed fibrils at fibripositors. A NMII- dependent cell-force model was proposed for the creation and dynamics of fibripositor structures in which fibrils in fibricarriers could grow to become fibrils in recessed fibripositors, then protruding fibripositors, and then released to the ECM. It was suggested that the initial collagen fibril nucleation can occur at the plasma membrane by accretion of collagen molecules or collagen aggregates. In this model fibricarriers could be a type of

Golgi-to-plasma membrane carrier or might be formed by the inclusion of short early fibrils that are not anchored in the ECM.

Paten et al. (2016) [42] demonstrated how tension can directly drive initial fibrillogenesis and maintain continuity of load-bearing collagenous tissue. It was showed that fibers can be formed by slowly drawing a microneedle from a slightly concentrated surface of a collagen solution droplet. A model for early connective tissue development was proposed in which extensional strain triggers fibril formation extracellularly into the path of force.

As the fibrils lengthen, strain rate between the initial fibrils amplifies and fibrils’ ends

which are separated by small gaps rapidly fuse by flow-induced crystallization (Figure

1.16). It was estimated that the required collagen concentration is locally achievable by

cells and cell contractions could generate equivalent flows. The strong suit of the flow- induced crystallization model is 1) its consistency with most observations of tendon morphogenesis such as abundance of short fibril segments during initial tendon 40

fibrillogenesis and their end-to-end growth, 2) providing a mechanism to grow collagenous

structures into the path of force, where it is needed, and establishing long-range

connectivity, and 3) synchronizing the role of , fibronectin, actin filaments, and in cooperation with local and global strains which have been all shown previously to be necessary for collagen fibrillogenesis.

Figure 1.16. Flow-induced crystallization model by Paten et al. (2016) [42]. Reprinted with permission. 41

Hyaluronic acid is synthesized by fibroblast cells besides secretion of collagen [169]. A

hyaluronic rich solution provides a viscous environment between the closely packed cells

to diminish the loss of collagen molecules by diffusion. This would maintain the local high

concentrations of collagen required in the flow-induced crystallization model and favour

the formation of fibrils. Remarkably, this role of hyaluronic rich was suggested in 1964 by

Goldberg and Green [131]. Furthermore, collagen fibrillogenesis dependency on integrins

[170], actin [171], and fibronectin [172] is well known. Disrupting the binding site of collagen-fibronectin is sufficient to eliminate collagen network assembly [173] and it has been hypothesized that the primary role of the collagen-fibronectin molecular interaction is to catalyse ECM network assembly [174]. Paten et al. (2019) [174] further demonstrated that the addition of fibronectin reduces that critical collagen concentration required for flow-induced crystallization of collagen fibers.

Lu et al. (2018) [175] studied type I collagen assembly dynamics in osteoblasts culture using live imaging of fluorescently labeled procollagen. The labeled collagens assembled into extracellular fibrils in the presence of ascorbate, while they remained intracellular without ascorbate. It was showed that collagen fibrillogenesis is tied to cell-generated forces. Fibril networks were continually stretched and physically reshaped by cell motion.

As a possibility, it was suggested that mechanical stretching of fibrils could unmask

binding sites on the molecules and promote molecular interactions (a mechanism that has

been shown for fibronectin). Using co-cultures of two different coloured collagen

expressing cells (mCherry- and GFPtpz-collagen) and double immunogold staining it was showed that individual collagen fibrils were formed by more than one cell. Therefore, it 42

was concluded that if fibrils were formed intracellularly, then initial fibrils formed by

different cells had to laterally associate to one another to explain the co-localization of

green and red collagen molecules on the same fibril.

It has been mostly accepted now that fibripositors are the site of fibril assembly in vivo

[18]. The original fibripositor idea was suggesting that fibril segments are formed

intracellularly and then discharged into extracellular space [12] without considering the

role of mechanics in either collagen assembly or fibril fusion. The fibripositor model has

been restructured and now suggest that fibrils are pulled toward the cell by nonmuscle

myosin II mechanism [168]. However, it still can’t explain synchronized alignment of

collagen fibrils with actin-rich cytoplasmic protrusions far from the main cell body [176].

Furthermore, and besides all the evidence regarding the phagocytosis of collagen fibrils in

rapidly remodeling tissues that were mentioned earlier, the fibripositor theory is puzzling

and can be brought into question regarding intracellular processing of procollagen. It has

been known that removal of the carboxyl propeptides lowers the solubility of procollagen

[177] and is an essential step for the assembly of collagen fibrils [178, 179]. Humphries et

al. [180] showed that procollagen processing can occur within intracellular compartments

of postnatal tendons. However, I) the enzymes for procollagen cleavage have been

previously only detected within the extracellular culture medium [66, 181-183] and not

extracts of the cells [184]; II) the required ionic calcium concentration for enzyme activity

[181] is orders of magnitude larger than intracellular calcium concentration [185]; III) the procollagen proteinases are neutral metalloproteinases [66, 182-184, 186] and have negligible activity at pH 6 or below [181, 187]; IV) no evidence of intracellular fibrils or 43

fibripositors were found even after the detected intracellular procollagen processing [180];

V) the acidic pH of Golgi network transport carriers and secretory vacuoles is incompatible with required condition for fibrillogenesis of collagen molecules and might even lead to formation of fibrils with disturbed structure and banding patterns.

1.3.2 Fibril Growth Mechanism

Like the site of initial fibril formation, the exact mechanism of fibril growth during development, ageing, and in pathological conditions is not clear and has been the subject of controversy. In vitro reconstitution of fibrils from collagen solutions and factors affecting the rate of precipitation and growth of collagen fibrils (such as pH, ionic strength, temperature and collagen concentration) are well studied. However, the manner in which molecules or fibril segments add to the growing fibril in vivo is not resolved. While reconstituted fibrils in vitro have a broad range of diameters, fibril diameter distribution in vivo is highly controlled and has a unimodal or multimodal distribution changing with age and tissue [188].

Embryonic growth occurs by an increase in fibril number and diameter [22, 189-191], resulting in different fibril thicknesses and shapes [22]. It has been suggested that collagen fibril growth can occurs by accretion of monomers onto the fibril surface and/or by interfibrillar fusion. But, in vivo collagen fibril growth is not completely an intrinsic self- assembly process. It also involves extrinsic regulation of fibril diameter by other fibril associated molecules, growth inhibitors, and mechano-biochemical environment of collagen fibrils [192]. 44

1.3.2.1 Fibril Growth Regulators

The growth of fibrils in embryonic and neonatal connective tissues is highly regulated. In

vivo fibrils are cylindrical with uniform diameter [193]. However, the mechanism of

growth is disrupted in reconstituted fibrils in vitro where they have a broad distribution of

diameter [97]. Presence of an upper limit for fibril diameter may be due to difficulty of the

addition of new molecules or fibril segments on energetic grounds [131]. Substantial

evidence points to the participation of water molecules, propeptides, proteoglycans, and

other type of collagens in this regulatory process:

1.3.2.1.1 Fibril Growth Regulation by Stabilization of Water Structure

Collagen structure and stability is affected by its interaction with water molecules [194-

201]. Its fibril formation is an endothermic, but entropy driven process [90, 202] arising

from release of water molecules into the surrounding liquid [203, 204]. Therefore, fibril growth (either through molecular association or fibril segments fusion) may be regulated by stabilization of water structures [28]. In general, it has been shown that breakers of water structure promote collagen fibril formation, while makers of water structure are inhibitory

[205].

Mature fibrils in vivo are cross-linked by covalent bonds between neighbouring molecules.

However, the young and growing fibrils are stabilized by non-covalent hydrogen bonds

[206] and have the potential to bind more water molecules [196]. It has been suggested that regulation of fibril growth may be related to proteoglycans [207] or hyaluronate [22, 208] which can stabilize the water layer associated with the collagen molecules. Release of these 45

trapped water molecules could provide the increase of entropy required for association of

molecules into the fibrils.

1.3.2.1.2 Fibril growth Regulation by Surface-Associated Small Proteoglycans

Proteoglycans are a superfamily of molecules in ECM that are distinguished by the

covalent attachment of one or more highly negatively charged chains

to their core proteins [209]. These proteoglycans with their sulfated

can regulate matrix hydration [210], interact with collagen fibrils [211], alter fibrillogenesis

rate [212-214], and change fibril orientation [215] and fibrillar structure of collagenous

ECM [216]. They have also been shown to alter fibril diameter [217] by directly

influencing molecular assembly [21] or lateral fusion of fibrils [22].

Besides stabilizing water structures by proteoglycans, an alternative mechanism for fibril

diameter regulation has been proposed by Scott and colleagues in which surface-associated

small proteoglycans and their glycosaminoglycan chains extend around the fibril and

through steric effects limit lateral fibril growth. Scott (1980) [218] using electron microscopy showed that there is a specific interaction between proteoglycan and collagen in tendon. Some proteoglycans appeared as filaments orthogonally arrayed along the collagen fibrils and were regularly separated by the collagen banding repeat distance.

Scott et al. (1981) [22] studied developing rat tail tendons up to 126 days post-partum using electron microscopy. It was reported that: I) Collagen fibril diameters increased with age while proteoglycans were seen at the periphery but not inside collagen fibrils. II)

Proteoglycans were apart by a constant distance, always equal to collagen fibril banding 46

even though the banding was increased with aging (approximately 41-68 nm). III) There

was a decrease in the ratio of proteoglycan to collagen with increasing age. It was shown

that loss of chondroitin sulphate was coinciding with the beginning of rapid growth phase

of collagen fibrils.

Based on these observations, a three phase model of fibrillogenesis and fibre maturation in

rat tail tendon was proposed. In phase 1 (up to day 40 after conception), tropocollagen

interacts with dermatan sulphate-rich proteoglycan during or immediately after formation

of microfibrils with ~53 nm periodicity. The hyaluronate and proteoglycan-rich

environment and collagen synthesis increase the number of thin fibrils, rather than growth

in diameter of established fibrils. In phase 2 (from day 40 to approx. day 120 after

conception), concentrations of chondroitin sulphate-rich proteoglycan and hyaluronate is decreased which promotes addition of collagen to established fibrils rather than formation of new fibrils, resulting in rapid increase of fibril diameter without axial periodicity change.

In phase 3 (day 120 after conception onwards), fibril growth slows down and axial periodicity increases to ~62 nm.

Scott and Orford (1981) [219] showed that dermatan sulphate-rich proteoglycans were located almost exclusively at the ‘d’ band, in the gap zone. It was suggested that the proteoglycans may limit access to the cross-linking sites and therefore hinder fibril radial growth by molecular accretion or fibril fusion.

Scott (1984) [208] also reported a rapid decrease in concentration of hyaluronate and chondroitin sulphate during the early periods of chick and calf limb tendon development.

However, as collagen concentration increased, dermatan sulphate concentration increased 47

gradually and then declined somewhat toward maturity. The relationship [X]/[collagen]r =

k, where [X] and [collagen] are concentrations of X (i.e. dermatan sulphate) and collagen,

and r is the fibril radius was used (and confirmed based on dermatan sulphate,

hydroxyproline and collagen fibril diameters measurements) to show that dermatan

sulphate is “regularly, specifically and entirely” associated with the periphery of collagen

fibrils during most of development. The same relationship was also used to show that

conversion of pN-collagen (present at the periphery of the collagen fibril) to collagen is

complete during the developmental period studied.

Direct in vivo evidence for the role of proteoglycans in regulation of collagen assembly

and growth was achieved by development of animals deficient in small leucine rich

proteoglycans (SLRPs). All of these animals showed collagen fibril abnormities in

different tissues [217, 220-222]. For instance, Danielson et al. (1997) [217] showed that decorin-deficient mice skin and tendon had coarser collagen fibrils with irregular size and shape. Two mechanism were considered by which decorin could alter fibril diameter shape:

First, increased concentration of fibril surface associated decorin inhibits fibril fusion.

Second, decorin maintains the regular shape of fibrils while fibrils have a natural tendency to be irregular. The second mechanism is not consistent with in vitro experiment were collagen monomers form fibrils with circular cross sections and laterally associate in the absence of decorin.

Corsi et al. (2002) [223] showed that decorin and biglycan deficiency in mice had similar effects on collagen fibril structure in the dermis but not in bone. While both deficiency resulted in thinner fibrils in the dermis, fibril diameter in bone increased as an effect of 48

biglycan deficiency and decreased as an effect of decorin deficiency. Robinson et al. (2017)

[224] also showed that absence of decorin and biglycan in the mature patellar mouse tendon

was associated with larger fibril diameter.

1.3.2.1.3 Fibril Growth Regulation by pN-collagen

There are several observations suggesting that N-propeptides are confined to the fibril surface where they block accretion of further molecules [24, 225-233]. As a result, further lateral growth would be regulated by enzymic cleavage of the propeptides. The important role of N-propeptide has been observed in the studies of dermatosparaxis and Ehlers-

Danlos syndrome type VIIB. Dermatosparaxis is caused by partial loss of procollagen N-

proteinase activity [234-236]. Presence of N-propeptide on the surface of these fibrils

results in a non-circular cross sections [237]. Remarkably, it has been shown that

dermatosparactic collagen fibrils will gain a normal appearance after implantation in

normal animals [238], suggesting the existence of a dynamic mechanism for fibril growth

and degradation. Also, Ehlers-Danlos syndrome type VIIB fibrils in which pN-collagen is

only partially cleaved have rough-bordered and non-circular cross sections [239, 240].

Therefore, the growth inhibitory role of pN-collagen has been used in some suggested

models:

Fleischmajer et al. (1981) [24] showed that extension N-propeptides are associated with thin fibrils (20-40 nm) of normal skin. A mechanism of fibril growth was proposed in which fibrils form with deposition of pN-collagens and fibril growth is regulated by removal of extension aminopropeptides. 49

Hulmes (1983) [23] proposed a growth model where the inhibition of lateral growth was the result of a growth inhibitor. A quasi-hexagonal packing of molecules [241] which has been detected in tendon was assumed. In this model, growth of pN-Collagen fibrils is inhibited due to the steric blocking of interaction sites by the N-propeptides. Additional growth with 4 nm increment in radius happens when these N-propeptides are removed by enzyme degradation and a new layer of N-propeptides forms on the fibril surface. There are evidence supporting this model: It has been shown that N-propeptides on the surface of fibrils can be cleaved by N-proteinase in vitro [240] and presence of N-propeptides in ECM can down-regulate collagen synthesis [242, 243].

Chapman (1989) [27] proposed a growth model for collagen fibrils in which accretion of collagen molecules are inhibited by an inhibitor (such as, but not limited to N-propeptides) on the surface of fibrils (Figure 1.17). No assumption for lateral association of molecules and no initial packing of molecules into microfibrils were considered. In the Chapman model, the long semi-flexible molecules [244] assemble together with some fluidity in lateral positioning and some small radial tilting in a way that each molecule has to tilt

laterally through about 4 molecules (~1.3֯ radial tilt). The growth inhibitor part of the

molecules (the N-end) are confined to the fibril surface and the C-ends are buried inside

the interior of the fibril. Since the growth inhibitors cannot act as a site for further accretion,

their surface density increases with lateral growth. Growth of fibril diameter continues till

steric hindrance happens at a critical diameter. This first critical diameter depends on the

lateral width of the inhibitor segment, allowing for growth of fibrils with preferred

diameters in different tissues. When fibril reaches a uniformity at this critical diameter, 50

accretion is limited to the fibril ends and growth is only in axial direction. Lateral growth can proceed to a second critical diameter after enzymatic removal of the growth inhibitor.

Successive cycles of accretion of molecule and removal of growth inhibitor led to calculations of fibril diameters based on the size of growth inhibitor.

Figure 1.17. Chapman model. A) Schematic representation of assembling collagen molecules. B) Schematic transverse sections through a fibril assembling from pN- collagen molecules. Sections labeled 5* represent the N-propeptide section of the molecule which are larger than other sections of the molecule labeled 1-5. C) At critical diameter, the growth inhibiting domains form a continues layer on the surface of the fibril, inhibiting further accretion of molecules. Reprinted with permission from Chapman (1989) [27]. 51

Holmes et al. (1991) [245] studied morphology of sheet-like assemblies of pN-collagen.

The results showed that the overlap:gap mass contrast was increased from 5:4 (the ratio in

a native collagen fibril) to 6:4. It was concluded that the N-propeptides fold back over the

overlap zone. Analysis of the mass thickness of the sheet indicated that the N-propeptide

domains are closely packed with adjacent domains (spaced 2.23 nm center to center instead

of 1.5 nm in collagen fibrils). Therefore, it was suggested that when surface density of N- propeptides reaches a critical value, fluidity in intermolecular contacts are restricted, further accretion is blocked, and fibril growth is inhibited.

Holmes et al. (1991) [246] studied the mechanisms directing lateral packing of molecules

in co-polymers formed from approximately 1:1 mixtures of collagen and pN-collagen using quantitative scanning-transmission electron microscopy. The transverse mass distribution of control collagen fibrils was symmetrical at all parts of the fibril. The control fibrils were circular in cross-section along their lengths and the tapers were microns in length. The tip

of the co-polymer fibrils was still symmetrical. However, the cross-sectional shape of the central shaft of the co-polymer fibrils was altered. It was suggested that the pN-collagen has a high affinity for the growing fibril. Since disruption of the transverse mass distribution occurred only at larger diameters where the surface-area/volume ratio is low, it was suggested that there is sufficient fluidity in the fibril structure to allow the N- propeptide domains remain at the fibril surface.

Watson et al. (1992) [240] studied copolymerization of collagen and abnormal pN-collagen from Ehlers-Danlos syndrome type VIIB. In Ehlers-Danlos syndrome type VIIB, procollagen processing is incomplete since proα2(I) chains lack the sequences containing 52

the N-proteinase cleavage site. Individuals with Ehlers-Danlos syndrome type VIIB have near circular fibrils with accumulated pNα2(I). The results showed that collagen and the abnormal pN-collagen copolymerize into fibrils with asymmetric distribution of mass in transverse cross-section. Treatment of these fibrils with N-proteinase resulted in partial cleavage of the abnormal pN-collagen in which all the pNα1(I) chains were cleaved but the abnormal pNα2(I) chains remained intact. Therefore, it was concluded that all the N- propeptides were located at the fibril surface. Furthermore, N-proteinase treatment resolved the circular cross-section of fibrils suggesting that there is considerable fluidity between collagen molecules in a fibril.

It has been also shown that fibrils lose their circularity as proportion of pN-collagen [247] or fibril size increases [246]. However, larger fibrils in healthy adult tissues do not contain

N-propeptides [24]. Therefore, pN-collagen cannot be the only growth regulating factor.

1.3.2.1.4 Fibril Growth Regulation by Collagen Type III

Other type of collagens are synthesized simultaneously with type I collagen in various cell lines [248-251]. The structural similarities of fibril forming types allow them to polymerize within the same fibrils [252]. However, structural differences between them may add additional information in case of fibril formation or growth. An unbalance in the ratio of type I and type III collagen has been reported in osteogenesis imperfecta [253], Ehlers

Danlos syndrome type IV [254], and fibrotic processes [255]. Therefore, studies have been performed to further illustrate the role of collagen type III: 53

Fleischmajer et al. (1990) [256] performed double labeling immunofluorescence and

immunoelectron microscopy with antibodies directed against the collagen molecule and

the aminopropeptide domains of type I and type III procollagens in embryonic and adult

human skin. The aminopropeptide of type I and type III procollagens were detected on the

same thin fibrils (20-30 nm) but the periphery of larger fibrils (90- 100 nm) contained type

III collagen and type III pN-collagen. It was suggested that the presence of type III pN-

collagen at the surface of thick hybrid collagen fibrils may stop the growth of fibrils in

adult skin and thus regulate their final diameter [19].

Romanic et al. (1991) [257] in an in vitro study demonstrated that pN-collagen III can co-

polymerize with collagen I, but cannot be deposited on previously assembled collagen I fibrils. It was shown that the presence of pN-collagen III can 1) inhibit the rate of collagen

I assembly, 2) decrease the amount of collagen I incorporated into fibrils, and 3) decrease the diameter of fibrils in comparison with fibrils generated under the same conditions from collagen I alone. Fibril diameter progressively decreased with increasing the initial molar ratio of pN-collagen III to collagen I. Therefore, it was concluded that pN-collagen III coats the surface of collagen I fibrils early in the process of fibril assembly and hinder lateral growth of the fibrils. But it does not bind to the growing tips of fibrils, resulting in formation of thin fibrils. Furthermore, Romanic et al. (1992) [258] investigated the co- polymerization of pN-collagen I with collagen and it was showed that, differing from the previous study, the presence of pN-collagen I 1) increased the lag time, 2) decreased the propagation rate, 3) altered fibrils circularity, and 4) increased the concentration of pN- 54

collagen remaining in solution at equilibrium. The differences between the two systems of

copolymerization is shown in Figure 1.18.

Figure 1.18. Schematic of the structures formed from the co-polymerization of pN- collagen III or pN-collagen I with collagen I. pN-collagen III and collagen I co- polymerize to form long, thin, and cylindrical fibrils with 67 nm periodicity (B) as well as thin and non-striated structures (A). Collagen I polymerizes to generate cylindrical fibrils that are shorter and thicker (C) than hybrid fibrils in (B). pN-collagen I co- polymerizes with collagen I to form ribbon-like structures that have a 67 nm pattern (D). pN-collagen I polymerizes by itself to generate large, thin sheet-like structures that are banded (E). From Romanic et al. (1992) [257].

1.3.2.1.5 Fibril Growth Regulation by Collagen Type V

Another collagen type that has shown growth inhibitory effect and retains its N-propeptides in fibrils is collagen type V [20]. Corneal stroma which contains collagen fibrils of uniformly small diameter [259] is relatively rich in type V collagen [260]. Studies of type

I/V interactions in the mature corneal stroma have shown that type I and type V collagen 55

co-assemble into fibrils [261-265]. The triple-helical domain of all the type V collagen molecules are buried (“masked”) within the heterotypic fibrils and the surface of the fibrils are coated only with type I collagen [260]. Type V collagen has a longer helix relative to type I collagen [266, 267] and its collagenase cleavage site is buried within fibrils [268]

(Figure 1.19).

Figure 1.19. Regulation of collagen fibril diameter and growth by type V collagen molecules. Type V amino-terminal domains project onto the fibril surface through the gap zones of the fibril, and when sufficient numbers have accumulated, they block further accretion of collagen monomers thru electrostatic hindrance or by interactions with other charged molecules such as the leucine-rich proteoglycans. Reprinted with permission from Birk (2001) [269] and Marchant et al. (1996) [270]. 56

Throughout embryonic development, the newly synthesized type V collagen molecules are

rapidly converted to the "masked" form [271]. Unmasking can be achieved 1) by treatment

of the tissue sections with dilute acetic acid [260], 2) by perturbing the structure of fibrils at 0 °C [261], or 3) by digestion of corneal sections with the type I-specific collagenase

while sections digested with the type V collagen-degrading enzyme showed no removal of

type V collagen [268]. Therefore, it has been suggested that masking of the type V collagen

molecules is due to fibrillar arrangement. However, NH2-terminal domains of the type V

collagen molecules are exposed at the fibril surface [272] (Figure 1.19). NH2-terminal domain of the native type V molecule is a multi-domain structure that consists of a kink, followed by a short rod, and terminating in a globular domain [273, 274].

Decreasing the levels of type V collagen secreted by corneal fibroblasts in situ resulted in assembly of large-diameter fibrils with a broad size distribution [270]. In vitro fibrillogenesis studies [20, 275] also showed that fibrils produced from only type I collagen were thicker than hybrid fibrils of type I and type V collagen. Fibrils produced from only type V collagen were thin and had no visible striations. However, striated fibrils were formed when small amount of type I collagen were added to the solution of type V collagen.

The greater the proportion of type V collagen included in the reaction mixture, the smaller the diameter of fibril. The amino-terminal domain of the type V collagen molecule was required for this regulatory effect and in its absence little diameter reducing activity was observed.

Models have been proposed [269, 270] for the arrangement of type I/V heterotypic collagen fibrils and the regulation of initial fibril assembly by the NH2-terminal domain of type V 57

collagen on the fibril surface (Figure 1.19). In these models, type I and type V molecules are arranged parallel to one another. The entire triple-helical domain of all the type V collagen molecule are buried within the fibril and type I collagen molecules are present along the fibril surface. Type V amino-terminal domains project onto the fibril surface through the gap zones of the fibril, and when sufficient numbers have accumulated, they block further accretion of collagen monomers and thereby limit growth in diameter. The amino-terminal domain is large and possesses a number of acidic residues and highly charged sulfate groups, and so may affect this block thru steric and/or electrostatic hindrance [276, 277] or by interactions with other charged molecules such as the leucine- rich proteoglycans.

1.3.2.2 Growth Models by Molecular Accretion or Interfibrillar Fusion

Numerous studies have asked the question “How fibrils grow?” and several models have been proposed to explain fibril growth mechanisms. Models in which fibrils grow by molecular accretion or interfibrillar fusion and regulated by mechanisms such as those mentioned in the section 1.3.2.1. In general, two types of models were postulated: diffusion-limited and interface-controlled models. In diffusion-limited growth models, the diffusion of collagen molecules to the fibril surface limits the growth of fibrils. In the interface-controlled models, the rate that molecules accrete onto the fibril surface or the rate that fibrils fuse together, limits the growth of fibrils.

In the latter group, growth will depend on the fibril surface structure and fibrils may grow radially or axially depending on the site of molecular accretion or fibril fusion. Beside the 58

fibril surface-associated growth regulators such as proteoglycans and amino-terminal

domains, fibril polarity can introduce additional structural inhomogeneity on the fibril

surface. Collagen fibrils in vertebrate tissues can be unipolar or bipolar. The unipolar fibrils

have a C-end and an N-end and all molecules are oriented in the same direction. The bipolar

fibrils have a polarity reversal region shown in Figure 1.20.

Figure 1.20. Unipolar and bipolar collagen fibrils from embryonic chick tendon. The molecular switch region of a bipolar fibril is shown in the lower panel. Reprinted with permission from Kadler et al. (2000) [89].

Holmes and Chapman (1979) [26] investigated the growth of native fibrils in acid-soluble collagen by quantitative dark-field electron microscopy. Based on axial mass distribution data it was showed that the number of molecules in an early fibril cross section increased linearly at the end regions of fibrils by 4-5 molecules per D-period at the N-terminal and

8-10 molecules per D-period at the C-terminal. The data from growing fibrils indicated that the rate of molecular accretion on to the fibril surface (∂R/∂t) decreases with increasing 59

the fibril radius (R). It was also mentioned that Haworth and Chapman (1973) showed that

the axial growth rate of the N-terminal was greater than that of the C-terminal.

Kadler et al. (1990) [278] studied longitudinal growth of collagen fibrils in vitro by enzymic cleavage of the partially processed precursor of collagen (pC-collagen). All fibrils initially formed with blunted and pointed ends, rapidly reached a maximal diameter, and then grew from pointed tips in the C- to N-terminal direction without any further increase in the diameter. It was suggested that the pointed tip of the fibril has the highest affinity and defines the critical concentration for fibril assembly. It was shown that the blunt end of the fibril can become a site of intermediate affinity when monomers bind in the reverse orientation and subsequently the blunt end becomes equivalent to a pointed tip for growth.

Scott (1990) [279] swelled rat tail tendon fibrils in acetate buffer, pH 5. Large fibrils disaggregated into two characteristic populations of smaller fibrils of 10-15 nm

(“protofibrils”) and ~25 nm (“subfibrils”) thickness. It was also shown that fully mature rat tail tendon consists of fibrils with diameters ranging from less than 20 nm to more than

600 nm with several fibrils in the process of fusion or separation. Intrafibrillar proteoglycan were observed inside large collagen fibrils in mature tendons. It was suggested that intrafibrillar proteoglycans are originally associated with protofibrils, and larger fibrils are formed by coalescence of smaller fibrils during development.

Silver et al. (1992) [280] developed a helical model for the growth of collagen fibrils based on the principle of nucleation and propagation [281] and which accounts for growth of fibrils from symmetrical parabolic shape tips with linear mass profiles. In this model, a distinctive structural nucleus with spiral or helical conformation forms at each end of a 60

growing fibril. Then, growth of the fibril takes place by propagation of the two structural nuclei. In this model, lateral growth was limited to account for observation of fibrils with the same diameters. At high monomer concentrations, the growth of the fibril was primarily by elongation resulting in tapered tips. At low monomer concentrations, growth occurs mainly by strand initiation resulting in blunter tips.

Holmes et al. (1992) [282] studied the tapered ends of collagen fibrils generated in vitro by enzymic removal of C-terminal propeptides from pC-collagen. Mass measurements by scanning-transmission electron microscopy showed that the fine tips of fibrils exhibited near-identical mass distributions. The mass of the fine tips increased at the rate of ~17 molecules per D-period, regardless of fibril length. The shape of the fine tips was approximately paraboloid of revolution. It was suggested that to maintain this shape throughout growth, accretion cannot be uniform over the fibril tip surface but must decrease as the diameter increases. In this model the end growth of a fibril is favoured at the expense of growth elsewhere (Figure 1.21).

Figure 1.21. Growth of a paraboloidal tip. Dotted curve shows the tip after a short interval of time. The growing tip remains unchanged in shape. From Holmes et al. (1992) [282]. 61

Parkinson et al. (1994 & 1995) [283, 284] used a model based on diffusion limited aggregation to simulate fibril growth. Aggregates were created with elongated morphology, a preference for tip growth, and intrafibrillar fluidity. The results of simulations suggested that fibril morphology is mostly a consequence of the aggregation of rod-like particles from solution, not surface diffusion. However, surface diffusion made the fibrils much more compact suggesting that intrafibrillar collagen-collagen interactions may be critical in the alignment of accreting molecules and to maintain stability in the fibril. In this model, the assembly does not depend on the specific interactions of binding sites in the monomers which is not in agreement with experimental data [285, 286].

Figure 1.22. A model for linear and lateral growth of fibril segments during development. Reprinted with permission from Birk et al. (1995) [287]. 62

Interfibrillar fusion can potentially involve tip-to-tip, tip-to-shaft, and shaft-to-shaft fusion.

Birk et al. (1995) [287] showed that fibril segments, isolated from developing tendons, would laterally associate and fuse into longer and larger diameter collagen fibrils. A model for fibril growth during tendon development was presented which is shown in Figure 1.22.

Based on this model and early in development, fibril-associated decorin or some other macromolecule are involved in the regulation of fibril growth through maintaining interfibrillar spacing and inhibition of segments fusion.

The extracted fibril segments were found to be molecularly bipolar and it was suggested that there is no preferred orientation of fibril segments during their fusion [287]. However,

Kadler et al. (1996) [76] showed that fusion of bipolar fibrils are inconsistent with existing experimental data. Bipolar fibrils with two C-ends or fibrils with multiple switch regions have not been found, either in vivo or in vitro (Figure 1.23). Fusion of unipolar and bipolar fibrils will decrease the unipolar fibril population. Therefore, an enriched bipolar fibril population, unable to fuse further, could determine the limit of fibril growth.

Birk et al. (1996) [207] isolated fibril segments from developing chicken cornea, dermis, and tendon by physical disruption. Fibril segments were asymmetric with long and short tapered ends and centrosymmetric with respect to molecular packing. The results indicated some differences between fibrils of different tissues: 1) Tendon fibrils grew between day

16 and 17 with a rapid increase in length and diameter. Dermis fibrils grew more linearly with respect to time. Corneal fibrils only grew longer with constant diameter. 2) Disruption of fibrils in phosphate-buffered saline resulted in larger diameter corneal fibrils, but tendon and dermis fibrils maintained their diameter. 3) Preserving corneal proteoglycans or the 63

water layer associated with the collagen molecules, maintained the diameter. Based on these observations it was suggested that fibrils grow by linear and lateral association of segments followed by molecular rearrangement. Therefore, generation of longer fibrils in cornea is the result of only end-to-end segments fusion.

Figure 1.23. Possible models of fibril fusion based on fibrils polarity. Arrows indicate molecular polarity within a fibril and pink boxes indicate regions of polarity reversal. From Kadler et al. (1996) [76].

Birk et al. (1997) [288] demonstrated that the fibrils become significantly longer between

14 to 18 days of tendon development. Fibril ends maintained their asymmetry, with no 64

difference observed in their structure. A model of connective tissue development, growth, and remodeling was proposed which is shown in Figure 1.24. In this model, thin fibril intermediates are formed by molecular accretion. Then, longer and larger diameter fibrils are produced by lateral associations of preformed segments. The longer fibrils would have multiple polarity changes which would determine the regions able to associate. Growth would follow by molecular rearrangement to reconstitute cylindrical fibrils. Enzymatic intervention was also considered in this model to degrade poorly cross-linked fibrils in regions of polarity reversal and generate short polar units that could participate in further growth.

Figure 1.24. A model for growth of tendon fibril segments by Birk et al. (1997) [288]. Reprinted with permission. 65

Trotter et al. (1998) [289] studied cross-sectional mass of intact fibrils from sea cucumbers dermis by the scanning transmission electron microscopy. The two ends of fibrils displayed similar mass distributions. There was a linear relationship between cross-sectional mass

and distance from the fibril tip. As the fibril grew in length, the paraboloidal tip became

blunter. The center of each fibril had the maximum cross-sectional mass which

exponentially increased with increasing fibril length. Since fibrils had symmetrical mass

distributions with a single transition zone in the center, therefore fusion as a growth process

was ruled out. Also, the model proposed by Silver et al. [280] which lead to in elongated

fibrils with constant diameters was not consistent with this experimental data. The data

suggested that fibrils grow by coordinated monomer addition at both fibrils centres and

ends, and that the rate of monomer addition at the fibril center should exceed that at the

fibril ends. Trotter et al. (2000) [290] had similar observations on the sea urchin spine

ligament fibrils, despite the greater shape variability of the sea cucumber fibrils. It was

suggested that the fibril tips produce independent axial growth, while lateral growth takes

place through a surface nucleation and propagation mechanism.

Holmes et al. (1998) [286] showed that C and N-terminal tips of collagen fibrils in

developing tendon were parabolic in shape and had linear axial mass distributions. Fibrils

showed an abrupt stop in lateral growth at multiples of five D-periods from the end of the

fibrils. In day 12, embryonic fibrils grew in length at constant diameter and tip shape. In

day 18, embryonic fibrils showed coarser tips and grew by lateral accretion onto the central

shafts (away from the growing tips). It was suggested that fibril assembly is an end-

regulated growth and relies on surface structural features which repeat every 5 D-periods. 66

Nurminskaya and Birk (1998) [291] presented a model of collagen fibril growth during chicken tendon development which is shown in Figure 1.25. It was hypothesized that fibrils grow in a three stage process regulated by stabilization and destabilization of fibrils surface. In the first stage of development, fibril segments are deposited in the ECM and stabilized through interaction with other ECM macromolecules. In the second stage, fibrils are destabilized and then grow in diameter and length by lateral fusion. In the last stage, mature fibrils are stabilized by a new set of fibril-associated molecules.

Figure 1.25. Model of fibril growth regulation during tendon development by Nurminskaya and Birk (1998) [291]. Reprinted with permission. 67

Ezura et al. (2000) [222] proposed a multi-step model for regulation of fibrillogenesis by lumican and fibromodulin in developing mouse tendons (Figure 1.26). In the first step,

fibril intermediates form by molecular accretion within extracellular compartments, under

cellular control, and regulated by type III collagen, decorin, lumican, and fibromodulin.

Fibril intermediates (~64 nm) are stabilized through their interactions with these leucine-

rich repeat proteoglycans. As fibromodulin displaces lumican on the fibril surface, fibril

intermediates generate more mature fibrils in a multi-step fusion growth process as shown

in the Figure 1.26.

Figure 1.26. A multi-step model for regulation of fibrillogenesis by lumican and fibromodulin in the mouse tendon. Fibril intermediates form by molecular accretion and stabilized through their interactions with leucine-rich repeat proteoglycans. The change in composition of the matrix proteoglycans leads to a multi-step fusion growth process. Reprinted with permission from Ezura et al. (2000) [222]. 68

Graham et al. (2000) [292] showed that fibroblasts synthesize two types of “early fibrils”

(~1 µm long): unipolar fibrils (with C and N ends) and bipolar fibrils (with two N-ends).

These early fibrils would generate longer fibrils by tip-to-tip fusion which requires the C-

end of a unipolar fibril. It was shown that fibril tips have less surface bound proteoglycans

compared to the fibril shaft. Also, removing proteoglycans of entire fibrils allowed lateral

association of fibrils to form fibril aggregates with distinguishable individual fibrils.

Therefore, it was suggested that fibrils fusion is regulated by collagen-proteoglycan interactions. Furthermore, Kadler et al. (2000) [89] showed that fibril fusion is not restricted to tip-to-tip fusion but can include tip-to-shaft in older tissues fibrils to generate branching networks of fibrils.

Cisneros et al. (2006) [293] observed self-assembly of acid-soluble collagen molecules into fibrils using time-lapse atomic force microscopy (AFM). Fibril growth initiated with fibrillar structures of ~1.5 nm thickness (about one collagen molecule thick). Fibrils grew with constant lateral and longitudinal growth rates into a final thickness of ~3 nm. Fibrils fused with each other and grew further. It was suggested that fibril self-assembly is a two- step process: 1) molecular assembly where molecules associated with each other or with fibrils, and 2) molecular rearrangement into microfibrils which form the building-blocks of fibrils.

Holmes et al. (2010) [294] isolated fibrils from avian embryonic tendon and incubated them in collagen solution. It was showed that fractured ends of fibrils could further grow in opposite axial direction by molecular accretion. A model of surface nucleation and 69

growth was proposed in which the ends of the broken fibrils act as nucleation sites and grow bidirectionally (Figure 1.27).

Figure 1.27. Growth of fibril ends. a) extracted fibril from 13-day chick embryonic tendon. Growth of fibril end after 30 min (b) and after 2 h (c) incubation in collagen solution. d) A schematic of nucleation and propagation growth model. From Holmes et al. (2010) [294].

Kalson et al. (2011) [295] investigated the effect of slow stretching on a cell culture tendon- like construct. The results showed an increase in fibril diameter and volume fraction. 70

Furthermore, it was shown that the increase in fibril volume fraction was due to an increase in fibril length and/or nucleation of new fibrils. It was noted that inter-fibrillar fusion alone could not explain the increase in fibril volume fraction.

Kalson et al. (2015) [13] presented a growth model based on 3D-electron microscopy of mouse tendon at three time points: in the embryo, at birth, and six weeks later (Figure 1.28).

Two stages in tendon development were identified. During the first stage (the embryonic growth stage), fibril number, diameter, and length increase by fibril nucleation and axial growth. During the second stage (the postnatal growth), fibril number remains constant but fibril diameter and length continue to grow likely by interface-limited molecular accretion.

Figure 1.28. Model of fibril nucleation and growth during tendon development by Kalson et al. (2015) [13]. 71

1.3.3 Role of Mechanics in Fibrillogenesis, Growth, and Remodeling of Collagenous Tissue

In most of the models and studies that were reviewed above, the role of local tissue micro-

mechanical environment, namely stresses and strains acting on collagen fibrils, have been

overlooked. However, there is sufficient evidence for the role that mechanics has to play

in regulating the formation, growth, and remodeling of collagenous structures in tissue,

fiber, and fibril levels. Mechanical stress can drastically enhance mechanical properties of

collagenous tissue in the path of load [50, 296, 297]. Lack of tension leads to disassociation

of existing collagen fibers [42] and formation of immature fibrils [298] with no preferred orientation [299].

At tissue level, the biomechanics of musculoskeletal system and connection of bone and connective tissue such as tendon and have been studied for centuries. Since

Wolff described the biological effect of applied load on bone, numerus studies have shown that lack of mechanical stresses due to immobilization leads to bone atrophy [300-304]. It is known that physical stresses (produced by muscle tension) are critical for bone remodeling [305-311], and continuous growth can be stimulated by the load environment

[312-314]. This growth-generated stresses and strains [315] at bone tissue may modulate growth and remodeling of surrounding simply due to the physical connection and necessity of growth harmony.

Direct studies of soft tissues also support the role of mechanics. Studies have shown that growth of load-bearing collagenous tissue of eye is directed by mechanical signals [316-

319]. In case of tendon, physical training results in an increased turnover of collagen in 72

local connective tissue [29-31]. On the other hand, immobilization decreases the tensile

strength, elastic stiffness, and total weight of the tendon [320]. However, part of these

changes might be due to decrease of proteoglycan and water content after immobility [321].

Still, occurrence of large fibrils in those tissues which are subjected to the highest tensile

loads like tendon versus occurrence of small fibrils in cornea is thought-provoking.

At fibrillar level, lack of stress during rehabilitation and healing process results in

haphazard and weaker fibrils [322]. However, physical training and immobilization

experiments have not been systematic and have resulted in less clear conclusions about

fibril number and diameter: Physical loading has resulted in an increased mean fibril

number and diameter [323]. Immobilization, on the other hand, has been shown to result

in a looser packing of the fibrils, decreased number of larger fibrils, and lower average

fibril diameter [324]. However, exposure to intense loading may also result in injury and

split of larger fibrils and therefore increase in the number of small fibrils [325, 326].

Conversely, immobilization has been shown to increase the number of larger fibrils and

decrease the number of smaller fibrils [327]. This was suggested to be due to a decrease in

collagen synthesis (resulting in fewer new and small fibrils) and degradation (resulting in

persistence of current fibrils and possibly fusion of smaller fibrils). Immobilization has

been also resulted in longitudinal splitting of fibrils (and therefore thinner fibrils) and

abnormal fibril orientation [328]. A more controlled experimental condition to prevent tissue injury due to excessive loading and to prevent growth due to increased collagen synthesis would be necessary to understand the direct effect of physical loading on fibril growth. Indeed, in a more relatively controlled experimental condition, slow stretching of 73

a cell culture tendon-like construct increased fibril diameter, fibril packing volume, and

mechanical stiffness [295].

Mechanics also play a significant role in cell-collagenous matrix interactions. Mechanical

stimulation can alter cell differentiation [329-331], growth [332, 333], migration [334-

336], orientation [337, 338], and collagen synthesis [339]. Cells cultured on elastic

substrates orient in the direction of tensile strain [340] and move toward the stiffer region

[341-343] with minimum strain energy [344]. In return, collagenous tissue remodeling is driven by cell-level mechanical stresses [345-347]. Fibroblasts are the primary cell type in collagenous matrix. They play an important role during fibril forming and remodeling conditions such as development [34, 35, 40] and wound healing [348-355]. In vitro, they

can exert mechanical forces on collagen matrix [168, 298, 356] to translocate [357], align

[33-36, 38, 334, 358, 359], and stabilize fibrils [360] and form tissue like structures [33].

The mechanical cues may not be necessarily provided by cells, instead may be purely under

local physical environment. Warren (1981) [361] showed that collagen fibrils of

regenerating frog tadpole tail orient in the tail axis prior to elongation of myogenic cells,

providing a directional developmental cue for muscle cells.

Overall, various studies are adding to our understanding of the functional adaptation of

collagenous tissue to its mechanosensitive environment. However, the experimental

knowledge of the effects of physical loading on individual fibrils is scarce. Therefore, the

question, how mechanics results in formation, growth, and remodeling of collagen fibrils,

remains perplexing. That is to say, the detailed mechanisms by which the actual collagen

molecules (and/or microfibrils) are added to or subtracted from the resident collagen fibrils 74

are not known. It is likely, however, that the mechanism will operate independently of the

cell surface or at some nominal distance from it [40], guided by spatial cues [41] and mechanical forces [42].

The purpose of this dissertation was to simply ask the question, does fibril strain promote

the molecular assembly of collagen? To investigate this question, individual collagen

fibrils were manipulated and stretched between two microneedles. The stretched fibrils were exposed to subthreshold concentration of fluorescently labeled collagen molecules to quantify molecular association onto loaded fibrils (chapter 4). To achieve this, it was necessary to 1) fluorescently label individual collagen monomers while minimally disturbing their fibril forming functionality (chapter 2) and 2) accurately measure the diameter of tested fibrils (chapter 3).

75

2 DYNAMIC TRACKING OF FLUORESCENTLY LABELED TYPE I COLLAGEN MOLECULES; DIRECT QUANTIFICATION OF MOLECULAR ASSOCIATION WITH NATIVE FIBRILS

2.1 Introduction

While type I collagen is the most abundant ECM protein and there have been numerous

studies on collagen biosynthesis and its regulatory stages, the fibrillogenesis, growth, and

remodeling mechanisms are less clear. To explore the possible mechanisms that govern the

fibrillogenesis mechanism and homeostasis at the fibril surface, it is important to be able

to directly observe the motion of individual collagen monomers entering and exiting the

fibril. Though collagen fibrils themselves are readily imaged via light and electron

microscopy, dynamic tracking of their surface interaction with individual collagen

monomers has not been performed.

The diameter of collagen molecules is generally far below the Abbe diffraction limit of

light and thus undetectable via light microscopy, and electron microscopy requires a

dehydrated, static sample [362]. Molecular labeling is a common strategy to enable light

microscopy to detect structures below the wavelength of light. However, these methods

may also require fixation or post processing of the sample [363, 364], alter protein structure 76

[365, 366] and synthesis [367, 368], detect other intracellular or ECM components [369],

or lack detailed characterization of collagen after labeling [370, 371]. Therefore,

examination and quantification of collagen network dynamics with molecular scale

resolution remains an important and unmet challenge.

In this section, common methods to detect or label collagen are reviewed and their

functionality to study the dynamic fibrillogenesis process and interaction of individual

collagen monomers with fibrils are discussed.

2.1.1 Radioactive and Nonradioactive Isotopically-Labeled Amino Acids

Labeling a protein during synthesis by incorporation of a labeled precursor is referred to

intrinsic (metabolic or biosynthetic) labeling. Radiolabeled amino acids like [methyl-11C]-

L-methionine [372-376], L-[3H]methionine [377], [3H]-L-proline [378], cis-4-[18F]fluoro-

L-proline [379-382], and trans-4-[18F]fluoro-L-proline [383] as well as azide-L-proline

[384, 385] have been used as collagen markers usually in clinical cancer imaging using positron emission tomography. Majority of these studies target proline for labeling collagen. The amino acid proline is frequent in collagen polypeptide chain sequence: Gly-

X-Y, where X is often an L-proline residue and Y is often a 4(R)-hydroxy-L-proline residue. This makes proline a candidate marker for collagen synthesis and detection.

These radiolabeled amino acids have been used widely to study collagen synthesis [386].

However, it has been known that slight abnormalities in collagen structure can change

thermal stability [387-389] and folding rate [390, 391] of its triple helical structure and 77

cause variety of connective tissue diseases [392-394]. Indeed, it has been shown that targeting a frequent amino acid like proline can affect collagen synthesis and fibrillogenesis. For instance, Takeuchi and Prockop [395] showed that different proline analogues decreased incorporation of [14C]proline into procollagen. They also showed

abnormal collagen containing L-azetide-2-carboxylic acid or cis-4-fluoro-L-proline are not extruded by cartilage cells due to altered kinetics for the enzymatic hydroxylation and decreased amount of hydroxyproline and hydroxylysine [365]. It has been also shown that incorporation of cis-4-hydroxy-L-proline interfere with the triple helical conformation so that the hydroxyproline is not resistant to pepsin [366].

Generally, labeling collagen using the isotopically-labeled amino acids is suitable for endogenous labeling of collagen monomers and requires additional detection imaging techniques such as positron emission tomography. While it is possible to extract the endogenously labeled collagen, our goal was to label tissue extracted collagen monomers exogenously for the purpose of simplicity. In conclusion from the above studies, a fluorescent probe that satisfies our requirements such as accurate quantitative measurements of labeled collagen molecules fibrillogenesis, should minimally disturb the collagen triple-helix with least possible number of labels.

2.1.2 Non-collagen Based Probes

Several studies have taken advantage of non-collagen molecules that have collagen binding

domains [396] to label collagen. Krahn et al. (2006) [370] fluorescently labeled collagen

binding protein domains present in bacterial adhesion proteins (CNA35 [397]) and 78

integrins (GST-α1I [398]) using either amine-reactive succinimide or cysteine-reactive

maleimide dyes. This probe has high specific affinity for human collagen type I and can be

used for imaging of fibril formation in long term tissue cultures. However, this labeling method adds more than 60 kDa to each collagen monomer which can drastically change its behaviour. This probe has been expanded by Aper et al. (2014) [399] to span the visible

spectrum.

Chilakamarthi et al. (2014) [400] presented a method for coupling of 5-(4-carboxyphenyl)-

10, 15, 20-triphenyl porphyrin (C-TPP) to the N-terminal of collagelin (a specific collagen binding peptide [401]) for imaging of collagen in live tissues. Their probe bound strongly and specifically to collagen type I in fibrotic liver tissue but not to normal liver tissue. This in vivo collagen detection probe is suitable for detection of liver fibrosis.

Biela et al. (2013) [402] used a low molecular weight fluorescent probe in which fluorescein (332.31 g/mol) conjugated to physostigmine (275.346 g/mol). This fluorescent probe can penetrate into tissue and non-covalently bind to ECM fibers. A major drawback of this method is its affinity to other ECM proteins such as . Also it was only used to label fibrillar collagen and its affinity was not tested for monomeric collagen.

Interestingly they performed a fluorescence recovery after photobleaching (FRAP) experiment and observed fluorescent recovery in the photobleached areas while fluorescent intensity was decreased in other regions. They explained the observation was due to weak chemical binding mechanism. However, this could be due to dynamic interactions between labeled collagen monomers and fibers. 79

In summary, non-collagen based labeling methods provide a great tool to detect fibrillar collagen. However, they are not optimized to track individual collagen monomers. These methods require addition of a collagen binding molecule and a fluorescent dye to the 300 kDa collagen molecule. While, it has been shown that only the addition of a collagen binding peptide can delay collagen fibrillogenesis [403]. For the purpose of our study, we need to preserve collagen monomer functionality by addition of only a small fluorescent tag.

2.1.3 Endogenous Labeling with GFP

Several studies have endogenously labeled the α1 or α2 chain using the green fluorescent protein (GFP). Krempen et al. (1999) [404] developed gene constructs containing the Col

1 a1 promoter driving the GFP reporter gene and tested their possible functions in transfection experiments and transgenic mice. Grant et al. (2000) [405] generated GFP-

Col2a1 producing mice for studying chondrocyte differentiation and chondrogenesis during skeletal development. Kalajzik et al. (2002) [406] produced GFP-Col1a1 expressing transgenic mice to study different levels of osteoblastic differentiation. Yata et al. (2003)

[407] generated a transgenic mouse line expressing GFP-Col1a1 to investigate the molecular mechanisms controlling increased collagen expression during fibrosis.

Yamaguchi et al. (2018) [408] generated a new GFP-Col1 transgenic mouse line based on

Yata et al. (2003) [407] procedure. They histologically confirmed the specificity of GFP expression in the mouse line and explored the possibility of type I collagen-producing cells other than fibroblasts. Stephens and Pepperkok (2002) [409] cloned human procollagen 80

type I α1 labeled with GFP to investigate ER-to-Golgi transport of procollagen. The GFP- procollagen behaved similarly to endogenous procollagen and was transported from the

ER to the Golgi complex in transport complexes. While these authors observed no differences in general development, health, and fertility between transgenic and wild-type

mice, the collagen fibrillogenesis process was not the subject of their studies and therefore

the effect of labeling on fibrillogenesis was not considered.

Most recently, Lu et al. (2018) [175] labeled collagen monomers by replacing the α2(I)-

procollagen N-terminal propeptide with GFPtpz or mCherry. The label was added to the

N-terminus to avoid the C-terminus for two reasons. First, C-terminus is critical for

nucleation of collagen triple helix. Second, it has been shown that patients with Ehlers

Danlos Syndrome type VII, which results in loss of the N-proteinase cleavage site in α2(I)

chain, can still make collagen triple helix and have no sign of bone disease [410]. This

imaging probe was transfected into osteoblast-like cells and fibronectin-null mouse

embryonic fibroblasts and used to image type I collagen assembly dynamics. Labeled

monomers were able to incorporate into extracellular fibril networks with normal D-

banding patterns. This labeling method has been extended by Morris et al. (2018) [411] to

generate an epidermal-specific GFP-collagen I transgenic zebrafish line to study the

dynamic nature of collagen fibril deposition and repair after wounding. While this labeling

method has provided a powerful tool for imaging of dynamic collagen fibrillogenesis in

live cultured cells, it still lacks some criteria to be used for kinetic studies of fibril

formation, growth, and remodeling. In fact, the GFP has a comparatively large molecular 81

weight (27 kDa) and can potentially affect collagen monomer-monomer or monomer-fibril

reactions.

2.1.4 Exogenous Labeling

Harris et al. (1984) [36] labeled collagen exogenously using fluorescein isothiocyanate

(FITC) which has a small molecular weight (389.382 g/mol). The collagen solution was

dialyzed against 0.25 M carbonate buffer (pH 9.5) and diluted to 1.6 mg/ml. Then FITC

was added progressively to a final concentration of 0.08 mg per mg of collagen and mixed

for 24 h at 4°C. A Sephadex G-25 column and 0.05 M phosphate containing 0.25 M NaCl

was used to separate unbound FITC. Labeled collagen was then dialysed against dilute

acetic acid. However, they did not quantify the effect of this labeling process on collagen

fibrillogenesis.

Stopak et al. (1985) [40] labeled collagen solutions from rat tail tendons with FITC. The collagen solution was dialyzed in 0.05 M phosphate containing 0.25 M NaCl. FITC was added to the collagen solution at a concentration of 0.08 mg per mg of collagen and mixed overnight. They also used gel filtration to remove the unbound FITC. They reported a degree of labeling of 1-3.5 molecule of FITC per 1 molecule of collagen. They added these labeled monomers to cell culture and observed no apparent difference in cell behavior or morphology compared to unlabeled collagen.

Cabral et al. (2003) [387] labeled procollagen with Cy2 (664.7 g/mol) and Cy5 (667.5 g/mol) fluorescent dyes. Fluorophores were added to 0.2 mg/mL collagen in 0.1 M sodium 82

carbonate and 0.5 M NaCl (pH 9.3) and stirred for 30 min at room temperature and stored

at -80 °C. Makareeva et al. (2006) [412] used the same method to label pepsin treated collagen. Han et al. (2008) [413] also used the similar method to label collagen with Cy5,

Alexa Fluor 488, and Alexa Fluor 568. They stated that labeling with Cy5 dyes altered the

fibrillogenesis kinetics curve and induced formation of nonfibrillar aggregates even at low

labeling fractions. On the other hand, they observed normal turbidity curves, critical

fibrillogenesis concentrations, and fibril morphology when collagen was labeled with less

than one Alexa Fluor dye per four to five collagen monomer. However, the data was not

shown. Han et al. (2010) [414] used the same labeling method and removed the unreacted

dye by collagen precipitation in 0.5 M acetic acid containing 0.9 M NaCl, dialysis, or size-

exclusion chromatography.

Johnson et al. (2003) [171] labeled collagen exogenously using FITC. First, collagen

monomers were dialyzed (8000 MWCO) against 100 mM sodium borate and 1M sodium chloride (pH 9.3) for 24 h at 4 °C. Then 20 mg/mL FITC in DMSO (approximately 56:1 molar ratio of dye to collagen) was added to collagen and gently stirred for 3 h at room temperature. Labeled monomers were dialyzed against 0.02 M acetic acid for 24 h at 4 °C to remove any unbound dye and to resolubilize the labeled collagen. Then labeled monomers were stored at 4°C. They added exogenous FITC-collagen (10-100 µg/mL) to the cultured smooth muscle cells and showed collagen assembly. However, their labeled monomers did not assemble under 100 µg/mL concentration in the absence of cells. Beside some TEM images that showed striated collagen fibrils (in low resolution), the morphology of fibrils and fibrillogenesis kinetics were not investigated. 83

Li et al. (2003) [170] labeled monomeric collagen to track fibril formation in both cellular

and noncellular environments. To label collagen, first they dialyzed tissue extracted and

soluble collagen against borate-buffered saline (170 mmol/L boric acid, 170 mmol/L sodium tetraborate, 75 mmol/L NaCl, pH 9.3). Then collagen was transferred to a solution of borate-buffered saline containing 30 mg/ml of either FITC or Texas Red and mixed in

the dark at 4 °C for 6 hours. Then unbound fluorophores were removed by dialysis against

0.1% acetic acid at 4 °C for 4 days. The concentration of labeled collagen was measured spectrophotometrically by measuring absorption at 280 nm (for collagen) and either 493 nm (for FITC) or 595 nm (for Texas Red) wavelengths. They showed that cultured human vascular smooth muscle cells in ascorbate-free medium (to prevent endogenous collagen production) used the added exogenous labeled monomers to form cross-banded fibrils with

67 nm periodicity. However, they did not investigate how their labeling process affected fibrillogenesis kinetics.

Meshel et al. (2005) [415] labeled lyophilized rat-tail collagen using Cy5 fluorophore. The labeling reaction was proceeded at pH 4 to prevent fibrillogenesis of collagen monomer into fibrils. Labeled collagen was then dialysed overnight in 0.2% acetic acid to remove unbound fluorophore. Labeling at lower pH prevented fibrillogenesis but also reduced the efficiency of labeling (10 times more fluorophore was used compared with the recommended amount by the manufacturer’s instructions).

Nevertheless, Yang et al. (2009) [416] using confocal reflectance and fluorescence microscopy studied fibril and network structure of FITC-labeled collagen gel. They used

FITC-conjugate type I collagen from Sigma Aldrich. They showed that even at a low 84

labeling density (3% FITC-labeled collagen) the labeling process inhibited lateral fibril bundling and induced changes in gel structure.

Overall, exogenous labeling of collagen monomers with a small molecular weight fluorophore has provided the most direct method and has shown the most promising results.

But there is still a need to fully characterize the labeled monomers to ensure uninterrupted fibrillogenesis kinetics and fibril morphology.

2.2 Objectives and Approaches

Our goal was to fluorescently label and track collagen monomers, while minimizing the effect of labeling on the kinetics of collagen fibrillogenesis and on the assembled fibril morphology. Proteins, in general, may be fluorescently tagged by the N-terminal α-amine, the C-terminal α-carboxyl, and the reactive groups at their side chains (e.g. primary amines, carboxylates, and disulfides) [417].

Figure 2.1. Amino acid lysine. The free amine group in its side chain is a potential binding side for amine reactive fluorophores. 85

Lysine (Figure 2.1) is a basic, charged, aliphatic amino acid. It contains an ε-amino group,

an α-amino group, and an α-carboxylic acid group. Lysine is an essential amino acid and

is required for formation of collagen. There are 38 positively charged and hydrophilic

lysines as potential binding site on each α-chain [418]. Collagen alpha-1(I) chain (Bovine) is shown in Figure 2.2.

Figure 2.2. Collagen alpha-1(I) chain amino acid sequence (bovine). The triple-helical region is highlighted and lysine is shown with letter K. There are 36 lysines in the triple- helical region and 1 lysine in each terminal telopeptides. Signal peptide (1-22), N- terminal propeptide (23-161), N-terminal telopeptides (162-177), triple-helical region (178-1191), C-terminal telopeptides (1192-1215), C-terminal propeptide (1218-1463). Reprinted with permission from UniProt: the universal protein knowledgebase [418]. 86

Abundance of lysine in collagen triple-helix, makes it a great target for fluorescent labeling.

It has been also shown that chemical combination of proteins with amine-reactive

fluorophores largely occurs through the amino group of the lysine [419] which is positively charged and located at protein surfaces [420, 421].

Another potential target for labeling collagen with an amine reactive dye is the N-terminal

α-amine. However, N-terminal might have major biological role in fibrillogenesis [229]. It

has been shown that pepsin treatment which partially degrades telopeptides, inhibits

irreversible covalent cross-linking between collagen monomers [422, 423] and alters the fibrillogenesis kinetics [422, 424]. Yet, labeling the N-terminal has not affected the triple- helix structure and their ability to form fibrils [175, 425].

Figure 2.3. A) Alexa Fluor 488 carboxylic acid, TFP ester (Molecular formula:

C39H44F4N4O11S2, Molecular weight: 884.91). B) Fluorescence excitation (—) and emission (- - -) of AF488 (Ex/Em: 494/519 nm). Image provided by supplier, Thermo Scientific. 87

In this investigation, we used Alexa Fluor 488 carboxylic acid, TFP ester fluorescent dye

(AF488) to label the free amine at the N-terminus and/or the ε-amine group of lysine throughout collagen molecule. The AF488 molecule has a relatively small molecular

weight of 884.91 g/mol compare with the collagen molecule (300 kDa). The AF488

molecular structure is shown in Figure 2.3.A and its fluorescence excitation and emission

are shown in Figure 2.3.B (Ex/Em: 494/519 nm).

Figure 2.4. A) Reaction of Alexa Fluor 488 with a free amine group. From www.thermofisher.com and the molecular probes handbook [426].

AF488 can bind to a free amine group on the collagen molecule. The reaction of AF488

with the free amine group is shown in Figure 2.4. Based on the manufacturer information,

AF488 is the most fluorescent and the most photostable available fluorescent dye (Figure

2.5.A). It is also suitable for multilabeling of each collagen molecule and allows for more

fluorophores to be attached to each collagen monomer before self-quenching becomes

apparent (Figure 2.5.B).

We targeted a labeling ratio of ~2 fluorophores per collagen molecule to minimize

disrupting the functionality of collagen, while still permitting us to determine the

orientation of the tagged molecules on the fibril surface. Such a tool has the potential to 88

not only track individual molecular associations in a complex biological system but also the directionality of the molecule with respect to the fibril axis. This will permit us to determine if the associations are functional or non-specific.

Figure 2.5. A) Alexa Fluor 488 photostability in comparison with Oregon Green 488 and Fluorescein. Samples were scanned 10 times with a 25 mW laser power and approximately 5 minutes each time. B) Relative fluorescence of Alexa Fluor 488 and FITC at multiple mole of dye per mole of protein. From www.thermofisher.com and the molecular probes handbook [426].

2.3 Experimental Methods

2.3.1 Collagen Labeling

Monomeric collagen was labeled using standard methods for fluorescently labeling antibodies [419]. Type I bovine collagen solution (5026-50ML, TeloCol, Advanced

BioMatrix) was diluted to 1 mg/ml using 10 mM hydrochloric acid (SA56-1, Fisher

Scientific), and then mixed 1:1 with 0.2 M sodium carbonate–bicarbonate buffer (24095, 89

Polysciences) containing 1 M sodium chloride (S671-3, Fisher Scientific) [40, 412]. The

labeling protocol was carried out at pH 7.5, 8.0, and 8.5 to probe the effect of pH on the

DOL and fibrillogenesis of labeled monomers. Lysine residues are most efficiently labeled

at pH values of 8.5-9.5; however, the lower pH range was selected to increase the

probability of attaching one fluorophore to the N-terminal [417] and therefore increase the average distance between fluorophores on each collagen molecule (Figure 2.6) for determining monomer orientation.

Figure 2.6. Schematic of collagen molecule labeled with 2 fluorophores. There are 38 positively charged and hydrophilic lysines and one N-terminal α-amine as potential binding site on each α-chain for the label [418]. Dimensions are not to scale.

AF488 (A37570, Alexa Fluor 488 TFP ester, Invitrogen) was dissolved in dimethyl sulfoxide (D12345, Life Technology) to a concentration of 0.5 mg/mL and added to collagen solution in 3, 6, 9, and 15x molar excess to achieve 1-10 fluorophores per collagen molecule. The reaction mixtures were stirred for 3 hours at room temperature.

2.3.2 Purifying the Labeled Collagen

Labeled collagen was mixed 1:1 with 1 M acetic acid (AC124040025, Acros Organics)

containing 1.3 M sodium chloride to precipitate the collagen. Next, the sample was 90

centrifuged for 1 hour at 10000×g, the supernatant was removed, and the pellet was dissolved in 10 mM hydrochloric acid for 12 hours at 4 °C. The solution of labeled monomers was injected into a 3.5 kDa MWCO dialysis cassette (66110, Thermo Scientific) and dialyzed for 3 days against 100 times the volume of 10 mM hydrochloric acid at 4 °C to remove any unbound dye from the protein solution. To ensure the efficiency of the dialysis process, AF488 (without collagen) was injected into 3.5 kDa MWCO dialysis cassettes. No free fluorophore was detected after 3 days in the dialysis cassettes.

2.3.3 Determining Collagen Concentration and Degree of Labeling

The collagen concentration of labeled monomers was determined using the DC Protein

Assay (500-0114, Bio-Rad Laboratories). Standard curves were produced using known concentrations of TeloCol collagen solution. The volume of samples was adjusted to make the procedure sensitive to our low working concentrations.

AF488 concentration was calculated based on the Beer-Lambert law:

A = lc (2.1)

ε where ‘A’ is the absorbance, ‘ε’ is the molar extinction coefficient, ‘l’ is the optical path length, and ‘c’ is the concentration. The labeled monomers’ absorbance was measured at

494 nm in a 1 cm pathlength cuvette. Molar extinction coefficient of AF488 was considered

71,000 based on the manufacturer data sheet. We used a standard 1 cm pathlength cuvette to improve measurement accuracy and repeatability. The cuvette was cleaned thoroughly with de-ionized water and filled with extra care to avoid formation of any bubbles. 91

Absorbance of 10 mM HCl in the 1 cm pathlength cuvette was also measured and

subtracted from data to calculate the absolute labeled monomers’ absorbance. The final

degree of labeling (DOL) was calculated as moles of dye per moles of collagen.

2.3.4 Spectrophotometry

To quantify the fibrillogenesis kinetics of each batch of labeled collagen solution, a plate

reader spectrophotometer (PowerWave XS, BioTek Instruments) was used to measure the

turbidity curve associated with assembly [87]. This was compared against unlabeled solution at the same concentration to determine the impact of labeling. Collagen solutions were prepared at 50 µg/mL in phosphate buffered saline (PBS), pH 7.3, and room temperature. The spectrophotometer temperature was set at 37 °C. 100 µL samples were held in a 96 well plate and absorbance of the collagen solutions were read every 30 seconds at 313 nm. The lag time and plateau time were calculated respectively as the time when the turbidity curve reached 10% and 90% of the maximum absorbance.

2.3.5 Scleral Fibril Extraction

Bovine eyes, 1-10 day old, were purchased from Research 87, Inc. and disinfected

immediately using Polyvinylpyrrolidone-iodine (PVP-I). PVP-I is an iodine-containing antiseptic compound with a broad spectrum of activity against bacteria, fungi, and viruses

[427-429]. Eyes were washed with sterile-filtered PBS and immersed in 0.1% PVP-I for

30 s [430]. It has been shown that high concentration and long exposure of PVP-I can cause 92

significant damage to tissue [431, 432]. Therefore, higher concentration and exposure time

were prevented.

Next, native collagen fibrils were trypsin extracted from the bovine sclera based on the

method of Liu et al. (2016) [433]. Briefly, the scleral tissue was isolated from the cornea,

optic nerve, muscle, and surrounding . The vitreous humor was removed and

the retina, tapetum lucidum, and choroid layers were debrided from the sclera, and the

sclera was then cut into smaller pieces (approximately 1 cm by 2 cm). Figure 2.7 shows

the sclera preparation and cleaning steps.

Figure 2.7. A) Bovine eye dissection. B) Cleaning the sclera. C) Cleaned sclera was cut into smaller pieces.

Some shallow cuts were made on each piece to increase surface area for fibril extraction.

Each piece of sclera was washed with sterile-filtered PBS, immersed in 0.1% PVP-I for 30

s, washed again with sterile-filtered PBS, and then placed into 20 mL of pH 7.5-8.0

extraction medium containing: >1000 BAEE unit/mL trypsin (9002-07-7, Sigma-Aldrich),

4 mM ethylenediaminetraacetic acid (60-00-4, Sigma-Aldrich), 0.1 M tris-HCl (1185-53- 93

1, Sigma-Aldrich), and 0.05% sodium azide (26628-22-8, Sigma-Aldrich). Samples were shaken for ~5 minutes until the sclera swelled and some fibrils were removed. Fibril suspensions were centrifuged at 5000×g for 30 minutes. The Supernatant was removed, and the fibril pellet was resuspended and stored in PBS containing 1% antibiotic/antimycotic (mixture of 10 mg/mL streptomycin and 25 mL/mL amphotericin

B) at 4 °C. Figure 2.8 demonstrates a schematic of fibril extraction process.

Figure 2.8. Schematic of fibril extraction process. Scleral tissue was dissected from 1-10 day old bovine eyeballs. Tissue was cut into 3-4 pieces and some shallow cuts were made on each piece. Then they were placed in 20 mL of the trypsin extraction solution. Samples were shaken for ~5 minutes until the sclera swelled and some fibrils were extracted. Pieces of sclera were then removed from solution and fibril suspension was centrifuged at 5000×g for 30 minutes. The Supernatant was removed, and the fibril pellet was resuspended and stored in PBS at 4 °C.

2.3.6 Proteoglycan Quantification

Fibril suspensions in trypsin extraction medium were centrifuged for 30 minutes at

14,500×g and the pellet was suspended overnight in 300 μL of 10 mM HCl at 4 ℃. 50 μL 94

of 8 M GuHCl solution (Acros Organics, 50-01-1) was added to each sample and incubated

overnight at 4 ℃. These sample preparation steps are shown in Figure 2.9.

Figure 2.9. Schematic of sample preparation steps for proteoglycan analysis. 300 μL of fibril suspension was used for each sample. Samples were centrifuged for 30 min at 14,500x g to isolate the supernatant and pellet. Supernatant was stored until 50 μL 8M GuHCl was added 24 hours prior to proteoglycan staining. The pellet was resuspended for 24 hours in 300 μL of 10mM HCl and then 50 μL 8M GuHCl was added 24 hours prior to proteoglycan staining.

Then, 50 μL of SAT solution (used to dilute alcian blue solution) containing 30 µL of 18

M sulfuric acid (Acros Organics, 7664-93-9), 75 µL of triton x-100 (Sigma-Aldrich, 9002-

93-1), and 10 mL water was added to each sample, vortexed, and incubated at room temperature for 5 min. Alcian blue solution containing 20 µL of 18 M sulfuric acid (Acros

Organics, 7664-93-9), 1 mL of 8 M GuHCl, 10 mL of 1% alcian blue in 3% acetic acid

(Sigma-Aldrich, B8438), and 10 mL water was prepared. 750 μL of alcian blue working solution (5 SAT: 9 water: 1 alcian blue solution) was added to each sample and incubated for 1 hour at 4 ℃. 95

Figure 2.10. Schematic of proteoglycans analysis process.

Samples were centrifuged for 15 min at 12,000×g and supernatant was discarded. Pellets were suspended in 500 μL washing solution containing 8mL dimethyl sulfoxide (Fisher

Scientific, 67-68-5), 0.2 g magnesium chloride hexahydrate (MP Biomedicals, 67-68-5), and 12 mL water. Samples were centrifuged for 15 min at 12,000×g and supernatant was discarded. Pellets were suspended in 500 μL of disassociation solution containing 6.6 mL of 8 M GuHCl, 3.3 mL of 2-propanol (Fisher Scientific, 67-63-0), and 25 µL of triton X-

100 (Sigma-Aldrich, 9002-93-1). Absorbance of samples were read at 600 nm and concentration of proteoglycans were measured using standard concentration of chondroitin sulfate (Sigma-Aldrich, 39455-18-0). The process is shown schematically in Figure 2.10. 96

2.3.7 Imaging

DIC and fluorescent images were taken on a Nikon inverted microscope (ECLIPSE

TE2000-E) equipped with a CoolSNAP EZ CCD Camera and a high Speed EMCCD

Camera (iXon Ultra 897, ANDOR) for detecting single molecules. The setup is shown in

Figure 2.11.

Figure 2.11. Nikon inverted microscope (ECLIPSE TE2000-E) equipped with a CoolSNAP EZ CCD Camera and a high Speed EMCCD Camera. 97

2.3.8 Fibril Diameter Measurement using TEM and DIC

Since DIC microscopy cannot directly measure submicron fibril diameters, a correlative

method was applied. For fibrils ranging between 60-240 nm, the fibril diameter can be

determined using light microscopy if the fibril diameter distribution and the DIC intensity

distribution are known for a given batch of fibrils [43]. This method has been expanded in

Chapter 3 to develop a more accurate method to measure fibril diameter using DIC microscopy.

The diameter distribution was obtained from TEM images (Figure 2.12.A). To prepare samples for TEM, 20 µL of sclera suspension was added on a 300 mesh Formvar coated

TEM grids (01701-F, Tedpella). After 10 minutes, when fibrils attached on the grid, the grid was wicked dry using a filter paper and stained 3 times on a droplet of 1.5% uranyl acetate for 3 seconds each time. The grid was dried with filter paper and imaged with a

JEOL JEM 1010 transmission electron microscope.

Intensity distribution measurements were performed on the same batch of sclera fibrils using DIC microscopy. First, the fibril suspension was pipetted onto a coverslip coated with 1% bovine serum albumin (BSA, A2153-10G, Sigma Aldrich) and given 1 hour to

adhere. Next, only the fibrils oriented parallel to the shear axis of the light path (northwest-

southeast for an inverted microscope) were selected for intensity measurements to

maximize signal strength and minimize error (Figure 2.12.B). A z-scan was used to find

the maximum intensity signal (Figure 2.12.C), and then 150 intensity measurements were

taken along a 20 m length using a custom MATLAB code. The average intensity shift

μ 98

was determined for 88 fibrils to establish the intensity distribution with a mean of 2687 ±

524 (arbitrary units) and correlated to the diameter of 800 fibrils measured with TEM

(116.7 ± 38.6 nm). The diameters found were in agreement with diameter distributions of

bovine [434, 435] sclera in the literature.

Figure 2.12. Diameter measurement of extracted sclera fibrils. A) Typical TEM image of trypsin extracted sclera fibrils. Images showed native D-banding periodicity and uniform diameter along the length of a single fibril. B) The DIC image shows a typical fibril (oriented in northwest-southeast direction) from the same sclera that was used for TEM imaging. C) The intensity profile across the fibril (along the black arrow) shown in B. Fibril diameters with a mean and standard deviation of 116.7 ± 38.6 nm were correlated to the intensity shift across fibrils with mean and standard deviation of 2687 ± 524 (arbitrary units).

2.3.9 Molecular Association of Labeled Monomers with Native Fibrils Experiment

A glass bottom, ITO coated Delta-T dish (04200417C, Bioptechs) and a perfusable coverglass lid (0420031216, Bioptechs) were plasma cleaned and coated with 1% BSA at 99

4 °C overnight. The dish was placed on the microscope stage and 1 mL of sclera fibril

suspension was added. After an hour incubation for fibril attachment, 20 mL of 1x PBS

was perfused into the dish at 20 ml/h using a syringe pump. A temperature controller

(5410429, Bioptechs) was set at either 25 or 30 °C, and the fibrils were imaged prior to the

addition of labeled monomers. Next 400 µL of 10 µg/mL labeled monomers was added to

the dish at 12 mL/h for 2 minutes to rapidly adjust the collagen concentration inside the

dish to 2 µg/mL (perfect mixing was assumed). Then 5 mL of 2 µg/mL labeled collagen

was added at 1 ml/h to maintain constant concentration inside the chamber. The collagen

concentration was kept at 2 µg/mL (sub-threshold for new fibril formation) to prevent self- assembly and formation of new fibrils [90, 170]. DIC (10 ms exposure time) and fluorescent (5 s exposure time) images were taken every 10 minutes for 6 hours using 60X oil objective (Nikon's CFI Apochromat TIRF Series, Numerical Aperture: 1.45) and NIS-

Element software (Nikon).

2.3.10 Image Analysis for Fluorescence

Incorporation rate and total accumulation of labeled collagen into the scleral fibrils were

measured by tracking the fluorescent intensity of a region of interest (ROI) defined along

the fibril. A 32 µm by 100 µm ROI was defined and the background-subtracted fluorescent

intensity along the fibril length was correlated to the number of collagen molecules [436]

within the effective depth of field. Depth of field (d ) was calculated as sum of the wave

f and geometrical optical depth of field: 100

× n n d = + e (2.2) NA M × NA λ f 2 where ‘λ’ is the wavelength of light, n is the refractive index of the immersion oil, ‘NA’ is

the objective numerical aperture, ‘M’ is the objective lateral magnification, and ‘e’ is the camera pixel size [437]. Fluorescent intensity was converted to collagen concentration by previously measuring the fluorescent intensity of standard fluorescent collagen solutions spanning 1-10 µg/mL.

2.3.11 Arrhenius Plot and Activation Energy Measurement

The reaction rate of labeled collagen monomers binding to the surface of a sclera fibril was

measured as the increase of fluorescent intensity with time. The reaction rate constant (k)

was then calculated using the reaction rate equation:

r = k [C ] [C ] (2.3) m n 1 2 where ‘r’ is the reaction rate and [C1] and [C2] are the collagen concentration in the solution

and on the fibril surface, respectively. The surface concentration was calculated under the

assumption that the monomers were packed with a lateral intramolecular Bragg spacing of

1.58 nm [438, 439] and a periodic gap region of 36 nm between the N-terminus of one

molecule and the C-terminus of the next molecule [440]. Therefore, [C2] was calculated as

-9 2 3.12x10 mol/m and [C1] was set at 2 µg/mL. The reaction orders, m and n, have been shown to be first order during in vitro collagen fibrillogenesis [441]. The reaction rate constant was determined at 25 and 30 °C and the Arrhenius equation: 101

k = Ae (2.4) −Ea⁄RT or

E ln k = ln(A) (2.5) RTa − was used to determine both the activation energy (Ea) and the pre-exponential factor (A) where ‘T’ is temperature in Kelvin and ‘R’ is the universal gas constant. Arrhenius plot is shown in Figure 2.13.

Figure 2.13. Arrhenius plot. By measuring the reaction rate constant at least at 2 different temperatures, the activation energy (Ea) and the pre-exponential factor (A) can be calculated from the equation of the line.

2.3.12 Orientation of Collagen Molecules

We wanted to determine if it is possible to detect the direction of the labeled monomers as they associate with single collagen fibrils. Labeled and unlabeled monomers were mixed 102

1:20000 in PBS at pH 7.3 and a total collagen concentration of 50 µg/mL. The mixture was incubated at 37 °C in a Delta-T Dish to allow for fibrillogenesis on the glass surface. After reaching equilibrium, reconstituted fibrils and fluorescently labeled collagen monomers were imaged at 60x with high speed EMCCD camera (iXon Ultra 897, ANDOR). The image processing to obtain the orientation of the ellipse major axis was performed in

Matlab (Mathworks, Inc., Natick MA) as follows: images were uploaded into Matlab, filtered using a Gaussian filter, and then background subtracted. The binary image was then processed using the function regionprops to obtain the orientation of the ellipse.

2.3.13 Statistical Information

Data are shown as mean ± standard deviation. No statistical tests were used to predetermine sample sizes, but our sample sizes are similar to those generally employed in the field.

Significance was set to P < 0.05. Significance of the mean was calculated using two-tailed t-test or one-way ANOVA. The sample sizes and the results of hypothesis tests are reported for each experiment in the results section.

2.4 Results

2.4.1 The Degree of Labeling (DOL)

Monomeric collagen was labeled with AF488 and the DOL was defined as moles of dye per mole of protein. The DOL achieved at multiple pH values is shown in Figure 2.14. The initial molar ratio of dye:collagen ranged from 3-15 and the resulting dye:collagen ratio 103

ranged from 2-10 fluorophores per collagen molecule. ANOVA test with a significance

level of 0.05 showed that labeling performance is not significantly different at pH 7.5, 8.0,

and 8.5 within each initial molar ratio of AF488:Coll groups. F values and degrees of

freedom of the ANOVAs for 3, 9, and 15 initial molar ratio of AF488:Coll groups were F-

value = 0.70, 0.20, and 0.25 and df = 8, 8, and 5, respectively.

Figure 2.14. DOL achieved at pH 7.5, 8.0, and 8.5 by adding 3, 9, and 15x excess moles of Alexa Fluor 488 to moles of collagen (AF488:Coll). ANOVA test with a significance level of 0.05 showed that labeling performance is not significantly different at pH 7.5, 8.0, and 8.5 within each initial molar ratio of AF488:Coll groups. However, labeling performance was significantly different between initial molar ratio of AF488:Coll groups. Data are expressed as mean ± standard deviation (n=3 replicates).

Therefore, subsequent experiments were performed at pH 7.5 to maximize the likelihood of attaching a fluorophore at the N-terminal of the monomer [417]. It is beneficial to bind

one fluorophore to the end of the protein because it maximizes the separation between the 104

N-terminal bound fluorophore and a secondary fluorophore closest to the C-terminal.

When the fluorophores span a greater distance on the protein, it enhances the ability to

determine the orientation of the protein after it binds to the fibril. Furthermore, statistical

analysis showed that increasing the initial molar ratio of AF488 to collagen increased the

DOL (ANOVA test with a significance level of 0.05). F value and degree of freedom of

the ANOVA were F-value = 53.22 and df=32.

2.4.2 De Novo Fibrillogenesis Experiments with Labeled Collagen

The first objective was to test the functionality of our fluorescently labeled collagen

molecules. To do this we took advantage of the well-characterized ability of collagen molecules to spontaneously undergo de novo fibrillogenesis under physiological conditions

[67].

The fibrillogenesis kinetics of labeled collagen was assessed via turbidity. The sigmoidal turbidity curve observed during collagen assembly has two distinct phases: I) a lag phase during which there is no detectable change in absorbance and II) a linear growth phase during which the fibrils grow in size and absorb more light [84]. The kinetics of fibrillogenesis of labeled monomers was studied and compared to unlabeled monomers by measuring the lag time, plateau time, and maximum change in absorbance at 313 nm.

Figure 2.15 shows the fibrillogenesis kinetics for collagen solutions where 100% of the monomers possessed a DOL of 0, 2, 4, and 9. Fibrillogenesis was affected even at a low

DOL (~ 2 dyes:molecule), whereby the lag phase increased and the total absorbance 105

decreased. Increasing DOL decreased the maximum absorbance and delayed the lag and

plateau time. Such a delayed growth phase has been reported for hetero/homotrimer

mixtures of type I collagen [413]. Fibrillogenesis of labeled monomers with DOL larger

than ~9 did not show the typical sigmoidal shape turbidity curve. Data are presented as

mean ± standard deviation (n=3 replicates per group). Table 2.1 summarizes the values of lag time, plateau time, and total absorbance change for each tested condition. Lag time, plateau time, and total absorbance change of labeled monomers with DOL of 2, 4, and 9 were significantly different (two-tailed t-test, α=0.05) from unlabeled monomers.

Figure 2.15. Turbidity of 50 µg/mL labeled collagen at 37 °C. In this experiment, 100% of the collagen molecules were labeled. Each labeled molecule had an average DOL of 0, 2, 4 or 9. Lag and plateau times were calculated as the time required to reach 10% and 90% of the maximum turbidity, respectively. Data are presented as mean ± standard deviation, n=3 replicates per group. 106

Table 2.1. Absorbance, lag time, and plateau time of sigmoidal turbidity curve of 50 µg/mL labeled collagen at 37 °C. In this experiment, 100% of the collagen molecules were labeled. Lag and plateau times were calculated as the time required to reach 10% and 90% of the maximum turbidity, respectively. Data are presented as mean ± standard deviation, n=3 replicates per group. The highlighted row shows lag time, plateau time, and total absorbance change of unlabeled monomers which were significantly different (two-tailed t-test, α=0.05) from labeled monomers.

The de novo fibril assembly data clearly demonstrate that the label interferes with the kinetics of assembly and that the effect decreases with degree of labeling. We then asked if the labels disrupted the morphology of the network or the nanostructure of the assembled fibrils. DIC, fluorescence, and TEM images were used to determine if the fluorophores altered the fibrillar structure and network architecture.

Figure 2.16.A shows collagen fibrils formed from a 50 µg/mL solution of unlabeled collagen molecules. A labeling ratio of DOL≈2 (Figure 2.16.C) produced an average fibril length and fibril count that was nearly indistinguishable from the unlabeled sample. Fibrils formed from collagen labeled with DOL≈4 were both shorter and sparser in number (Figure

2.16.E). When the labeling was increased to DOL≈9, fibrillogenesis was strongly disrupted and non-fibrillar aggregates were mostly observed (Figure 2.16.G). 107

Figure 2.16. Collagen fibrils formed in neutralized solutions of 50 µg/ml collagen at 37°C. A) DIC and C), E), and D) Fluorescent images of fibrils formed by monomers with DOL of 0, 2, 4, and 9 respectively. We used DIC imaging for the unlabeled fibrils (DOL =0) because there was no fluorescence signal. B), D), F), and H) TEM images of fibrils formed by monomers with DOL of 0, 2, 4, and 9 respectively. 108

The visibility of the D-banding - a hallmark of collagen fibril formation - in TEM images

decreased with increasing the DOL (Figure 2.16.B, D, F, and H). However, analysis of

TEM images showed that the D-banding period remained ~67 nm, regardless of the DOL.

This would indicate that despite the difference in network morphology, the labeling process did not impact the ultrastructure of the fibrils. We thus concluded that while the small label affected the kinetics of assembly, the final fibrillar structure remained “native”. Our next goal was to adjust the concentration of labeled monomer to determine what concentration ratio (labeled/unlabeled) would yield nearly normal kinetics.

To minimize the effect on the assembly kinetics and network morphology, the remaining experiments were performed using a DOL≈2. However, it is important to remove as much label-induced artefact as possible from the assembling system. Our hypothesis was that including a small number of labeled monomers relative to unlabeled monomers would restore the kinetics to baseline at some threshold value. To find that threshold, we re- examined the assembly kinetics at our lowest DOL (~ 2 dyes:molecule), while the percentage of labeled monomers was varied from 100% to 1%. As shown in Table 2.2, the fibrillogenesis kinetics were not significantly altered from the unlabeled control when up to 5% labeled monomers was used. As the percentage of labeled monomers increased, the effect was more pronounced with 100% labeled monomers reducing the rate of fibrillogenesis by ~70%, increasing the lag time by ~56% and lowering the optical density by ~34%. We thus felt comfortable that using our lowest DOL which still permits tracking molecular orientation would permit us to observe the kinetics of assembly while minimizing perturbation of the process by the dye molecule. 109

Table 2.2. Fibrillogenesis of labeled (DOL≈2) and unlabeled monomers mixture. Changes in kinetics of fibrillogenesis is negligible (less than 10% error) in mixtures containing 5% or less labeled monomers (shown in highlighted rows). Data are presented as mean ± standard deviation (n=3 replicates per group).

2.4.3 Single Molecule, Multi-label Fluorescence Orientation Microscopy (SMO Microscopy)

The work in this section was completed with contribution from Dr. Charles DiMarzio and

Dr. Monica Susilo.

One of our goals from fluorescently labeling collagen molecules is to determine whether exogenous collagen is associating non-specifically or functionally with the fibrils. Non-

specific associations of monomers would show a random molecular orientation distribution

relative to the fibril axis, while functional associations would exhibit molecular co-

alignment with the fibril axis. Since the fluorescent molecules were located at two points

along the collagen molecule, the epi-fluorescence image of a doubly-labeled collagen monomer is the incoherent superposition of two point-spread functions (Figure 2.17.A and

B) of the microscope at the emission wavelength with each one centred on one of the two

fluorophores. Typically, when the separation is smaller than the resolution, the image can 110

be approximated into an ellipse (Figure 2.17.C), where the major axis represents the collagen molecule orientation. As a proof of concept, collagen fibrils were reconstituted with mixtures of labeled and unlabeled monomers. After reaching equilibrium, single collagen molecules that possess two fluorophores were detected (Figure 2.17.D). The pixels of the fluorescing molecules were fitted with ellipse (Figure 2.17.E), where the major axis represents the molecule orientation (Figure 2.17.F). These initial results show the potential to establish a new single molecule microscopy method (SMO microscopy) capable of determining the orientation of monomers as they interact with fibrils.

We further produced synthetic images of labeled collagen monomers to estimate stochastic distribution of fluorophores along the collagen molecule and microscope parameters that will provide orientation information. Analysis of the images shows that there are subsets of the collagen molecules that generate point spread functions with equivalent major and minor axis. This can be a characteristic of a singly-labeled monomer, or a doubly-labeled monomer where the fluorophore separation exceeds the resolution of the microscope and the signal-to-noise ratio is sufficient. In the latter case, two distinct point spread functions will appear in the image, and the orientation of the monomer will be the vector direction from one to the other. However, in most cases the separation will be smaller than the resolution. In this case, a computationally simple approach can be used to calculate the covariance matrix of the two-dimensional position vector. Finding the eigenvalues and eigenvectors of this matrix approximates the image as an ellipse with the higher and lower eigenvalues representing the lengths of the major and minor axes and the corresponding eigenvectors representing their directions. 111

Figure 2.17. SMO microscopy showing orientation of type I bovine collagen molecules associated with fibrils. When collagen is labeled with two fluorophores separated by a large enough distance (A), it is possible to fit an ellipse to the pixel representation (B). Thus, we can determine if the single molecules are co-aligned with the fibril axis or just associating (C, D, and E). Courtesy of Dr. Monica Susilo.

The solution of the eigenvalue problem is limited by the spacing of the fluorophores

relative to the size of the point-spread function and the Poisson distribution of photons detected in each pixel of the image. Additionally, noise can result from the detector, from incomplete rejection of scattered illumination, and from autofluorescence. Although the mean of this noise can be subtracted, its variations cannot. 112

We have computed receiver operating characteristics (ROC) for different spacings and

numbers of photons per fluorophore. To accomplish this, we generated synthetic images

with Poisson random noise, computed the eccentricity as the ratio of major to minor axis

and compared to a threshold. If the eccentricity exceeded threshold we recorded a

detection. We repeated the simulation with one fluorophore and if the eccentricity exceeded

threshold we recorded a false alarm. Each set was repeated 200 times and the probability

of detection (PD) was plotted against the probability of false alarm (PFA).

Using microscope and fluorophore parameters we anticipate a maximum of a few thousand photons per fluorophore, but a low illumination level is desired in order to observe signals for a period of time without photobleaching. Thus in our computation we used between 25 and 2500 photons per fluorophore. We began with one fluorophore on each end of the monomer (spacing of 300 nm), and successively divided the spacing by 2 until the area under the ROC curves became close to 0.5.

Figure 2.18 shows three synthetic images with the major and minor axes plotted. Figure

2.18-A shows the image for 25 photons per fluorophore at a spacing of 300 nm. With this large spacing, the two fluorophores are clearly resolved even with so few photons. Figure

2.18.B shows a single fluorophore for comparison. Figure 2.18.C shows the image for 2500

photons per fluorophore at a spacing of 68 nm. At this point it becomes very difficult to

determine that there are two fluorophores present.

Figure 2.19 presents ROC curves for the same two spacings. The curves in each case were computed with, from top to bottom 2500, 500, 250, 100, 50, and 25 photons per fluorophore. For the larger one, the detection statistics are nearly perfect at all numbers of photons. For the smaller one, there is a slight improvement over random chance only for

113

very large numbers of photons. We note that for low numbers of photons the ROC curve

is below the 45-degree line, which represents random chance. This result occurs because

with small numbers of photons, a few random detection errors can alter the symmetry of

the image. The doubly-labeled monomers, with twice as many photons are less susceptible

to this phenomenon.

Figure 2.18. Synthetic images of labeled collagen monomer with the major and minor axes plotted. The colour is representative of the number of photons per pixel. A) Synthetic image of doubly-labeled monomer for 25 photons per fluorophore at a spacing of 300 nm. B) Synthetic image of singly-labeled monomer for 25 photons per fluorophore for comparison. C) Synthetic image of doubly-labeled monomer for 2500 photons per fluorophore at a spacing of 68 nm. Courtesy of Dr. Charles A. DiMarzio.

Aberrations in the microscope optics, sampling limitations imposed by the camera, and non-uniform noise may further reduce performance. On the other hand, more rigorous processing approaches may improve performance. At present, we anticipate being able to determine the orientation of a useful number of monomers with fluorophore spacings exceeding about 70 nm. Beside the N-terminal α-amine, there are 38 lysines that are potential locations of the label. 79% of these lysines are farther than 70 nm from the N- terminal α-amine [418]. Given a high probability (e.g. more than 50%) that one of the

114

labels is on the N-terminal, we expect that at least 40% of the labeled collagens will have

detectable orientation ellipses.

Figure 2.19. ROC curves for A) 68 nm and B) 300 nm spacings. The curves in each case were computed with, from top to bottom 2500, 500, 250, 100, 50, and 25 photons per fluorophore. For the 300 nm spacing, the detection statistics are nearly perfect at all numbers of photons. Courtesy of Dr. Charles A. DiMarzio.

2.4.4 Labeled Collagen/Native Fibril Association Experiments

Young, native collagen fibrils were trypsin extracted from 1-10-day old bovine sclera.

Since proteoglycans can influence the association of collagen monomers with fibrils [21], we determined if they were present on our extracted fibrils. Extracted fibrils were separated from the extraction solution and stored in 10 mM hydrochloric acid. The proteoglycan analysis showed that more than 99% of proteoglycans were removed from scleral fibrils during extraction. The concentration of proteoglycans in the extraction solution (without collagen fibrils) and the collagen fibril solution were measured as 65 ± 4.2 and 0.25 ± 1.2

µg/mL, respectively (n=6). Thus, our system is testing the ability of collagen to interact with fibrils in the absence of proteoglycan control [71, 442] (e.g. at the maximum association rate). Figure 2.20 shows the standard curve and the calculated proteoglycans concentration.

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To determine the maximum association rate of labeled collagen with fibrils, proteoglycan- free scleral fibrils were incubated in the presence of labeled collagen monomers. Collagen monomer solutions at 2 µg/mL and comprising either 5% or 100% labeled monomers associated with the native fibrils with the same lag and plateau times at 25 °C, showing that the reaction rate was not affected by the number of fluorescently labeled monomers

(unlike during de novo fibrillogenesis) and labeled monomers were not outcompeted by unlabeled monomers. There was no significant difference (two-tailed t-test, α=0.05) in lag time (t(12 degree of freedom)=-0.10, tcritical=2.18, P=0.92) and plateau time (t(12 degree of freedom)=1.10, tcritical=2.31, P=0.30) when 5% or 100% of monomers were labeled.

Figure 2.20. Quantification of proteoglycans with Alcian blue (n=6). The concentration of proteoglycans was measured using the standard concentration of chondroitin sulfate. “Pellet” refers to the sample containing separated fibrils after extraction and “Supernatant” refers to the sample containing the fibril extraction suspension without the fibrils. Note that some error bars are too small to be seen. Courtesy of Alexandra A. Silverman.

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Figure 2.21. Incorporation of labeled collagen into fibrils. Single, native sclera collagen fibrils incorporate exogenous labeled monomer with kinetics similar to fibril assembly. (A), (B), and (C) Fluorescent images show incorporation of labeled monomers on a typical fibril over time at 25 °C. (D) The graph shows typical accumulation of labeled monomers into sclera fibrils at 25 and 30 °C. Lag and plateau times in minutes (the times when the fibril intensity reached 10% and 90% of its maximum intensity) were measured as 59.9 ± 19.4 and 111.8 ± 37.4 at 25 °C (n=21 replicates) and 43.5 ± 11.7 and 97.8 ± 20.7 at 30 °C (n=23 replicates), respectively.

Therefore, collagen monomer solutions at 2 µg/mL and comprising 100% labeled monomers were added to the native fibrils at 25 and 30 °C. Figure 2.21 shows that monomers actively engage with the native fibril surface in a two-phase process (i.e. lag

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and growth) similar to that observed by turbidity during collagen fibrillogenesis de novo.

The association rate and net accumulation (until saturation) of labeled monomers into/onto

scleral fibrils were measured as 2.8 ± 1.2 and 4.0 ± 1.5 molecules per m2 per minute and

114.3 ± 26.8 and 207.1 ± 55.4 molecules per m2 at 25 °C (21 replicates)𝜇𝜇 and 30 °C (23

replicates), respectively. Significance of the means𝜇𝜇 was calculated using two-tailed t-test and α=0.05; t(42 degree of freedom)=2.76, tcritical=2.02, P=0.008 for association rates and

-8 t(42 degree of freedom)=-7.00, tcritical=2.02, P=1.44×10 for net accumulations. Given our

estimated fibril diameters, at the plateau (saturation), the available space on the fibril

surface would be 1/16 and 1/9 covered by exogenous monomer at 25 °C and 30 °C,

respectively. Using an Arrhenius plot, activation energy and pre-exponential factor were

measured as 12.6 kcal/mol and 6.5x109 m3/(s.mol).

Figure 2.22. Photobleaching of fibrils after reaching equilibrium with 2 µg/mL labeled monomers. Fibrils were imaged continually with 1 second exposure time.

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To consider the effect of photobleaching, 4 fibrils were incubated at 30 °C with 2 µg/mL

labeled monomers for 6 hours in a similar experimental setup. After reaching equilibrium,

images were captured every 5 seconds continually with 1 second exposure time. As it can

be seen in Figure 2.22, fibrils intensity was not changed significantly at least for 200 s exposure time. Therefore, we assumed no photobleaching during this experiment.

2.5 Discussion

It has been shown that collagen molecules are produced at a relatively high rate by resident

cells during tissue expansion [443] and that the growing fibrils are bathed in a monomeric

collagen solution that is constantly recycled [444]. Collagen fibrils assembled in vitro are likely to be in dynamic equilibrium with collagen monomers in the local solution with an equilibrium concentration of less than 10 µg/mL [80]. Kadler et al. [90] reported a temperature dependent critical concentration decreasing from 4.73 to 0.12 µg/mL when temperature was increased from 29 °C to 41 °C. We hypothesize that growing collagen fibrils are likely to be constantly exchanging molecules at their surfaces and that the surface exchange rates (kon and koff) are modulated by both chemical and mechanical [445] signals.

However, the baseline exchange rates for native collagen fibrils are not known.

Part of the difficulty in establishing kon and koff is the insoluble nature of the collagen fibrils and the uncertainty of the nature of their association with soluble monomers/microfibrils.

While estimation of the rate constants for collagen fibril surface erosion and adsorption requires relatively simple quantitative fluorescence microscopy, determining whether collagen monomers are incorporating functionally into extant fibrils is more difficult. The goal of this study was to develop methods which will enable us to ask quantitative questions

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about the dynamics of molecular assembly and disassembly at the collagen fibril surface.

Figure 2.17 demonstrates our potential ability to determine the orientation of single

collagen molecules, using SMO, that have been labeled with at least two fluorophores as they associate with single collagen fibrils. When collagen is labeled with two or more fluorophores separated by a large enough distance, it is possible to fit an ellipse to the pixel representation. Thus, theoretically, we can determine if the single molecules are co-aligned

with the fibril axis (incorporating) or simply associating. The data we have provided show

both association and alignment for some of the exogenous labeled molecules along the

fibrils, indicating that SMO has the potential to differentiate between functional and non-

functional association. However, more work is needed to verify the methodology. We have

thus restricted our analysis to the general accumulation of collagen monomer on the fibrils.

The activation energy of 12.6 kcal/mol calculated in the present study was lower than

values reported in the literature for de novo collagen assembly in vitro (27 – 58 kcal/mol)

[80, 90, 441], suggesting that the activation energy for growth of a native fibril is less than

formation and growth of a reconstituted fibril in vitro which one would expect. However,

it is possible that the difference might be caused by systemic inaccuracies in our fibril

diameter measurement. It has been shown that electron microscopy images taken on

processed collagen fibrils underestimate fibril diameters due to dehydration of samples

[446]. Corneal fibrils in normal hydration and free solution have diameters around 40%

larger than dehydrated fibrils [447]. Conversely, fibrils might flatten and adhere to the glass

surface and lose their round cross sectional shape [121], partially blocking the available surface area on the fibril to react with monomers.

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Moreover, it has been shown that proteoglycans are removed from collagenous tissue by

trypsin treatment [71, 442] similar to that which we employed. Therefore, the lateral

growth of our scleral fibrils and the low activation energy might be attributed to the absence

of proteoglycans. Proteoglycans are a superfamily of molecules in ECM that are

distinguished by the covalent attachment of one or more highly negatively charged

glycosaminoglycan chains to their core proteins [209]. The scleral extracellular matrix has

been shown to contain three major proteoglycans; biglycan, decorin [448], and aggrecan

[449]. These proteoglycans with their sulfated glycosaminoglycans can regulate matrix

hydration [210], interact with collagen fibrils [211], inhibit fibrillogenesis rate [212], and

change fibril orientation [215] and fibrillar structure [216] of collagenous ECM. They have

also been shown to control fibril diameter [217] by directly influencing molecular assembly

[21] or lateral fusion of fibrils [22]. Although this study was performed in the absence of

proteoglycans, we recognize the capacity of other collagen binding molecules, such as

proteoglycans and fibronectin, to modify fibril diameters [25] and formation kinetics [174].

Future studies with a more inclusive set of biological molecules will continue to refine our

understanding of molecularly driven growth.

During initial fibrillogenesis (Figure 2.15) the lag time we observe is likely due to the lack

of pre-existing nucleation sites that must be first established. However, this is not the case in Figure 2.21 where monomers are incorporating into the already extant native fibrils. The lag time we observed might be due to formation of microfibrils on the tested fibril’s nucleation sites. Note that performing the experiment under the critical concentration was to prevent formation of de novo fibrils. However, monomers can still associate with the

native fibril directly and/or initiate microfibrils on the fibril surface at a concentration lower

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than the critical concentration. Presence of proteoglycan-free native fibrils can reduce the activation energy of monomer/fibril reaction by providing nucleation sites on the fibril surface.

The plateau region of the turbidity experiment (Figure 2.15) is due to consumption of the

reactant, while our monomer/fibril association experiment (Figure 2.21) had a constant supply of monomers. Our results suggest that at the plateau region of monomer/fibril association experiment, 6% and 11% of fibril surfaces were occupied by labeled monomers at 25 °C and 30 °C, respectively. Considering the constant supply of subcritical concentration of labeled monomers at both temperatures (preventing formation of microfibrils) and saturation of scleral fibrils by different amounts of labeled monomers shows that in addition to proteoglycans, there must be other inhibiting factors that alter the availability of binding sites on fibrils and arrest the fibril’s growth in our system. However, we do not currently have a good explanation for the plateauing effect. Ultimately, we suspect that collagen fibrils reside in a state of dynamic equilibrium with the local extracellular milieu and that this equilibrium can potentially be altered by multiple factors

(i.e. mechanical forces, matrix proteins, ionic concentration, etc.) that could shift the delicate balance of the molecular association rates (kon and koff) between fibrils and molecules.

2.6 Summary

In summary, we have developed and validated a small “functional” fluorescent labeling

method for collagen molecules which permits their tracking during assembly of fibrils

while minimally affecting the assembly kinetics and their ability to form native-like fibrils.

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We provide detailed information about the number of labels that can be tolerated and the

concentration of labeled monomers that can be included in an assembling solution without

appreciably influencing the kinetics, the network structure or the collagen fibril nanoscale

organization. We then used the label to assess the kinetics of monomer association rate

with native extant collagen fibrils. The results tell us that fibrils attract monomer with an

energy barrier lower than that required for initial collagen formation and that the shape of

the kinetics curve is sigmoidal, displaying an upper limit on association.

Our next accomplishment was to introduce a new form of dynamic microscopy that has the

potential to not only track collagen monomer association with fibrils, but can determine if

that association is functional. The method uses the separation of two labels to detect the

molecular orientation which should co-align with the fibril axis if it is a functional

association rather than just non-specific. We discuss the optical algorithm and showed initial molecular orientation tracking data as molecules associate with reconstituted collagen fibrils.

Taken together, these developments will make it possible to dynamically track and assess molecular association with fibrils, paving the way for live, dynamic imaging of matrix assembly in vitro in optically accessible living systems.

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3 A CORRELATIVE METHOD TO MEASURE COLLAGEN FIBRIL DIAMETER IN DIFFERENTIAL INTERFERENCE CONTRAST MICROSCOPY

3.1 Introduction

Collagen is the most abundant protein in vertebrates and the principle load bearing

molecule in their tissues. While there has been over 50 years of research into collagen fibril

assembly and growth, we are still remarkably deficient in our knowledge of the

mechanisms that drive animal structure formation and evolution. Collagen fibrils, 20 nm

to a few hundred nanometers in diameter [188, 450], are the basic building block in the

complex hierarchical structure of collagenous tissues. However, our ability to conduct

quantitative studies of fibril dynamics and mechanics is limited. The theoretical limit of

visible light microscopy, 1.22λ/(2NA), translates to approximately 220 nm. This resolution

limit does not permit very accurate determination of collagen fibril diameter or changes in

diameter. Such measurements are critical to understand basic assembly and degradation

dynamics in controlled systems.

Fibril diameter has been measured using different methods such as small-angle X-ray

scattering (SAXS), TEM, SEM, AFM, and second-harmonic generation microscopy

(SHG). All of these methods have been extremely valuable in studying of collagen fibrils

but they all lack the ability to measure collagen fibril diameter in live studies performed

124 with light microscopy. Alternatively, DIC microscopy has been used to visualize dynamic structures that are as small as microtubules (25 nm diameter) and has been shown to be sensitive to the size of objects smaller than the wavelength of light [451]. In this investigation, we take advantage of DIC microscopy’s ability to report dimensions of nanometer scale objects to generate a curve that relates collagen diameter to DIC edge intensity shift (DIC-EIS). This simple, non-destructive, label free method should advance our ability to directly examine fibril dynamics under conditions that are physiologically relevant. In this section, common methods to measure collagen fibril diameter are reviewed and their limitations are discussed.

3.2 Current Methods for Measuring Collagen Fibril Diameter

3.2.1 SAXS

SAXS is a common method to measure average collagen fibril diameter of a tissue in its native state without any additional treatment. When X-rays pass through a collagenous tissue, they provide detailed information on arrangement of collagen molecules and fibrils

[452-454]. SAXS has been used widely to measure fibril diameter in many studies such as developing chick metatarsal tendons [455], across human cornea [456, 457], cornea of different species [438], as a function of human cornea age [439].

Figure 3.1 shows a schematic of SAXS from collagen fibrils in cornea. Packing of collagen fibrils in cornea gives rise to a series of equatorial patterns that represent fibril diameter as well as lateral spacing between fibrils. The meridional reflections represent the D- periodicity of fibrils.

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Figure 3.1. Small-angle x-ray diffraction from corneal collagen fibrils. From Boote et al. (2003) [456].

Overall, SAXS is non-destructive and allows for a statistically robust sampling. However, it provides measurement of average collagen fibril diameter of a tissue and it has not been used for individual fibrils.

3.2.2 TEM

TEM has been widely used in study of collagen fibrils since 1940s (these early studies are summarized by Harkness [458]) to quantify the dimensions of collagen fibrils in variety of tissues such as skin [459], ligaments [327, 460-463], cornea [165], rat tail tendon [188,

190, 464], limb tendons [323, 465], adult human tendon [466], developing chick metatarsal

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tendon [467] as well as reconstituted fibrils [67, 82, 468]. TEM enables direct measurement of diameters of collagen fibrils as well as three-dimensional organization of fibrils [469].

Figure 3.2. Electron micrograph of type I collagen fibrils in both transverse and longitudinal orientations. Scale bar is 300 nm. Reprinted with permission from Cassella et al. [470].

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To prepare samples for TEM, tissue need to be dehydrated, fixed, embedded in resin, cut

into extremely thin slices, and stained. Transverse sections are usually used to measure fibril diameter. Alternatively, extracted fibrils or reconstituted fibrils can be dried and stained on TEM grids which will lead to a longitudinal fibril view. Figure 3.2 shows type

I collagen fibrils in both transverse and longitudinal orientations.

Nevertheless, TEM preparation procedure is destructive due to dehydration [464], fixation

[190], and sectioning [323]. It has been show that fibrils can shrink relative to the wet

fibrils in physiological conditions [467] and flatten on the support surface of TEM grid

[294].

3.2.3 SEM

TEM was the main tool available for studying the fine morphology of the collagen fibrils

until invention of high resolution SEM in late 1960s. SEM enabled the ability to visualise

the three-dimensional collagen network and provided further insight into the nature of the

collagen fibrils and their size [471, 472]. Since then, SEM has been used to measure fibrils

diameter in different tissues such as articular cartilage [473, 474], human teeth [475],

intervertebral discs [476], ligament and tendon [477], or to study the effect of

proteoglycans in the process of fibril formation [478, 479]. Samples are usually dried,

mounted on a metal stub and then coated to be observed under a SEM. Figure 3.3 shows a

high resolution SEM image appropriate to measure collagen fibril diameter.

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Figure 3.3. High-resolution SEM view of collagen fibrils from the rat tail tendon. Reprinted from Ohtani (1992) [480].

3.2.4 AFM

The surface of collagen fibrils without any specific preparation can be investigated in ambient conditions using AFM. AFM images are obtained by scanning the fibril surface with a finely shaped tip held at an atomic distance and probing the interactions between the tip and the fibril surface. Atoms at the apex of the tip and the atoms on the fibril surface interact with each other via short-range chemical forces and long-range Van der Waals and

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electrostatic forces. Researches have used AFM to measure collagen fibril diameter of

reconstituted fibrils [481-483] as well as native fibrils of different tissues such as cornea

and sclera [484-486] and rat tail tendon [487]. Figure 3.4 shows an AFM image of reconstituted collagen fibrils.

Figure 3.4. AFM images of reconstituted collagen fibrils. Reprinted with permission from Revenko et al. (1994) [483].

The most important advantage of the AFM compared to electron microscopy methods is the ability to image samples in their native state. However, there are still some limitations:

AFM has a rather slow temporal resolution, it is necessary to consider image distortion due to the tip size, and fibril adhesion to surfaces could change the fibril’s shape [486].

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3.2.5 SHG

When an anisotropic molecule is placed in the electric field of an exciting light, it creates

an oscillating field at twice the frequency (second harmonic). Collagen has a highly

anisotropic molecular structure which makes it a strong source of SHG [488, 489]. SHG

of collagen fibrils was first demonstrated by Freund et al. [490] and since then has been

used to study of collagen fibrils both in vivo [491] and in vitro [492] and for detecting a

wide range of pathological conditions such as fibrotic tissues [493, 494], cancerous tissues

[495-499], and osteogenesis imperfecta [500].

Figure 3.5. Correlative SHG-TEM imaging. A) SHG and B) TEM images of unstained collagen fibrils (scale bar, 10 mm). C and D) Zoomed regions of interest from A and B (scale bar, 5 mm). E) SHG photon number as a function of the fibril diameter measured on the TEM image. F) All samples using diameter bins of 50 nm. Reprinted with permission from Bancelin et al. (2014) [501].

However, measurement of fibril diameter is not directly possible from the nonlinear optical signal of SHG microscopy. Bancelin et al. [501] developed a correlative method to

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determine collagen fibril diameter using SHG and TEM images. They reported absolute measurement of individual fibrils as small as 30 nm in diameter with ± 25 nm accuracy

(Figure 3.5).

SHG has the ability to detect collagen fibrils without introducing any exogenous probes.

Although, there are still some limitations since SHG microscopy is expensive and

technically demanding. Also, the SHG can only be used for unipolar fibrils with constant

collagen density and the signal depends on the fibril angle which can introduce inaccuracy

in quantitative studies [502].

3.3 DIC Microscopy

All of the above methods have been extremely valuable in studying of collagen fibrils but

they all lack the ability to measure collagen fibril diameter in live studies. Alternatively,

DIC microscopy is one of the most popular methods to visualize unstained fibrillar

collagen.

In DIC microscopy, the illuminating light is first plane polarized by the polariser and then

split by a Wollaston prism into two orthogonal components with a slight lateral shear between them. After passing through the condenser lens, specimen, and objective lens, the lateral shear is removed by a second Wollaston prism. And finally the two components are made to interfere by an appropriately oriented analyser. The produced pseudo-3D image demonstrates the specimen features like differences in refractive index or thickness. The

DIC optical path is shown in Figure 3.6.

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Figure 3.6. Schematic diagram of the DIC optical path showing the optical components (left) and their effects on the orthogonally-polarised light (right). From Cogswell et al. (1997) [503].

In DIC microscopy, the linearity and contrast of the final image are dependent on the bias phase which is usually introduced by rotational polariser or the lateral position of the objective Wollaston prism. At a bias phase setting of zero, a highly non-linear phase only image can be obtained which shows bright regions wherever positive or negative phase gradients occur. Instead, at a bias phase setting of 45 degrees, the most linear response to specimen phase gradients occurs. However, the final image is a combination of phase and amplitude information which makes it difficult to measure phase alone. The behaviour of a DIC system for different values of bias retardation is shown in Figure 3.7.

A major advantage of DIC is that collagen fibrils can be studied directly in solution or in a bioreactor [334] without previously being dehydrated, fixed, or stained. Also, its highest

133 spatial frequencies are tightly confined to the plane of focus of the microscope which rejects most of out-of-focus blurring effects.

Figure 3.7. Behaviour of A) the signal strength, B) the contrast from a phase change, C) the linearity of the response, and D) the signal-to-noise ratio of a DIC system for different values of bias retardation. Reprinted with permission from Cogswell and Sheppard (1992) [504].

Major progress in science is often enabled by the development of new technologies or methodologies. Biological specimens are weak-phase objects and their conventional images have low contrast and poor visibility. DIC microscopy was introduced over a half of a century ago for the study of phase objects and has been widely used by biologist since then. Later, a video-enhanced DIC method was introduced by Allen et al. [451] to improve performance of DIC microscope and image microtubules as small as 25 nm diameter. The method was widely used, for example, to measure microtubules growth velocity [505] and image reconstitution of physiological microtubule dynamics [506], and enabled the

134 discovery of kinesin [507]. However, due to innate nonlinearities, DIC microscopy has been only used for qualitative imaging and was therefore not appropriate for accurate diameter measurements of those small biological samples.

Previously, attempts were made to measure collagen fibril diameter with DIC microscopy

[43, 508] and fibril images were simulated using MIE scattering theory. A linear relationship between fibril diameter and DIC Edge Intensity Shift (DIC-EIS) was predicted at that time (Figure 3.8). However, due to vibration of fibrils in the thin field of view of

DIC, fibril diameter was not resolved experimentally. In this investigation we used electron microscopy methods in combination with DIC to show not only the sensitivity of DIC-EIS to diameter change, also to correlate DIC-EIS to fibril diameter without needing further electron microscopy processing.

Figure 3.8. DIC-EIS for simulated DIC images with different diameter fibrils (for a 60X objective and 1.45 NA). Note the linear region between 60 nm and 240 nm diameter fibrils. From Brendan Flynn dissertation, Northeastern University (2012) [509].

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3.4 Experimental Methods

3.4.1 Fibril Preparation for TEM-DIC imaging

Type I collagen fibrils were extracted from bovine scleras as previously described in

Chapter 2. Referenced TEM grids (01841-F, Ted Pella Inc.) were placed on a rotatable dish and between two small cover glasses (1217H20, Thomas Scientific) to not directly touch the bottom glass and prevent distortion of the formvar layer. A 20 µL volume of the fibril suspension was pipetted on top of the grid. Fibrils were air-dried on the TEM grid and imaged with DIC microscopy at different angles to the shear axis of the light path. The experimental setup is demonstrated schematically in Figure 3.9.

Figure 3.9. The experimental setup where collagen fibrils were air-dried on a referenced TEM grid and imaged with DIC microscopy. The collagen fibril is not in scale.

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3.4.2 DIC Imaging

All DIC images were taken on a Nikon inverted microscope (ECLIPSE TE2000-E)

equipped with a CoolSNAP EZ CCD Camera and a 40X objective (Plan Fluor ELWD 40x

Nikon, NA 0.6). Fibrils were scanned in z-direction (0.1 µm steps) to find the best plane of focus.

3.4.3 TEM Processing

TEM grids were stained three times on a droplet of 1.5% uranyl acetate for three seconds

each time. The grids were dried with filter paper and imaged with a JEOL JEM 1010

transmission electron microscope.

3.4.4 Microfabrication of Trenches in PDMS Sheets

Standard UV lithography processes were used to create a mold for casting the

microstructures. Trenches of 25 µm wide and 20 µm deep were designed in SolidWorks

software and printed out on high-resolution transparencies. Single side polished silicon (Si) was used to fabricate the master mold using photolithography process with a negative near-

UV photoresist (SU-8 2050 Microchem Corp.). The Si wafer was spin-coated with the photoresist, then baked for solvent removal and exposed to UV light through the printed

mask. The exposed wafer was then post-baked and developed to obtain a master mold

containing the microstructures.

Once the mold fabrication was complete, trenches of PDMS were casted out of the SU-8

master mold by performing standard soft lithography method. PDMS prepolymer mixture

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(Sylgard184, DowCorning) was prepared from the elastomer base and curing agent at a

10:1 weight ratio. The mixture was then vacuumed to remove air bubbles. The silicon mold

was spin-coated with the mixture (100 µm thickness) and then cured at 60 ᵒC for 6 h.

Hardened PDMS with microstructures was peeled off the mold and cut into 5 µm by 15

µm pieces. Figure 3.10 shows an SEM image of the microfabricated trenches on PDMS sheet.

Figure 3.10. SEM image of 25 µm wide and 20 µm deep trenches on 100 µm thick PDMS sheet. Scale bar is 20 µm. Courtesy of Dr. Pooyan Tirandazi.

3.4.5 Fibril Preparation for SEM-DIC imaging

PDMS sheets were plasma cleaned and placed on a dish as shown in Figure 3.11. A 50 µL

volume of collagen fibrils was pipetted on an 8 mm coverglass that was placed beside the

PDMS sheet. A glass microneedle was used to draw collagen fibrils out of fibril solution.

Then using a second microneedle, fibrils were placed over trenches of PDMS. The

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microneedles were held by capillary holders and moved by electronic micromanipulators

(TransferMan NK 2, Eppendorf). Fibrils oriented northwest-southeast (perpendicular to the shear axis of the light path for an inverted microscope) were imaged in DIC microscopy

before SEM imaging.

Figure 3.11. A) The microscope stage equipped with micromanipulators. B) Schematic of pulling collagen fibrils out of fibril suspension using microneedles and placement of fibrils over trenches of PDMS.

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3.4.6 SEM Processing

PDMS sheets with collagen fibrils on them were placed on SEM stubs and coated with 5

nm of platinum using a sputter coater. Collagen fibrils were imaged using a Hitachi S-4800

at 1.5 kV. Images were captured with higher quality when samples were elevated as close

as possible to the upper detector.

3.4.7 Image Processing

The image processing to obtain fibril diameter from SEM images and DIC-EIS from DIC

images were performed in Matlab (Mathworks, Inc., Natick MA) as follows: SEM images were uploaded into Matlab. A region of interest (ROI) was defined around the fibril and edges of the fibril were found by defining a threshold intensity.

DIC images were uploaded into Matlab. An ROI was defined around the middle section of the fibril (10 µm) over trenches of PDMS. The maximum and the minimum intensities of all columns (across the fibril) were found and DIC-EIS was calculated as the difference between the maximum and the minimum intensities. This process was repeated for all frames (fibrils were scanned in z-direction with 0.1 µm steps) to find the best plane of focus.

TEM images were stitched together in Photoshop software to create large images of fibrils.

The large images were used to measure fibril diameter as a function of position along the fibril in ImageJ software.

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3.4.8 Statistical Information

All images were processed 5 times in MATLAB and results were reported as mean ±

standard deviation. ANOVA tests (P < 0.05) were used to indicate the consistency of image

processing (n=3) and also to indicate the consistency of DIC-EIS of hydrated fibrils in PBS

after 10 minutes, 30 minutes, 2 hours, 6 hours, 1 day, and 2 days. T-tests with P < 0.05 were used to show the statistical difference for DIC-EIS of fibrils in different orientation and also DIC-EIS of wet and dry fibrils.

3.5 Results

3.5.1 DIC-EIS is sensitive to collagen fibril diameter change

To measure the sensitivity of DIC-EIS to changes in fibril diameter, it was necessary to perform correlative TEM. Single collagen fibrils extracted from bovine sclera were placed on referenced TEM grids (Figure 3.9) and first imaged with DIC microscopy (Figure

3.12.A) oriented in northwest-southeast direction. Each fibril was then stained and imaged with TEM. TEM images were stitched together to produce large mosaics containing the full length of fibrils (Figure 3.12.B). For each fibril, continuous end-to-end measured fibril diameter was compared to the continuous end-to-end DIC-EIS representation of the fibril.

Figure 3.12.C shows a selected region of the fibril at high magnification. Measured diameters from this TEM image and their corresponding DIC-EIS are shown in Figure

3.12.D along the fibril section. Figure 3.12.D clearly demonstrates that DIC-EIS is sensitive to changes in collagen fibril diameter along the same fibril. However, the numerical results of DIC-EIS in our experimental setup was not comparable between

141 fibrils. This inconsistency was thought to be due to local distortion of formvar layer after fibril deposition which could alter the DIC-EIS signal.

Figure 3.12. A) DIC image of a representative fibril at 40x magnification. The fibril was oriented in northwest-southeast direction for DIC imaging. However, the image is rotated here for comparison with TEM image. B) TEM image of the same fibril. C) High magnification of a selected region of the fibril with a rapid diameter change. D) Correlation of DIC-EIS and diameter along the selected region. The x axis represents position along the fibril starting from the left side of the fibril.

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3.5.2 DIC-EIS is unaffected by small changes in fibril direction

Theoretically the maximum edge intensity happens when the fibrils are oriented in northwest-southeast direction since highest contrast is obtained in shearing direction. The vibration directions of analyser and polarizer and the resulting shear direction of Nikon

ECLIPSE TE2000 DIC inverted microscope is shown in Figure 3.13.

Figure 3.13. Shearing direction in Nikon ECLIPSE TE2000 DIC inverted microscope. From Nikon’s instructions manual.

It is possible to rotate the specimen or move the DIC prism [510-512] in order to achieve this maximum edge intensity. However, it is not always practical to image fibrils at this

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favorable angle. This can take considerable time and also not be suitable for imaging of

live systems. Therefore, we investigated the sensitivity of the DIC-EIS to fibril direction, while fibrils were rotated up to 90° away from northwest-southeast direction. As the fibril was rotating away from northwest-southeast direction, the change in DIC-EIS was more pronounced, lowering the DIC-EIS by 2%, 4%, 10%, 18%, and 34% for fibrils 15°-30°,

30°-45° ,45°-60°, 60°-75°, and 75°-90° deviated from the northwest-southeast direction, respectively. As shown in Figure 3.14, the DIC-EIS was not significantly altered for fibrils up to 45° deviated from the northwest-southeast direction. Data is normalized to max DIC-

EIS values (fibrils deviated 0°-15° from the northwest-southeast direction). Statistical analysis was performed first by testing equality of variances using F-test. Then significance of the data was tested using two-tail t-test assuming equal or unequal variances (P < 0.05).

Figure 3.14. Change of DIC-EIS as a function of fibril direction (n = 35 fibrils). Asterisks show statistical difference (P < 0.05).

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3.5.3 DIC-EIS is linearly correlated to collagen fibril diameter

Figure 3.15. DIC image of a fibril over the 25 μm wide gap. The middle section of the fibril was used to calculate DIC-EIS across the fibril. The scale bar is 10 µm.

Our goal was to calculate fibril diameter directly from DIC images. To overcome the inconsistency of DIC-EIS values of fibrils on TEM grids, fibrils were placed over trenches of PDMS to be imaged with SEM and DIC microscopy. The procedure was carried on the microscope stage (Figure 3.11.A) and single bovine sclera fibrils were pulled out of a fibril suspension using glass microneedles (Figure 3.11.B) and transferred to the PDMS sheet

(Figure 3.10). Fibrils were imaged by DIC (Figure 3.15) and then SEM (Figure 3.16). The middle section of each fibril (magnified portion of the fibril in Figure 3.16) was used to

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measure DIC-EIS and diameter. The DIC-EIS and fibril diameter were positively

correlated (0.97) for fibrils with diameter between ~100 to ~300 nm (n = 23 fibrils). Figure

3.17 shows this linear region. Using this linear curve, fibril’s diameter could be estimated

with ±11 nm accuracy. DIC-EIS was insensitive to diameter change for fibrils larger than

300 nm and smaller than 100 nm.

Figure 3.16. SEM image of a fibril over the 25 μm wide gap. The middle section of the fibril which was used to measure fibril diameter is shown in high magnification.

Note that due to high energy of focused beam of electrons, fibrils usually broke when images were captured repeatedly from the same fibril section (Figure 3.18). Therefore, each data point on Figure 3.17 represents the average diameter of one SEM image with no error bar. Also, the standard deviations of DIC-EIS data (148 AU in average) were too small to show on Figure 3.17.

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Figure 3.17. DIC-EIS as a function of fibril diameter (n = 30 fibrils). A linear region can be seen for fibrils with diameter between ~100 nm to ~300 nm (n = 23 fibrils). Note that diameter values are measurements of dehydrated fibrils with 5 nm platinum coating.

Figure 3.18. SEM image of a broken fibril due to high energy of focused beam of electrons. The broken fibril formed a spring-like structure. The scale bar is 1 µm.

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3.5.4 DIC-EIS is capable of live measurements of hydrated fibrils

The actual value of this method is its ability to look at live measurements. Therefore, we

correlated the dry to the wet DIC-EIS signal. Fibrils were placed over trenches of PDMS

and imaged dry and then in PBS with DIC. Five fibrils were imaged wet with predicted

diameters of 133, 155, 163, 204, and 289 nm (diameter of fibrils were predicted based on

the linear correlation that was found in Figure 3.17). Each fibril was hydrated and imaged

6 times in PBS: 10 minutes, 30 minutes, 2 hours, 6 hours, 1 day, and 2 days after adding

PBS. DIC-EIS of wet fibrils was not significantly different over these 6 sets of images

(ANOVA test, p < 0.05). Furthermore, DIC-EIS of all five wet fibrils were significantly different (T-test, p < 0.05) and showed a positive correlation (0.99) to fibril diameter

(Figure 3.19). Using this linear curve, fibril diameters could be estimated with ±4 nm accuracy.

Figure 3.19. Correlation of DIC-EIS for wet and dry fibrils. DIC-EIS of fibrils in PBS were also linearly correlated to fibril diameters. Y-axis error bars were smaller than 53 AU leading to less than ±2 nm uncertainty in predicted diameter (too small to show on the figure). An asterisk indicates a significant difference, p < 0.05, between conditions.

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3.5.5 DIC-EIS shows a reversible diameter change due to dehydration

Since dehydration and rehydration of fibrils over trenches of PDMS was destructive and fibrils usually broke under the process, DIC-EIS of fibrils on glass was used to investigate the reversibility of fibril diameter change after dehydration. Figure 3.20.A and B show DIC images of a typical fibril in dry and wet conditions with the same exposure time.

Figure 3.20. DIC images of a typical fibril in A) dry and B) wet conditions on glass. C) DIC-EIS of the fibril shown in A and B.

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Figure 3.20.C shows the sensitivity of DIC-EIS to diameter change for the fibril shown in

Figure 3.20.A and B. To investigate the reversibility of the signal, collagen fibril

suspension in PBS was added to the glass bottom dish and allowed for fibrils setting on the

glass. DIC images of fibrils were captured in PBS before drying, after air drying, and finally

after rehydration in PBS. DIC-EIS of four fibrils were analysed and the data shows a

reversible diameter change after drying and rehydration of fibrils (Figure 3.21). A

reversible diameter and D-banding periodicity change has been reported previously [513].

Figure 3.21. Reversible DIC-EIS of fibrils after air drying on glass.

3.6 Discussion

A powerful tool to measure absolute collagen fibril diameter from DIC-EIS through the use of correlative DIC-SEM and DIC-TEM imaging has been presented. The method is suitable for extracting information from fibrils with dry diameters between ~100 nm to

~300 nm.

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Collagen fibrils have diameter usually between 20 to 500 nm. Above 300 nm, we can begin

to image the fibril using any light microscopy method and so we don't need this technique.

The significant achievement is that using this technique we can extend the region over

which we can measure fibril diameter to a lower limit of 100 nm. DIC-EIS can be further

improved by using higher numerical aperture objectives and video-enhanced DIC to further extend the lower limit of measurement. This will allow quantitative studies of dynamic collagen fibril assembly, degradation, mechanics and mechanochemistry. While previously it was necessary to apply a secondary method to obtain fibril diameter [174], using this method direct DIC microscopy can be used to measure fibril diameter without the need to fix, stain, or label the fibril.

It is important to note that the DIC signal is sensitive to changes in experimental setup when the optical path is altered (e.g. changes in the index of refraction of fibril milieu, wavelength of light [514, 515], light exposure time, filters, and objective). However, given the generally linearity in the DIC-EIS vs diameter curve it can be easily calibrated with a few referenced fibrils.

When working in air, it is necessary to consider the water content of the fibrils, since it has been shown that the fibril diameter increases with hydration [454]. Fratzl and Daxer [516] have suggested a two-stage drying model for collagen fibrils in the corneal stroma in which first, water is removed from interfibrillar substance (mostly proteoglycans) and second, complete drying happens by removing water from inside the fibril. This suggests that fibril associated water is bound relatively strongly and that it affects the fibril diameter.

Measurements from electron microscopy of wet frozen [447] corneas and X-ray studies of hydrated corneas [517] showed a ~45% increase of diameter compared to dehydrated fibrils

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[518]. Another X-ray diffraction study of cornea showed ~40% increase of fibrils diameters on hydration [519]. We imaged dried and hydrated fibrils and showed that in both conditions, DIC-EIS is linearly correlated to the fibril diameter. We also showed that DIC-

EIS of hydrated fibrils were not changed over 2 days. These collagen fibrils were trypsin extracted [520] from bovine sclera and previously showed no proteoglycan content. Lack of proteoglycans can explain the rapid hydration of fibrils in PBS and also similar range of linearity in the DIC-EIS vs diameter curve (Figure 3.19).

As is well-known, the DIC signal is sensitive to fibril orientation showing maximum edge intensity when fibrils are oriented perpendicular to the shear direction. However, our data show that the DIC-EIS of fibrils oriented in different directions has an effect that is small at small angles and that the effect can be calibrated.

Fibril cross section shape needs to be considered carefully. We took our measurements on fibrils that were on formvar or stretched across trenches in a PDMS substrate. In the latter case, we could expect no surface effects on our measurements. However, it has been shown that fibrils can flatten when they are adhered to surfaces [521]. Flattening will strongly affect the measurements of DIC-EIS through three mechanisms: 1) The edge-to-edge distance will increase, 2) The presence of the second interface can change the gradient in the index of refraction, and 3) the curvature on the sides will change, altering the optical path gradient. While, DIC-EIS signals can be obtained from surface adhered fibrils, it is likely that there will be substantial shifting of the DIC-EIS/diameter curve.

Nonetheless, the method is only suitable for single fibrils. DIC microscopy is unable to visually differentiate between multiple small fibrils closely associated together and a larger single fibril. In case of a double-fibril that is shown in Figure 3.22, the DIC-EIS was not

152 appropriate to calculate the double-fibril diameter. This can be misleading and should be prevented through extraction and storage procedures.

Figure 3.22. Merged fibrils in A) DIC, B) SEM, and C) higher magnification of region shown B. Scale bars are 10 µm in A and B and 2 µm in C.

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4 EFFECT OF FIBRIL STRAIN ON FIBRIL/MONOMER ASSOCIATION

4.1 Introduction

In chapter 1, the role of mechanics on cell-collagenous matrix interaction and growth and

remodeling of collagenous structures in tissue levels was reviewed. It was indicated how

local tissue mechanical environment can affect growth and remodelling of collagenous

tissue. In this section, the possible mechanisms by which mechanics can alter growth and

remodeling of individual collagen fibrils are discussed.

4.1.1 Tensile forces on fibrils can regulate the dynamic equilibrium between molecular accretion and enzymatic degradation

During development, collagen fibrils are not yet entirely cross-linked [522] and are

dynamically subjected to degradation or growth [518, 523-525]. The collagen fibril

homeostasis is governed by a balance of monomer synthesis and fibril degradation.

Collagen monomers, present in subcritical concentration [199, 526, 527] in fibril’s milieu,

may dynamically associate to or dissociate from the developing fibril. Molecular

association has shown by Stopak et al. [40], where exogenous collagen incorporated into

connective tissue of developing embryonic chicken. Lu et al. (2018) [175] also showed

that fluorescently labeled collagen monomers assemble into extracellular collagen fibrils

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of cultured cells. However, the mechanism that guides molecular accretion and fibril

degradation and ultimately causes fibril growth is not clear.

It is known that, highly-stressed tissues like those in the rotator cuff show more collagen

remodeling compared with less stressed tissues like those in the forearm [528]. Indeed, several studies have shown that mechanical loading upregulates both collagen [529-531]

and enzyme [530] expression when structure formation is needed. Collagen fibrils are

initially crimped and respond to an applied force by straightening before stretching.

Enzymes bind to periodic buckling sites of relaxed collagen fibrils and degrade them [532].

It has been hypothesized that the fibril enzymatic degradation is an energy activated

process and the activation energy is increased by the axial strain energy density of the fibril

[533]. It has been shown that tensile stress not only increases the Gibbs free energy required for thermal denaturation [534, 535], but also inhibits enzyme degradation of the fibril [45,

46].

While the role of mechanics in inhibition of thermal and enzyme degradation has been

explained in the concept of ‘use it or lose it’ [536], still the exact mechanism by which mechanical stresses/strains can lead to fibril growth remains unclear. It is possible that the same mechanism that hinders the fibril degradation, also promotes the fibril growth by accelerating molecular accretion. We thus hypothesized that tensile forces on collagen fibrils can be viewed as a delicate mechanochemical switch [46] shifting the balance of collagen monomers association/dissociation at the surface of loaded fibrils.

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4.1.2 Tensile forces on fibril can regulate the thermodynamics of fibril growth reaction

Thermodynamically, collagen structure and stability are affected by collagen interaction

with water molecules: Collagen thermal stability increases by dehydration [194-198].

Molecules assembled in fibrils are more thermally stable than the molecules in solution

[199, 200], showing more than 23 °C higher denaturation temperature [201]. Besides,

collagen fibril formation and growth is an endothermic reaction, but entropy driven process

[90, 202] arising from release of water molecules into the surrounding liquid [203, 204].

The rate of fibril formation and growth increases with temperature up to the denaturation

temperature of the collagen molecules.

In general, breakers of water structure promote collagen fibril formation, while makers of

water structure are inhibitory [205]. Mature fibrils in vivo are cross-linked by covalent

bonds between neighbouring molecules. However, the young and growing fibrils are

stabilized by non-covalent hydrogen bonds [206] and have the potential to bind more water molecules [196]. Collagen structural models [537-541] suggest that there are two types of

intermolecular and intramolecular hydrogen bonds in fibrils: I) a direct interchain hydrogen bond forms between the glycine residue and the residue in the second position of the

neighbouring chain (Figure 4.1.a), and II) an additional hydrogen bond which links two

adjacent tropocollagens using a bridging water molecule (Figure 4.1.b). This water-

mediated hydrogen bonding makes two thirds of hydrogen bonds that connect

neighbouring peptides [542] and therefore is a dominant interaction in stabilizing the fibrillar structure [543, 544]. These water bridges are dynamically linked with freely exchangeable hydrogen atoms [545].

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Figure 4.1. (a) A direct intermolecular hydrogen bond. (b) A bridging water molecule between polar groups of two tropocollagens. (c) Water molecule trapped between two hydrophobic side chains of neighbouring tropocollagens. Reprinted with permission from Streeter and Leeuw (2011) [203].

Furthermore, water molecules can be sandwiched in between hydrophobic groups of neighbouring tropocollagens [546] to maximize the number of water-water hydrogen bonds [547, 548] (Figure 4.1.c). Since molecular assembly is driven by decreasing the number of unfulfilled hydrogen-binding opportunities at the protein-water interface [549],

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the trapped water molecules and the water bridges may have an important role in the

collagen molecular assembly during fibril growth and remodeling.

Mechanical load on fibrils could in principle change the molecular conformation of fibrils

and release these water molecules. This tension-induced mechanism could provide the increase of entropy required for association of molecules into the fibrils. Polymer mechanochemistry has shown that mechanical forces can lower the reaction barrier of mechanochemical processes at molecular level and facilitate a number of otherwise

kinetically inaccessible processes [550, 551]. The effect of mechanobiological factors on

molecular association/dissociation with collagen fibrils may be interpreted as a

consequence of their effects on water-structure [28].

4.2 Experimental Methods

Our goal was to assess the role of fibril strain on the incorporation rate of collagen

monomers onto fibrils. Briefly, fibrils were extracted from bovine sclera and then stretched to 0%, 4%, and 6% strain and kept under static strain using two microneedles. Subthreshold concentration of labeled monomers were added to the stretched fibril’s medium and the fibril’s fluorescent intensity was recorded and analysed.

4.2.1 Fibril Attachment Between Microneedles

A micropipette puller (Figure 4.2, #P-97, Sutter Instrument, Novato, CA) and borosilicate

glass rods (10 cm long and 1 mm in diameter) were used to create microneedles with fine

tips (Figure 4.3).

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Figure 4.2. P-97 Sutter Instrument micropipette puller. From www.sutter.com.

Type I collagen fibrils were extracted from bovine scleras as previously described in

Chapter 2. A delta T dish was plasma cleaned and placed on the microscope stage as shown previously in Figure 3.9 and Figure 3.11. To keep the dish clean and free of unwanted collagen fibrils during the experiment, an 8 mm cover glass was placed on the dish and a

75 µL volume of fibril suspension was pipetted on the 8 mm coverglass. Two microneedles were positioned above the fibril suspension using two Eppendorf Transferman NK2 micromanipulators (Figure 3.11).

Figure 4.3. A representative microneedle that was used to stretch fibrils.

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A fibril, around 200 µm long was chosen near the droplet surface. One of the needles was placed under and near one end of the fibril. Using this needle, the fibril was drawn from

the fibril suspension, but the other end was kept inside the droplet. The second microneedle was used to grab the other end of the fibril that was still in the droplet. Once the fibril was attached to both microneedles, it was moved away from the droplet surface. The fibril suspension and the 8 mm coverglass were carefully removed from the dish.

Figure 4.4. A fibril that was drawn from the fibril suspension and wrapped around microneedles in air and hold slightly tight to adhere to the glass.

Then, the fibril was wrapped around the needles by rotating one needle around the other one at least 3 times. During the process, the fibril was kept slightly tight to facilitate

bonding of the fibril to the glass by the force of adhesion. The fibril was kept slightly tight

for another 1 hour to completely adhere to the glass (Figure 4.4). It was made sure to have

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the fibril tight enough to adhere to the needles, but not too tight to deflect the fine tip of the microneedles. After the fibril was completely air dried, a sheet of trenches of PDMS was placed on the dish. The fibril was positioned over a trench while attached between

microneedles (Figure 4.5). DIC images were captured to estimate fibril diameter as

explained in the previous chapter.

Figure 4.5. DIC image of a fibril over trenches of PDMS. After the fibril was wrapped around microneedles, it was moved over trenches of PDMS to measure DIC-EIS and estimate its diameter.

4.2.2 Fibril Transfer into PBS, Preconditioning, and stretching

The fibril was then raised to remove the PDMS sheet and the 8 mm cover glass from the

dish. Then, the dish was filled with PBS and the fibril was gently lowered into the PBS.

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The temperature was set at 35 °C and water was added to the dish using a pump to count for evaporation during the experiment. The fibril was kept in PBS for 30 minutes to allow for complete rehydration.

Then, using the two microneedles, fibrils were pulled to 0%, 4%, or 6% strain. These fibrils usually had some kinks or twists which could open after small amount of stretching.

Therefore, fibrils preconditioned to prevent immediate stress relaxation and increase of fibril initial length after straightening the initial twists. The preconditioning was performed by stretching fibrils 3 times to 0%, 4%, or 6% strain. At the beginning of each cycle, fibrils were stretched to the point that it caused deflection in at least one of the microneedles. The fibril length (distance between the two microneedles) at this stage was recorded as the fibril initial length for that cycle. At all times, one microneedle was kept stationary and the other one moved 1 µm/s manually to stretch or relax the fibrils.

At the end of the third cycle, fibrils were held under static strain (0%, 4%, or 6% strain).

Note that preliminary results indicated that holding fibrils at 10% strain and higher caused fibril failure in under 1 hour. Therefore, it was decided to test fibrils under 3 conditions: no load (0% strain), low load (4% strain), and high load (8% strain). However, after designing the experiment and performing some conditions, the 8% strain condition was no more stable during the whole length of the experiment. Therefore, the high load condition was changed to 6% strain. Figure 4.6 shows two representative fibrils that were kept under 0%

(panel A) and 6% strain (panel B). Note that stress relaxation was still pronounced for the fibrils being stretched to 4% or 6% strain. The relaxation was noticeable by the change in microneedles deflection.

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Figure 4.6. Fibrils secured between needles inside PBS. The fibril in the panel A is tight, but not loaded (0% strain). The fibril in the panel B is stretched to 6% strain. Note the needles deflection in the panel B. Also, the coiled section of the fibril can be seen on the top needle in the panel A.

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4.2.3 Label monomer addition and recording

After fibrils were hold at 0%, 4%, or 6% strain, labeled monomers (DOL=0.6) were added

to the dish at a final concentration of 0.5 µg/mL and mixed well by pipetting 1 mL of the

solution up and down for 10 times. DIC (10 ms exposure time) and Fluorescent (5 s

exposure time) images were captured every 1 minute for 1 hour on a Nikon inverted

microscope (ECLIPSE TE2000-E) equipped with a CoolSNAP EZ CCD Camera and a

40X objective (Plan Fluor ELWD 40x Nikon, NA 0.6). Labeled monomers were added and

mixed inside the dish between the first and second frames.

4.2.4 Image Processing and Assessment of Incorporation Rate

DIC images were processed to measure DIC-EIS and estimate fibril diameter as explained in the previous chapter. Fluorescent images were processed in MATLAB as following: An

ROI was defined around the middle section of the fibril. In some cases, there were some artefacts on the middle part of the fibril. Therefore, the ROI was defined avoiding the artefact. A background ROI was also defined adjacent to the fibril to measure the fluorescent intensity of the background. Fluorescent intensity of the background ROI was scaled to the size of the first ROI and then subtracted from it. The background subtracted fibril intensity was then normalized per surface area of 1 µm length of the fibril.

The incorporation rate was calculated based on the fibril fluorescent intensity. Results showed that total fibril intensity rapidly increased during the first 30 minutes, followed by a plateau. Therefore, it was assumed that Kon (association rate) >> Koff (dissociation rate)

in the first 30 minutes.

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Furthermore, two photobleaching functions were assumed for labeled monomers:

photobleaching of monomers 1) on the fibril and 2) in the solution. Since monomers on the

fibril are directly exposed to the exciting light, the data provided by the AF488

manufacturer was used for their photobleaching (Figure 4.7). While, photobleaching of the background ROI was considered as the photobleaching of the monomers in the solution.

Figure 4.7. Alexa Fluor 488 photobleaching. The data is extrapolated from the manufacturer website.

It was assumed that the total concentration of monomers in the dish is constant during the experiment. A simple calculation shows that there are ~3*1012 collagen molecules in a dish

when it is filed with 3 mL of 0.5 µg/mL collagen solution. Also it would take ~140000

monomers to add 1 layer of monomers on a fibril’s surface which is 200 nm in diameter

and 100 µm long. One layer of monomers corresponds to ~3 nm fibril diameter increase.

Therefore, it would be reasonable to assume constant soluble monomer concentration in

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the dish during the experiment. Note that there was only one fibril in the dish and also no

other new microfibrils were formed away from the tested fibrils. It was also assumed that

after the initial mixing, the collagen concentration remained uniform throughout the dish.

The diffusion coefficient of collagen molecules in a free and dilute solution is ~8 µm2/s

[552].

The fibril intensity at each time point (In) was considered as the sum of the intensity of fibril at the previous time point in addition to the intensity of new monomers added to the fibril at the current time point (Kn). In addition, all monomers carry their photobleaching

history. Meaning that if a monomer is added to the fibril at the time point of ‘n’, it has been photobleached in the solution ‘n-1’ times and will be photobleached on the fibril after that.

This has been formulated as:

= + + + +

𝐼𝐼𝑛𝑛 𝐾𝐾1𝐽𝐽1𝐹𝐹𝑛𝑛 𝐾𝐾2𝐽𝐽2𝐹𝐹𝑛𝑛−1 𝐾𝐾3𝐽𝐽3𝐹𝐹𝑛𝑛−2 ⋯ 𝐾𝐾𝑛𝑛𝐽𝐽𝑛𝑛𝐹𝐹1 Where ‘J’ is the photobleaching function for the monomers in the solution and ‘F’ is the

photobleaching function for the monomers on the fibril. A custom MATLAB code was

used to solve for Kn.

4.2.5 Control Test and Long-Term Experiments

Alternatively, some fibrils were held under strain for 3 hours in the presence of labeled

monomers. Furthermore, as a control test to examine the effect of handling fibrils using

microneedles, some fibrils were allowed to sink onto the glass and exposed to the same

concentration of labeled monomers for 3 hours.

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4.3 Results

4.3.1 Labeled Monomers Incorporate onto the Fibrils

Bovine sclera fibrils were stretched to 0%, 4%, and 6% strain (n = 3 fibrils for each condition) and incubated in 0.5 µg/mL fluorescently labeled collagen molecules at 35 °C

in a temperature controlled open dish. For all tested fibrils, the fibril intensity was increased rapidly and reached a plateau during the first 30 minutes (Figure 4.8).

Figure 4.8. Normalized fibril intensity. The fibrils’ intensity rapidly increased and reached a plateau during the first 30 minutes. Fibrils which were under 4% and 6% strain reached their maximum intensity significantly faster than unloaded fibrils (t-test, p < 0.05).

Figure 4.9 shows a representative fibril before adding labeled monomers and 30 minutes after adding labeled monomers. All fibrils showed random bright punctuated spots which were unavoidable in the fibril analysis and selection of regions of interests. Therefore, the

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absolute values of fibril intensity per fibril’s surface area were not comparable between the

tested fibrils regardless of being stretched or not. However, normalizing each fibril’s

intensity by its maximum intensity at the plateau showed that the fibrils which were under

4% or 6% strain reached their maximum intensity faster than the 0% strain fibrils. The plateau times (time to reach 90% of the maximum intensity) were 15.6 ± 5.3, 7.0 ± 1.7, and

6.0 ± 2.4 for fibrils under 0%, 4%, and 6% strain, respectively. ANOVA test (p < 0.05) showed that the means of the three populations are not all equal (F-value = 8.15 and F- critical = 3.98). T-tests were used to test each pair of means. T-tests (p < 0.05) showed that the plateau times were significantly different between 0%-4% and 0%-6%, but not between

4%-6% groups.

Figure 4.9. DIC and fluorescent images of a representative fibril stretched to 6% strain between needles. Panel A (DIC) and B (fluorescent) show the fibril at time 0, right before adding the labeled monomers. Panel C (DIC) and D (fluorescent) show the fibril at 30 minutes after adding the labeled monomers. Note the punctuated bright spots on the fibril after 30 minutes in panel D.

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Exponential curves were fitted to the normalized fibril intensity data. The fitted curves

predict that fibrils reach their maximum intensity after 30.0, 12.8, and 10.6 minutes for

fibrils under 0%, 4%, and 6% strain, respectively. Figure 4.10 clearly demonstrates that fibrils under 4% and 6% strain reach their maximum intensity before the unstretched fibrils.

Figure 4.10. Exponential fitted curves on the normalized fibril intensity data. The dash lines represent the 95% prediction intervals.

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4.3.2 Microfibrils Form on the Fibrils between Microneedles

After 30 minutes, microfibrils were formed and continued to grow (at least up to 3 hours)

on the tested fibrils between needles (Figure 4.11). Therefore, the fibrils’ intensity was not predictable after 30 minutes due to presence of microfibrils in the selected ROIs.

Alternatively, when fibrils were allowed to sink on the glass bottom dish, air dried,

rehydrated in PBS, and incubated in 0.5 µg/mL labeled collagen at 35 °C, no microfibrils

were seen in association with tested fibrils at least up to 3 hours.

Figure 4.11. Formation of microfibrils around the tested fibrils between microneedles.

4.3.3 Incorporation rate onto 6% strained fibrils remained significantly higher than 0% and 4% strained fibrils

It was assumed that during the first 30 minutes, the Kon (association rate) >> Koff

(dissociation rate). Considering the photobleaching of labeled monomers in the solution

and on the fibrils, incorporation of new labeled monomers at each time point was estimated

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as explained in section 4.2.4 and normalized in Figure 4.12 to compare between groups.

All fibrils showed their highest monomer incorporation immediately after addition of

labeled monomers. Incorporation rapidly decreased during the first 10 minutes, and

remained constant afterward until 30 minutes. However, the fibrils under 6% strain

maintained a higher incorporation after 10 minutes. Normalized incorporation remained

fairly constant after 10 minutes at 0.26 ± 0.03, 0.25 ± 0.04, and 0.49 ± 0.15 for fibrils under

0%, 4%, and 6% strain, respectively. ANOVA test (p < 0.05) showed that the means of the

three populations are not all equal (F-value = 8.20 and F-critical = 3.98). T-tests were used to test each pair of means. T-tests (p < 0.05) showed that the incorporation onto 6% strained fibrils remained significantly higher than 0% and 4% strained fibrils after 10 minutes.

Figure 4.12. Labeled monomers incorporation onto fibrils. While all fibrils started with their highest monomer incorporation, the normalized incorporation onto 6% strained fibrils remained significantly higher than 0% and 4% strained fibrils.

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Figure 4.13. Exponential fitted curves on the normalized labeled monomer incorporation data. The dash lines represent the 95% prediction intervals.

Exponential curves were fitted to the normalized labeled monomer incorporation data

(Figure 4.13). The fitted curves predict that fibrils under 6% strain have a higher

incorporation rate compared to fibrils under 0% and 4% strain, throughout the experiment.

The normalized incorporation values were obtained from the fitted curves at 5, 10, and 30 minutes and are shown in Table 4.1.

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Table 4.1. Normalized incorporation values obtained from the fitted curves at 5, 10, and 30 minutes.

4.4 Discussions

A simple method was developed to mechanically stretch single native fibrils using only microneedles and micromanipulators. In combination with 1) the single molecule fluorescent labeling method and 2) the fibril diameter estimation method by DIC-EIS, we will be able to ask fundamental questions on how mechanics can affect fibril growth and remodeling. Here, we investigated the effect of static loading on the growth of native fibrils. Fibrils were stretched to 0%, 4%, and 6% strain. It was shown that labeled monomers rapidly incorporate onto all tested fibrils. However, the association rate remained significantly higher for fibrils under 6% strain compare to fibrils under 0% and

4% strain.

In our experimental setup, due to formation of microfibrils, the fibril intensity and consequently the monomers incorporation were not quantifiable after 30 minutes. Before

30 minutes, it was assumed that the dissociation rate is negligible and therefore fibril intensity is due to only labeled monomers incorporation and their photobleaching in the solution and on the fibril. It is expected to start with a high incorporation rate to replace the monomers that were lost during the fibril extraction and experimental procedure. However, to maintain a constant diameter, the association and dissociation rates have to become equal

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after reaching equilibrium. Therefore, as incubation time increases, the dissociation rate

should increase. That means our results underestimate the incorporation rate towards the

end of 30 minutes.

A simple calculation shows that there are ~3*1012 collagen molecules in a dish when it is

filed with 3 mL of 0.5 µg/mL collagen solution. It would take ~140000 monomers to add

1 layer of monomers on a fibril’s surface (200 nm in diameter and 100 µm long). One layer

of monomers corresponds to ~3 nm fibril diameter increase. Also, the diffusion coefficient

of collagen molecules in a free and dilute solution is ~8 µm2/s [552]. Therefore, it was concluded that the concentration of labeled monomers around the tested fibrils remains constant during the experiment. Consequently, the higher association rate of fibrils under

6% strain supports the structural limited growth models for the fibril growth and shows that mechanical strain can put fibrils in a lower state of energy and stimulate fibril growth.

Incorporation of labeled monomers onto the fibrils under 0% and 4% strain were not

significantly different. This could be due to mechanical tension experienced by all fibrils

during the experimental procedure (dehydration, submerging into PBS, and being wrapped around needles). Furthermore, the experiments were not completely strain-controlled. The

fibrils that started under 4% or 6% strain, partially relaxed during the experiment. This

could minimize the difference between the 0% and 4% experimental group and lead to

similar incorporation rate.

Formation of microfibrils on all the fibrils tested between microneedles, but not on the fibrils on the glass-bottom dish, show that the experimental procedure causes permanent changes on the fibrils between needles and decreases the activation energy required for nucleation of new microfibrils on the fibrils between needles. Both fibrils, on the glass and

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between needles, were extracted with the same method and dehydrated and rehydrated

during the experimental procedure. The only difference was that the fibrils between needles were exposed to some inevitable mechanical tension (even the 0% strain group). Therefore, this change in activation energy has to be due to applied mechanical tension on the fibrils and no other possibilities such as dehydration. Furthermore, the punctuated bright spots showed in Figure 4.9 could be the damaged areas on the fibril. These damaged spots could be the potential site of monomers incorporation and microfibrils nucleation. It has been shown previously by Holmes et al. (2010) [294] that fractured ends of fibrils could grow by molecular accretion.

Therefore, mechanical stresses and strains could increase fibril growth by two mechanisms.

First, partial damage in fibril structure decreases the activation energy required for reaction

between free monomers in solution (subthreshold concentration) and fibrils. Formation of

punctuated bright spots and microfibrils on the tested fibrils support this conclusion. The

required activation energy could also decrease in vivo by loss of surface bound

proteoglycans or N-propeptides. Second, mechanical strains could put fibrils in a lower

state of energy, increasing the association rate of free monomers and fibrils. Fibrils can take a lower state of energy by either loss of bound water molecules [205] or molecular twisting within fibril [25].

Finally, the results support the growth and remodeling theory [43] that has been formulated in Dr. Ruberti’s research group. It suggests that collagen monomers and enzymes are in a dynamic equilibrium with existing fibrils and tensile forces on fibrils can shift this

equilibrium and change the balance between molecular association (kon) and dissociation

(koff). It has been previously shown in our laboratory that 1) tension can directly drive initial

175

fibrillogenesis and structures into the path of force [42], 2) applied mechanical strain preferentially preserves type I collagen against enzymatic degradation in molecule [44],

fibril [45, 46], and tissue [47-49] levels, 3) in the absence of cells, unloading of fibers can

lead to disassociation of collagen fibers [42], 4) and cyclic stress leads to strengthening of

the collagenous structure [50]. All together suggest that application of mechanical forces

can induce fibril formation, strengthen fibrils, and decrease molecular dissociation rate.

Here we showed that stresses and strains on collagen fibrils can promote the molecular

assembly and fibril growth. The results elucidate the role that mechanics could play in the

collagen fibrils growth mechanism and provide new opportunities to develop treatments

for variety of pathological conditions by promoting growth or degradation when/where

needed.

4.5 Summary and Conclusions

In this PhD dissertation, we sought to answer this question: Does fibril strain promote the

molecular assembly of collagen? To investigate this question, individual collagen fibrils

were manipulated and stretched between two microneedles. The stretched fibrils were

exposed to a subthreshold concentration of fluorescently labeled collagen molecules to

quantify molecular association onto the stretched fibrils. To achieve this, it was necessary to 1) fluorescently label individual collagen monomers while minimally disturbing their fibril forming functionality and 2) accurately measure the diameter of tested fibrils.

In chapter 2, to address the observation-interference problem, exogenous collagen

molecules were tagged with multiple, small fluorophores and the effect on morphology and

kinetics was examined. While the labeling procedure could alter fibrillogenesis kinetics, it

176 was shown that adding two labels (permitting orientation measurement) to each molecule and keeping the percentage of labeled monomers below 5% of the added collagen, preserved both the self-assembly kinetics and the fibril morphology. Exposure of native young collagen fibrils to labelled collagen demonstrated that exogenous molecules associate with and likely incorporate into the fibrils at low, but measurable rates. The incorporation rate and total accumulation (to reach saturation) of labeled collagen into the scleral fibrils (predominantly type I collagen) was measured as 2.8 ± 1.2 and 4.0 ± 1.5 molecules/(µm2.minute) and 114.3 ± 26.8 and 207.1 ± 55.4 molecules/µm2 at 25 and 30

°C, respectively. The reaction of the monomers with the fibril surfaces produced a kinetic signature similar to de novo collagen assembly, with an activation energy of 12.6 kcal/mol.

We further showed that at least 40% of the labelled collagens will have detectable orientation to determine whether exogenous collagen is associating non-specifically or functionally with the fibrils.

In chapter 3, a method using a simple standard optical microscopy technique, DIC, to measure fibril diameter has been developed and calibrated. The theoretical limit of visible light microscopy is approximately 220 nm. This resolution limit does not permit very accurate determination of fibril diameter or changes in diameter. We were able to demonstrate a direct, linear correlation between fibril diameter and the DIC-EIS. Using a non-oil immersion, 40x objective (NA 0.6), collagen fibril diameters between ~100 nm to

~300 nm could be obtained with ±11 and ±4 nm accuracy for dehydrated and hydrated fibrils, respectively. Above 300 nm, diameters can be determined using conventional imaging techniques and the lower limit of detection can be decreased by using a higher NA objective. This simple, non-destructive, label free method should advance our ability to

177

directly examine fibril dynamics under experimental conditions that are physiologically

relevant.

In chapter 4, the effect of static loading on the growth of native fibrils was investigated.

Fibrils were stretched to 0%, 4%, and 6% strain and exposed to subthreshold concentration of fluorescently labeled collagen molecules. It was shown that labeled monomers rapidly incorporate onto all tested fibrils and reach a plateau. The time to reach plateau was significantly faster for the stretched fibrils. Analysis of the fibril intensity and photobleaching data indicated that the association rate was significantly higher for fibrils under 6% strain compared to fibrils under 0% and 4% strain, increasing the association rate by 100%. It was concluded that mechanical stresses and strains could increase fibril growth by decreasing the activation energy required for reaction between monomers and fibrils

and also by setting fibrils in a lower state of energy, increasing the association rate of

monomers and fibrils.

Understanding the very local mechanism of collagen fibril growth and remodeling could

indeed shed light on the biomechanics of collagenous tissue. Molecular mechanistic insight

gained from these studies will likely provide new opportunities to develop treatments of a

broad range of ECM-associated pathological conditions by promoting growth or

degradation when/where needed.

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