Building a Synthetic Periodontal Ligament: Collagen

BUILDING A SYNTHETIC PERIODONTAL LIGAMENT: COLLAGEN

NANOSTRUCTURES ON TITANIUM

ELOISE PEYTON MILLER

Submitted in partial fulfillment of the requirements

For the degree of Master of Science

Thesis Advisor: Dr. Steven Eppell

Department of Biomedical Engineering

CASE WESTERN RESERVE UNIVERSITY

May 2020

CASE WESTERN RESERVE UNIVERSITY SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

Eloise Miller candidate for the degree of Master of Science in Biomedical Engineering*.

Committee Chair Steven Eppell

Committee Member

Anirban Sen Gupta

Committee Member

Leena Palomo

Committee Member

Jonathan Pokorski

Date of Defense

March 10, 2020

*We also certify that written approval has been obtained for any proprietary material contained therein.

Table of Contents

Table of Contents ...... iii

List of Tables ...... vii

List of Figures ...... viii

Abstract ...... 1

Chapter 1: Introduction and Background ...... 2

1.1 Dental Implants ...... 2 1.2 Periodontal Ligament ...... 5 1.3 Artificial Ligament Systems ...... 8 1.4 Collagen Self-Assembly: Fibrillogenisis ...... 10 1.5 Non-Specific Covalent Bonds Between Proteins and Surfaces ...... 14 1.6 Carbonyl Formation and Oxyamine Conjugation ...... 15

Pyridoxal-5-Phosphate as a Transamination Reagent ...... 15 Rapoport’s Salt as a Transamination Reagent ...... 25 Summary of Carbonyl Formation and Oxyamine Conjugation...... 26

1.7 Silane Surface Reactions ...... 27

Chapter 2: Study Design ...... 29

2.1 Study Goal ...... 29 2.2 Experimental Design ...... 30 2.3 Developing the Experimental Protocol ...... 31

Step 1: Silanization of Titanium ...... 31 Step 2: Ketone-Collagen Production...... 42 Step 3: Binding Ketone-Collagen to Oxyamine-Silanized Titanium .... 55 Step 4: Nucleation and Growth of Collagen Fibrils on Titanium ...... 60 Summary of Developed Protocol ...... 65

Chapter 3: Manuscript ...... 67

iii

3.1 Preface ...... 67 3.2 Manuscript ...... 67

3.2.1 Abstract ...... 67 3.2.2 Introduction ...... 68 3.2.3 Results and Discussion ...... 70 3.2.4 Conclusions ...... 79 3.2.5 Methods ...... 80

Chapter 4: Conclusions and Future Work ...... 84

4.1 Conclusions ...... 84 4.2 Future Work ...... 85

Loading Fibril Surfaces ...... 87 Animal Modeling ...... 89

4.3 Other Potential Applications ...... 91

Space-safe collagen adhesive ...... 91 Dynamic Testing of Fibrils Synthesized In-vitro ...... 92

Protocols ...... 94

Titanium Slides...... 94

Necessary Components ...... 94 Protocol for obtaining titanium squares: ...... 95 To remove titanium from holders ...... 95

Cleaning Procedures ...... 95

Set up the Sonicator ...... 96 Sonicating Titanium ...... 96 Washing holders with soapy water ...... 98 Sonicating Holders ...... 98

Protocols – Chemical Steps to Produce Fibrils on a Titanium Surface ..... 100

Ketone-Collagen ...... 100

Plasma Treatment...... 106 Oxyamine Surfaces ...... 107 Oxyamine-Ketone Collagen Surfaces (oxime linkage reaction) ...... 112 Nucleation and Growth of fibrils (fibrillogenesis) ...... 114

Buffers and Rinsing Solutions...... 116

Safety...... 116 Transamination MES Buffer: pH 6.5 with NaCl ...... 117

Oxime Linkage MES Buffer: pH 5.5 with MgCl2 ...... 118

KeCol Rinsing Solution: 150 mM MgCl2 (“ketone-collagen”)...... 119 PBS: Phosphate Buffered Saline...... 120

Determining Efficacy ...... 120

UV-Vis Spectroscopy: Oxyamine-TAMRA reaction and analysis ..... 120 XPS ...... 124 Immunostaining: Immunofluorescent Imaging ...... 126 SEM ...... 130

3D-Printed Holders ...... 133

Ordering Oxyamine Molecule ...... 135

Use of Selected Tools ...... 136

Using the Scale ...... 136

Necessary Components: ...... 136 Protocol: ...... 136

Using the pH Meter ...... 137

Calibration ...... 138 Use ...... 139

Squeeze Bottles ...... 139

Soap ...... 140 Alcohols – Methanol, ethanol, isopropanol ...... 140

Waste – How to dispose of used solvents and chemicals ...... 140

Alcohol used for cleaning...... 140 Oxyamine-contaminated solvent ...... 141 Acetic acid buffer, PBS, collagen, ketone collagen, soapy water, water ...... 141

UV-Vis Spectroscopy, NanoDrop 1000 ...... 142

Why not use a Bradford Assay? ...... 143 Sample Preparation ...... 143

Concentration for NanoDrop ...... 143 Dilution for NanoDrop ...... 143

Using the Nanodrop 1000 ...... 144

ImageJ processing of Immunofluorescence images ...... 150

ImmunoFluorBrightness.m ...... 152

Plasma Cleaner Operation ...... 154

Necessary Components: ...... 154 Notes: ...... 154 Before you begin: ...... 156 Protocol: ...... 157 Troubleshooting:...... 160

Bibliography: ...... 163

List of Tables

Table 1: Solution conditions that lead to the formation of collagen aggregates

that are not D-banded fibrils...... 13

Table 2: Results of a study using PLP as a transamination agent on various

proteins...... 19

Table 3: Results of 20-tetrapeptide library screening for N-terminal residue PLP activity...... 21

Table 4: Bond energies, lengths, and energy-to-length ratios of covalent bonds in

the synthetic ligament system ...... 88

vii

List of Figures

Figure 1.1: Simple periodontal ligament...... 5

Figure 1.2: Structure of a collagen fibril...... 6

Figure 1.3: Cellular and vascular periodontal ligament...... 7

Figure 1.4: Typical Turbidity vs. Time profile of collagen self-assembly...... 11

Figure 1.5: Transamination and Oxime Formation scheme...... 15

Figure 1.6: Amine-catalyzed silane surface reactions...... 28

Figure 2.1: Simple overview of the structure of a single collagen fibril bound to a

titanium surface...... 30

Figure 2.2: Amine-carbonyl reaction with cyanoborohydride reduction...... 32

Figure 2.3: Assessing plasma treatment by hydrophilicity...... 34

Figure 2.4: XPS spectra of a titanium slide exposed to hydrolyzed APTES (top

purple spectrum) and a titanium slide exposed to non-hydrolyzed APTES (bottom green

spectrum...... 35

Figure 2.5: 3D-printed fluid exchange holders...... 37

Figure 2.6: Clockwise diagram showing the fluid exchange process with 3D- printed holder cross-sections...... 38

Figure 2.7: XPS spectra of APTES titanium slides with positive results...... 39

Figure 2.8: XPS spectra of a clean titanium slide without surface modification

(bottom blue spectrum) and a titanium slide modified by surface addition of oxyamine-

silanes (top red spectrum)...... 40

Figure 2.9: Scheme of oxyamine-silane surface chemistry...... 41

Figure 2.10: FTIR spectra of titanium slides...... 44

viii

Figure 2.11: FTIR spectrum of a titanium surface with a dried spot of PureCol

collagen...... 45

Figure 2.12: PLP gel mass...... 46

Figure 2.13: PLP gels on a pipette tip...... 46

Figure 2.14: Chemical structure of pyridoxal 5'-phosphate (PLP)...... 48

Figure 2.15: Chemical structure of N-methylpyridinium-4-carboxaldehyde

benzosulfate salt, A.K.A. Rapoport's salt...... 49

Figure 2.16: SDS-collagen gel...... 49

Figure 2.17: Chemical structure of 3-(1-pyridinio)-1-propanesulfonate, a “bulky” salt with a similar chemical structure to RS...... 50

Figure 2.18: Conversion of N-terminal amines of collagen using N-

Methylpyridinium-4-carboxaldehyde as a transamination agent...... 53

Figure 2.19: UV-vis spectra of collagen solutions...... 54

Figure 2.20: Oxime linkage formation between surface-bound oxyamine-silane

molecules and ketone-modified collagen N-terminus...... 55

Figure 2.21: Histograms of immunofluorescence grayscale values for the

detection of collagen molecules on activated titanium surfaces...... 56

Figure 2.22: Histograms of fluorescent intensity found on the acid-hydrolyzed slides...... 59

Figure 2.23: SEM image of a collagen gel on a titanium slide formed using a

standard fibrillogenesis procedure...... 61

Figure 2.24: SEM images of titanium surfaces...... 63

Figure 2.25: SEM images of zoomed-in titanium surfaces...... 64

Figure 2.26: Overview of the chemical steps necessary to produce collagen

fibrils conjugated end-on to a titanium surface...... 66

Figure 3.1: Overview of the chemical steps necessary to produce collagen fibrils

conjugated end-on to a titanium surface...... 70

Figure 3.2: XPS spectra of a clean titanium slide without surface modification

(bottom blue spectrum) and a titanium slide modified by surface addition of oxyamine-

silanes (top red spectrum)...... 72

Figure 3.3: UV-vis spectra of collagen solutions...... 73

Figure 3.4: Histograms of immunofluorescence grayscale values for the

detection of collagen molecules on activated titanium surfaces...... 76

Figure 3.5: SEM images of titanium surfaces...... 78

Figure 3.6: SEM images of zoomed-in titanium surfaces...... 79

Figure 4.1: Chemical structure of surface-bound ketone collagen molecules...... 87

Figure 4.2: Diagram of bio-inspired adhesive using two titanium pieces functionalized with collagen fibrils...... 91

Building a Synthetic Periodontal Ligament: Collagen Nanostructures on

Titanium

Abstract

ELOISE PEYTON MILLER

Clinical complications revolving around dental implants present significant clinical

problems. Many of these complications, such as infection and aesthetic issues, could be remedied by the addition of an artificial biomimetic periodontal ligament at the interface of the titanium implant and the alveolar bone. This study seeks to mimic the natural structure of the periodontal ligament found around natural teeth roots by conjugating collagen fibrils perpendicularly to a titanium surface. The results of this study show proof-of-concept that collagen molecules can be bound end-on to a titanium surface,

laying the groundwork for the development of artificial periodontal ligament. Further

development of the ligament-building protocol is necessary to produce a bio-functional

tissue.

Chapter 1: Introduction and Background

1.1 Dental Implants

With over 2 million dental implants placed annually in the United States,1 dental

implants have become a common alternative to fixed and removable partial dentures for

the treatment of tooth loss.2 A two-piece dental implant is comprised of three parts: a dental implant, an abutment, and a crown. The dental implant is analogous to the tooth

root and is expected to osseointegrate with the jaw, the crown is the dental prosthesis,

and the abutment connects the two and interfaces with the gums. Dental implants offer

many functional benefits over dentures, such as improved speech and ease of eating, but

also important aesthetic benefits related to appearance and self-esteem.3 However, long-

term dental implant complications are critical issues which have called into question the

definition of implant success along with celebrated 5% failure rates.2 When implants

were first introduced, an implant surviving in the mouth after 5 years of function was

deemed successful. However, over the last decade, research has identified significant

consequences in surviving implants involving infection, inflammation, and declining

aesthetics. This is especially poignant in light of reports that up to 48% of implants show

soft tissue inflammation.4–9 This inflammation can be triggered by a variety of highly

prevalent factors such as the implant components themselves, host response to residual

cement, bone morphology, soft tissue barrier, implant loading conditions,

provisionalization protocols, and bacterial plaque.10–13 Clinical resolution of the

pathologies around implants remains a significant challenge in clinical practice with no

gold standard therapeutic protocols to address these complications.14–16 Dental implant

complications can be categorized into three genres: disease, function, and aesthetics, all

of which can be traced back to the implant-bone interface.

Soft-tissue disease around natural teeth and dental implants occur at about the

same rate, in that approximately 47% of adult Americans suffer from gingivitis17,18 and

48% suffer from peri-implant mucositis.7 Peri‐implant mucositis is described as an

inflammation of the soft tissue around dental implant without any signs of marginal bone resorption. A more severe complication is peri-implantitis, where more than 2 mm

of bone surrounding the implant resorbs as a result of bacterial infiltration. Peri-

implantitis has an incidence rate of approximately 22%.2 While dental implant diseases

are analogous to tooth diseases, typically infection and the related tissue destruction

spreads more rapidly and more severely around dental implants.19 One explanation for

this difference is the differing tooth-root interfaces that the dental implant and natural tooth experience. Naturally, the cementum of teeth and bone are attached by periodontal ligament. Current fixation involves screwing the tooth root replacement into the bone, generally resulting in a patchwork of different interfaces which are all unlike the natural

interface.20,21 While the current dogma of dental implants is that full osseointegration

should occur, full ankylosis around implants is quite rare, points of ankylosis being

more common. Where implant ankylosis does not occur, commonly patches of scar-like

long-junction epithelium form. This fills the void between the implant and the bone, but

is very weak and does not provide a sufficient barrier in either strength or bacterial

resistance.22 After implantation, the exposed titanium surfaces can accumulate

glycoproteins and form a dental pellicle, allowing bacteria to infiltrate and begin biofilm

formation. Any points of ankylosis are also susceptible to bacterial invasion as there is

no periodontal ligament to act as a buffer zone.19 Implementing a synthetic periodontal

ligament at the interface of the bone and the titanium implant may reduce incidence

rates of peri-implantitis and peri-implant mucositis by providing that biomimetic barrier to bacteria.

The function of a dental implant is related to how well it can withstand typical forces of mastication, and how well it maintains the strength of the surrounding bone. A dental implant, its abutment, and its crown should be able to withstand the forces of mastication. A single molar implant might experience about 120N in the axial direction,23 and there may also be short-term force maximums up to 847N in healthy young men and 595N in women.24 Ankylosed teeth and dental implants exhibit lessened

alveolar bone strain under the above forces compared to teeth having a periodontal

ligament due to an insufficient transfer of load.24 Following Wolfe’s Law, this deficit in

alveolar bone strain can lead to deterioration and weakening of the bone, eventually

causing functional detachment of the implant from the jaw years after implantation.25

Adding a synthetic periodontal ligament to the bone-cementum interface may cause

more biomimetic stress to be applied to the bone, hindering resorption.

As patients receiving dental implants live longer, issues surrounding dental

aesthetics become more prominent. The jaw continues to grow as humans age even past thirty. Natural teeth that are surrounded by periodontal ligament can be moved via orthodontia, but ankylosed implants are mechanically fixated to the jaw bone. As such, as the jaw moves, artificial teeth follow the path of the jaw, causing them to fall out of alignment with other teeth.26,27 This process can not only hinder the function of the

prostheses in mastication and also lead to unsightly crooked artificial teeth. An artificial

periodontal ligament surrounding the dental implant could allow for orthodontia

following implantation and the healing process, therefore allowing any damaging

aesthetic changes to be repaired.

1.2 Periodontal Ligament

Between the cementum

of a tooth and the alveolar bone

of the jaw, there is an

unmineralized, largely

collagenous section of tissue

visible by clinical cone-beam x- Figure 1.1: Simple periodontal ligament. ray imaging.28,29. This short Structure of the periodontal ligament and corresponding cementum and alveolar bone interfaces. The brown fibers represent Sharpey’s fibers. unmineralized section

separating the mineralized tissues is referred to as the periodontal ligament (PDL).

Periodontal ligament separates teeth from bone, allows the teeth to move during

mastication, as the jaw grows, and makes orthodontia possible. It is a fibrous tissue that is resilient, responsive, and fulfills many functions in maintaining the health of the tooth and bone. Figure 1.1 includes a simplified diagram of the periodontal ligament, showing only the main structural elements of the periodontal ligament and its interfaces with the cementum and alveolar bone. The main structural components of the ligament, the

Sharpey’s Fibers depicted in brown, insert perpendicularly and mineralize into surrounding tissues. These fibers act as the chief mechanical loading structure of the ligament. The main structural elements of the Sharpey’s fibers are anisotropically aligned type I collagen fibrils. Figure 1.2 shows a diagram of the highly organized structure of type I collagen fibrils. Collagen is a fibrous extracellular matrix protein

with a molecular weight of 300 kDa, a length of ~300 nm, and a diameter of ~1 nm and

are comprised of a triple helix made of three alpha peptide chains Within the fibril, the

collagen molecules arrange into a repeating structure called a D-staggered array with a

characteristic 67 nm banding pattern, where the ends of adjacent molecules are

separated by a 40 nm gap zone. Collagen fibrils are typically 50-100 nm in diameter and

many microns long.30

Figure 1.2: Structure of a collagen fibril. Collagen molecules (left) are arranged in fibrils (right) in a staggered fashion such that the ends of adjacent molecules are separated by a 40 nm gap zone. This configuration results in D-banded physiological fibrils.

The periodontal ligament is highly vascular and cellular. Figure 1.3 shows a

more complete diagram of the anatomy of the periodontal ligament including the non-

collagenous components.31 The largest functional component of the PDL’s extracellular

matrix is the collagen fibrils (Sharpey’s fibers) that span the length of the ligament, about 100-400 µm, between the alveolar bone and root cementum.32 Other extracellular

matrix components include elastin fibers and the ground substance.33 The ground

substance is a hydrogel consisting mostly of water, proteoglycans (e.g. decorin,

Figure 1.3: Cellular and vascular periodontal ligament. The periodontal ligament is highly cellular and vascular and contains many other extracellular matrix components around the collagen fibers. The collagen fibrils are depicted in brown, and the other extracellular components are in the pink background.

versican, perlecan), glycosaminoglycans (e.g. hyaluronic acid, heparin sulfate), and glycoproteins (e.g. fibronectin, laminin, tenascin). The PDL also has an extensive blood supply, a neural network, and a large cell population. These cells include fibroblasts, bone cells, cementum cells, endothelial cells, epithelial rests of Malassez (ERM), neural

cells and the various stem and precursor cells of the bone and cementum. Fibroblasts are

the predominant cell type of the PDL and are responsible for the production and

maintenance of the extracellular matrix.31–34

There are six principal functions of the periodontal ligament: homeostasis,

inhibition of periodontal disease, bone remodeling, sensory feedback, nutrition, and

tooth support.35 A homeostatic balance exists between periodontal fibroblasts and

osteocytes lining the alveolar bone socket, preventing encroachment of the bone and

resisting ankylosis.36 The collagen fibrils and other matrix components in periodontal

ligament act as a buffer for bacterial infiltration, hindering periodontal disease.19

Additionally, the vascularity of the ligament along with its population of immune cells can fight bacterial infiltration if the mechanical barrier does not suffice. The structures of the ligament and surrounding bone are constantly undergoing remodeling, and the ligament responds to tooth movement, injury repair, and occlusal forces. In these functions, pressure on the ligament stimulates bone resorption while fibril tension stimulates cementum and bone formation, maintaining the distance between the cementum and alveolar bone.37 Mechanoreceptors called Ruffini nerve endings within the ligament respond to force application, such as particles wedged between teeth or stuck on the occlusal surfaces, and provide sensory input to aid in injury prevention.35

The matrix of the periodontal ligament protects the bone, blood vessels, and nerves necessary for the health and nutrition of the surrounding tissue from occlusal forces.38 A vital function of the periodontal ligament, and one that is a major focus of synthetic periodontal ligament design, is the tooth support mechanism. The ligament acts as a medium of force transfer during chewing and, given its viscoelastic behavior, behaves as a shock absorber while also transferring higher bone strains compared to ankylosed teeth.25,38 A synthetic periodontal ligament, principally the mechanical aspects, surrounding artificial tooth roots could address the clinical complications which detract from the success of dental implants.

1.3 Artificial Ligament Systems

For this thesis, we are only concerned with producing an artificial scaffold for periodontal ligament: a brush of Sharpey’s fibers consisting of anisotropic collagen fibrils. These fibers may act as a strong structural interface to limit the bacterial, loading, and aesthetic complications found on existing dental implants. We want to

create structures that closely match the mechanical properties, function, and

biochemistry of the native Sharpey’s fibers.39 In the future, work can be done to seed the

Sharpey’s fibers with cells and/or add more of the extracellular matrix components

found in natural periodontal ligament if necessary.

The regeneration and replacement of ligaments have been of interest to

researchers for many years. Replacement fibers created with synthetic polymers have

been largely unsuccessful due to long-term deterioration or inflammatory responses.40

For this reason, we decided to use collagen as the basis for biomimetic artificial

Sharpey’s fibers bound to a dental implant. As an initial proof-of-concept, we sought to covalently conjugate collagen fibrils end-on to a titanium surface to mimic the dentin-

Sparpey’s fibers interface. This goal could be accomplished in one of two ways: conjugating pre-made collagen fibrils to a surface, or by binding single collagen molecules to a surface and building that molecule into a fibril via a self-assembly process known as fibrillogenesis. Animal-made collagen fibrils are often extracted from animal tissue such as bovine skin and rat tail.41 Fibrillogenisis is a spontaneous

procedure whereby solution-phase collagen molecules self-assemble into fibrils under

physiological conditions. These free collagen molecules are also typically sourced from

bovine dermis and are usually in the form of pepsin-digested atelocollagen. Figure 1.2

shows how collagen molecules assemble into fibrils.

While other research groups have created structures out of anisotropically

aligned collagen fibrils, no one has ever covalently bound collagen fibrils upright on a

surface. Aligned fibril mats can be made by flow-induced crystallization42 or

electrochemical fabrication,43 but these processes result in flat planes of collagen fibrils

that are not bonded to a surface, much less end-on. A post-fibrillogenesis chemical

conjugation step would have to take place to turn these mats into implant-bound

Sharpey’s fibers. Another group developed a surface of electrostatically-bound bovine

collagen fibrils that do stand perpendicular to a surface.44 This design, however, relies

on non-covalent interactions between surface-bound poly-L-lysine and the C-termini of

collagen molecules, which is non-specific to the end of the collagen fibrils. There are

only three carboxyl-termini per collagen molecule, yet there are 71 negatively-charged glutamic acids and 43 aspartic acids per 1,000 amino acid residues. This method of conjugation relies on positive-negative amine-carboxyl interactions, which are weak with respect to covalent bonds. The method also has a high potential to bind negatively- charged amino acids, rather than the C-termini of the final collagen molecules on the fibril, to the poly-L-lysine surfaces. Additionally, there is a possibility that the C-termini of the collagen monomers within the collagen fibril could bind to the surface.

1.4 Collagen Self-Assembly: Fibrillogenisis

Given the issues associated with using pre-formed fibrils outlined in section 1.3,

I decided to use a system that takes advantage of fibrillogenesis, or collagen’s ability to self-assemble into physiological fibril, directly on the surface of a titanium implant. This system would require that individual collagen molecules be bound to a titanium surface, which could then be used as nucleation sites for a fibrillogenesis procedure.

Physiological fibrils exhibit a pattern known as D-banding, outlined in Figure 1.2.

Though the exact mechanism for fibril formation is still being debated, evidence suggests that D-banded fibrils form in a three-step process consisting of pre-nucleation,

nucleation, and growth as outlined by the Fred Silver group.45 These steps all occur

under a range of solution conditions, which are discussed later in this section. The pre-

nucleation step involves the formation of unstable, reversible aggregates containing

about five collagen molecules. Pre-nucleation continues as these five-membered

aggregates arrange laterally into still unstable pre-nuclei. Nucleation follows this

procedure and involves the formation of a “unit fiber” which consists of approximately

105 collagen molecules gathered from the pre-nuclei. The unit fiber is a stable aggregate,

which can then rapidly grow longitudinally. The unit fibers are discussed in this thesis

as nuclei.

Fibrillogenesis can be monitored through turbidity-time assays. As fibrils form in solution, the liquid becomes turbid, and this increase in optical density is directly related to the rate of collagen fibril formation and the number of fibrils formed. Figure

1.4 shows a typical turbidity curve for Type I collagen fibrillogenesis. The lag phase, denoted in Figure 1.4 between points a and b, represents the period where nuclei are being formed

(pre-nucleation and nucleation). During this period, no longitudinal

fibril growth occurs and the

Figure 1.4: Typical Turbidity vs. Time profile of collagen self-assembly. nuclei are too small to This diagram reveals the kinetic process, model, and mechanism of fibrillogenesis. a and b represent the beginning and end of the lag period scatter light, so there is no respectively, where collagen nucleation occurs. Point C represents the end of the growth period and the beginning of the plateau period. change in optical density.

Following the lag phase is the growth period, where fibrils grow longitudinally, causing

the optical density of the fluid to increase. Finally, fibril growth plateaus due to the lack

of free collagen molecules in solution. During nucleation, the number of nuclei formed

is dependent on the collagen concentration, where a higher monomer concentration

leads to more nuclei. The temperature of the fibrillogenesis solution determines the rate

of nucleation, as increased temperature leads to a shorter lag-phase in turbidity curves.

For this thesis, I will discuss pre-nucleation and nucleation as a single process

simply called nucleation. Thus, in this thesis, fibrillogenesis is broken up into two steps:

nucleation and growth, which can be visualized by the lag and growth periods,

respectively, in the turbidity vs. time profile. I chose to combine pre-nucleation and

nucleation because, as Farber et. al. describes it, nucleation is the rate-limiting step and

there is a clear delineation in turbidity curves between nucleation and growth.

Collagen is easily removed from mammalian tissues via a pepsin digest, so all

collagen referred to from this point on is atelocollagen, where the non-helical ends of

the collagen molecules have been removed.46 These atelocollagen molecules, such as

those in the commercial Type I collagen solution called “PureCol”, still assemble into

D-banded fibrils. (PureCol, Advanced BioMatrix, San Diego, CA) Several

fibrillogenesis methods work to produce physiological fibrils as outlined by Holmes et.

al.47 The Eppell lab has found success using their “Simultaneous neutralization and

warming” fibrillogenesis procedure. This Holmes protocol calls for a 4℃ solution of 1.8 mg/mL collagen to be diluted 10x in a 37℃ buffer such that the final solution

temperature is 34℃ with a collagen concentration of 0.18 mg/mL, a phosphate

concentration of 12 mM, and a pH of 7.4. Though this original protocol does not specify

the other salts included in the buffer, the most common phosphate-buffered saline

solutions also contain 137 mM of NaCl, which is what is used in the Eppell lab.48,49

Other than the above recipe, collagen fibrillogenesis can be carried out in a wide

range of different conditions, all of which lead to D-banded fibrils. In a buffer containing a 10 mM phosphate, D-banded fibrils form with NaCl concentrations ranging from 50 to 500 mM, where higher NaCl concentrations lead to more tightly-packed collagen molecules.50 Lower NaCl concentrations between 10-20 mM lead to the

formation of diffuse fibrils with early indications of banding. In a buffer containing 150

mM NaCl, D-banded fibrils from with phosphate concentrations ranging from 10-50

mM.51,52 All of these conditions will lead to D-banded fibrils in a temperature range of

22-37℃ and a pH range of 6.5 to 8.46,50,51,53 Collagen will also aggregate under an

extensive list of different conditions involving phosphate and NaCl, but these conditions

are not conducive to creating D-banded fibrils. Fibrils formed may be smooth (lacking

banding) indicating a lack of ordered structure, or may have different banding patterns

such as an 11 nm repeat rather than the normal 67 nm repeat. A number of these

conditions garnered from the Lewis group are outlined in Table 1 below.

Table 1: Solution conditions that lead to the formation of collagen aggregates that are not D-banded fibrils. All fibrils were formed at a temperature of 22℃ and a collagen concentration of 0.02 mg/ml. Fibrils were assayed by transmission electron microscopy.

2+ pH [NaCl] (mM) [PO4 ] (mM)

2.5- 5.0 150 2051

4.0 10-500 050

7.0 0 10-50050

1.5 Non-Specific Covalent Bonds Between Proteins and Surfaces

The system developed for this thesis requires that individual collagen molecules

be bound end-on to a titanium surface and that those bound molecules are used as

nucleation sites for a fibrillogenesis procedure. There are many possible ways to

covalently bind proteins to surfaces. Any reaction used to bind a collagen molecule end-

on to a surface will involve either the terminal amine or terminal carboxyl groups of the

molecules. As such, I will focus on conjugation methods involving terminal amines or

carboxyls. EDC (1-Ethyl-3-(3-dimethylaminopropyl)carbodiimide) or EDC/NHS (N-

Hydroxysuccinimide) reactions are the most popular techniques used to form covalent

peptide bonds between carboxyl and primary-amine groups.54 However, amines and

carboxyls are not unique to the termini of proteins and collagen is no exception. There

are only 3 terminal amines and 3 terminal carboxyls per collagen molecule (due to is

having three peptide chains), but around 240 residues containing carboxylate side-

groups (aspartic and glutamic acids) and 100 containing primary amine side-groups

(lysines).55,56 Since these side-groups are much more prevalent than the N- or C- termini, carbodiimide-based procedures are insufficient to site-specifically conjugate

collagen end-on to a surface.

1.6 Carbonyl Formation and Oxyamine Conjugation

Figure 1.5: Transamination and Oxime Formation scheme. Scheme presented by the Francis group for the transamination of protein n-termini followed by oxime linkage to an oxyamine-containing molecule. The blue globular structure represents the rest of the protein or peptide, and the orange star represents the rest of the oxyamine-containing molecule. Examples of transamination reagents include pyridoxal-5-phosphate (PLP) and N-methylpyridinium-4-carboxaldehyde benzenesulfonate (Rapoport's salt, RS)

Carbonyl-reactive conjugation methods have become more popular in recent

years because ketone groups are relatively rare to most proteins, and they can be

introduced by biological transamination reactions. Oxyamine groups react selectively

with these aldehyde or ketone groups to form oxime linkages, thus allowing various

ketone- and oxyamine-containing molecules to be conjugated together.57 Oxyamine

groups may also be referred to as hydroxylamine or aminooxy groups. Figure 1.5 shows

an overview of this reaction scheme. Examples of transamination reagents include

pyridoxal-5-phosphate (PLP) and N-methylpyridinium-4-carboxaldehyde

benzenesulfonate (Rapoport's salt, RS). While some proteins do have aldehyde or ketone

groups naturally occurring in their amino acid sequence, collagen does not.55,56,58

Transamination reactions performed with these aldehyde reagents are highly specific to the N-termini of proteins.59 For these reasons, the ketones introduced to the N-termini of

collagen molecules would be unique, allowing for site-specific conjugation of collagen

molecules to oxyamine-containing molecules. A surface decorated with oxyamine functionalities could, therefore, be used in conjunction with the transamination chemistry to produce a surface covered in end-bound collagen molecules.

Pyridoxal-5-Phosphate as a Transamination Reagent

In 2006, the Francis group introduced a new method for the transamination of

the N-termini of angiotensin I using pyridoxal-5-phosphate (PLP), an enzyme cofactor otherwise known as active vitamin B6.60 In 2010, the group published an article

detailing methods to apply this novel approach to a wide range of other peptides and

proteins.59 The recorded protocol involves mixing a protein or peptide with PLP in

phosphate-buffered saline at pH 6.5 and incubating at 37C for 2-24 hours. The resulting

keto-peptide is then reacted with an oxyamine-containing molecule, resulting in oxime

bond formation between the two molecules. Figure 1.5 shows an overview of this

reaction scheme. The pKa of N-terminal α-amines (pKa = 6–8) is lower than that of

other aliphatic amines (pKa 10.5) such as the lysine side group, so this PLP reaction

can target N-terminal amines∼ by carrying out the reaction at pH 6.5.61 The Francis group

has done a large amount of work characterizing the effects that the N-terminal residues

have on the final product following exposure to PLP. The following is a timeline of the

evolution of the Francis group’s transamination procedure.

In their first 2006 paper, the Francis group was assaying for a better way to do

site-specific protein modifications.62 They wanted to find an alternative to popular

techniques targeting unique sulfhydryl groups on cysteine residues, which can be done

with reactive maleimide chemistry.63 This sulfhydryl technique is not site-specific when

proteins contain more than one cysteine residue, can be inconvenient or impossible on

some proteins, and can result in changes within the structure of a protein thereby

influencing its functionality. Targeting the N-terminus of proteins would alleviate these

issues. N-terminal modification had been previously accomplished by oxidation of the

N-terminal amine to an imine followed by hydrolysis into a carbonyl group (aldehyde or

ketone) through the use of glyoxylic acid, copper(II) salts, 1 M pyridine, or 1 M acetic acid.64 These reaction conditions, however, are harsh and affect the folded structure of most proteins.62 Experiments with glyoxylic acid led to the discovery that aldehyde- containing molecules could participate in site-specific transamination reactions with the

N-termini of peptide sequences. Through assays of eight different aldehyde-containing reagents, they found that PLP was the most effective in transaminating the N-terminal amino groups of angiotensin I. They assessed the yields of their reactions with PEG‐ alkoxyamine. The analysis was done by SDS‐PAGE with Coomassie staining. They further tested the effectiveness of the PLP reaction on other proteins, and the results are displayed in

Table 2.62 Thus, they developed a bioconjugation strategy that is compatible with proteins containing free cysteine residues and can also be used to label proteins when chemistry appropriate for cysteine groups is not available.

Table 2: Results of a study using PLP as a transamination agent on various proteins.

Protein Properties N-terminal T Conv.

residue [℃] [%]

Angiotensin I Globular Aspartic 37 65

1.3 kDa Acid

Hormone

Myoglobin Globular Glycine 37 69

16.7 kDa

Oxygen carrier

GFP Globular Valine 55 67

26.9 kDa

Fluorescent

GFP, mut. “” Glycine 37 41

GFP, mut. “” Glycine 55 80

RNase A Globular Lysine 37 50

13.5 kDa

Enzyme

Thioredoxin Globular Glycine 37 50

14 kDa

Redox protein

Protein G’ Globular Methionine 41 30

65 kDa

Bacterial cell wall protein

Following the success of the PLP reaction in the 2006 paper, they moved on to

modifying monoclonal antibodies and published a paper on the topic in 2007.60,62

Monoclonal antibodies have many potential applications as imaging agents, targeting

groups for therapeutic drug delivery, components of diagnostic arrays, and catalysts.

These applications all require the addition of new functionalities to these antibodies,

such as fluorophores, drugs, or nanoparticles. Popular conjugation techniques target

amino acid residue side chains, leading to these functionalities appearing at random

locations across the antibody. Thus, the Francis group applied the PLP reaction and

subsequent oxime formation to monoclonal mouse anti-FLAG IgG as a regiospecific

modification strategy. They assayed for reaction efficiencies using AlexaFluor 488

alkoxyamine and SDS-PAGE, and found 25% conversion of the light chains at 37°C

and 47% at 50°C, but were unable to quantify the low reaction efficiencies of the heavy

chains.

Given that the previous study revealed that different protein N-terminal residues displayed varying levels of transamination with PLP60, in 2008 they developed a method

for screening a 20 member tetrapeptide library for PLP reactivity.65 In this study, they

quantified the different potential products for each of these N-terminal residues, thereby

predicting the success of a protein or peptide transamination reaction. The tetrapeptides

tested had amino acid sequences X-KWA, where X represents one of the 20 naturally

occurring amino acid residues. They generated the 20-member peptide library using

solid-phase peptide synthesis via Fmoc-based chemistry along with liquid

chromatography-mass spectrometry analysis. For the PLP reaction, the peptides were

treated with 10 mM PLP in 50 mM phosphate buffer at pH 6.5 with 10% N,N-

dimethylformamide (DMF) for 18−20 h at room temperature. Following extensive washing, the peptides were then incubated with 250 mM O-benzylhydroxylamine hydrochloride (BnONH2) for 1.5 h at room temperature. Table 3 shows an overview of the results of this experiment.

Table 3: Results of 20-tetrapeptide library screening for N-terminal residue PLP activity.

N-terminal Residue Expected Product

A, G, D, E, N High conversion to oxime (>80%)

C, R, T, Y, L, S, M, F, or Varying levels of conversion

Q Transamination, but minimal oxime conversion

W, H Pictet-Spengler product in high yield

P PLP adduct, some oxime formation

K Mostly cyclic imine and resulting PLP adducts. Some species may undergo oxime conversion Through this screening, they discovered that the terminal amino acid residue is not the sole determining factor for the success of a reaction.65 For example, in the 2006 Gilmore et. al. paper, they were able to modify GFP with an N-terminal Valine with good yield

(67% conversion to oxime)62, but in this study, valine is one of the poorest residues for transformation, with only a 10% conversion to oxime. Beyond predicting the success of a transamination reaction, the Francis group also used this information to transaminate tobacco mosaic virus (TMV) coat proteins with native Serine N-terminal residues. These proteins originally showed no transamination following PLP exposure, so they changed the N-terminal residue of these proteins through mutation to a highly reactive Alanine, and added a spacer residue to prevent steric hindrance. These results suggest that the

secondary and tertiary structure of proteins can lead to an N-terminal environment that is not compatible with modification by PLP.

The group then published two more papers in 2008 regarding polymer hydrogels that are impregnated with protein crosslinks. In the first paper, the Francis group used a

PLP reaction to introduce ketones onto the N-termini of GFP, and used a separate protein engineering strategy to produce ketones on the C-termini.66 Using mass spectrometry, they found that the PLP reaction had a conversion rate of about 50%. The polymer N‐(3‐aminopropyl)methacrylamide was functionalized to bear oxyamine groups. The protein and the polymer were cross-linked via oxime bond formation. In the second paper, they used their site-specific PLP chemistry to create pea-metallothionein- containing hydrogels for the removal of heavy metals in water treatment systems.67 Pea metallothioneins capture toxic metal ions, such as copper and zinc, by condensing from an expanded state to form binding pockets. To maintain the pocket-forming function, the primary amino acid sequence must be free to condense, and as such the effective strategy involves N- and C-terminal conjugation of these proteins into polymer matrices.

Using mass spectrometry, the found that the PLP reaction had a conversion rate of about

38%. These groups are then used to crosslink alkoxyamine-containing polymers through oxime formation.

In 2010, the group published a paper reiterating the data garnered from the 2008

Scheck et. al. paper and included an extensive protocol to generalize their PLP process to any number of proteins.59,65 The general protocol is as follows: mix 10 mM PLP and

10 to 500 μM protein with PBS at pH 6.5; Incubate at 37 °C for 2–24 h; Remove PLP using one of a variety of size exclusion methods. In this 2010 paper, the group included

crucial details about the process, such as the importance of using a fresh solution of

PLP, checking and adjusting the pH following PLP addition, and storing the solid PLP

at 4°C.

Following the results of the initial tetrapeptide screening library discussed in the

2008 Scheck at. al. study65, the Francis group sought to increase their screening process to include not just the final amino acid residue, but the final three. In the 2008 paper,

they found that the N-terminal sequence of the protein can significantly influence the

overall success of this PLP transamination strategy. Thus, in 2010 they developed an

8,000-member combinatorial peptide library where all peptides had the form

XXXWSNAG.68, where X represents one of the 20 naturally occurring amino acid

residues. Each peptide in the library was subjected to PLP-mediated transamination and

subsequent oxime formation with a Disperse Red alkoxyamine dye. Through this

screening, they found that AKT was a particularly reactive amino acid sequence with a

conversion rate of ~90%.

In 2012, the Francis group published a paper outlining the site-specific N-

terminal modification of the ~4,200 coat proteins on filamentous phage.69 These were

PLP-modified so that they can then serve as chemispecific handles for the attachment of

alkoxyamine groups through oxime formation. The overall protein recovery for the

transamination and oxime formation steps ranged from 55–95%, with 80% being a

typical value.

Many other groups have used PLP to modify proteins and conjugate them to

various surfaces.

In 2007, Christman et. al. used PLP to immobilize streptavidin onto to a

micropatterned 2-hydroxyethyl methacrylate and Boc-protected aminooxy-

tetra(ethylene glycol) methacrylate copolymer.70 This work hoped to further the field of

diagnostic arrays and biomaterials, where the orientation and activity of proteins are

essential. Streptavidin has an N-terminal alanine residue, and at this point, the alanine

residue had not yet been tested with PLP. Streptavidin is a bacterial globular protein

with a molecular weight of 52.8 kDa and a strong binding affinity to the protein biotin.

They modified streptavidin with a 50 mM PLP solution in a pH 6.5 phosphate buffer. To

show the efficacy of this reaction, they incubated the modified streptavidin with the

previously mentioned micropatterned polymer films. They then visualized the

attachment of streptavidin via antibody staining with a primary anti-streptavidin

antibody followed by an Alexa Fluor 488 secondary antibody. Significant fluorescence

was found on surfaces where the protective Boc- group of aminooxy functionalities

were removed.

In 2009, the Chilkoti group modified myoglobin using PLP to conjugate the

protein to PEG, thereby improving its pharmacokinetics and solubility.71 Myoglobin

was reacted with a 20 mM PLP solution in 25 mM phosphate buffer, pH 6.5 and

incubated 37 °C for 36 hours. The PLP was then removed via ultracentrifugation. The

protein was then reacted with the aminooxy-containing atom transfer radical polymerization initiator (for the conjugation of PEG), giving an oxime yield of approximately 75%.

In 2009, Lempens et. al. modified glutathione S‐transferase (GST) and protein G′ with PLP to make surface‐based assays.72 GST is a 25 kDa globular catalyst protein.

Each protein was combined with 6.7 mM PLP in 50 mM, pH 6.5 phosphate buffer and

incubated overnight at 41 °C. Using a 5 kDa PEG-alkoxyamine and SDS‐PAGE

analysis, the found 22 % conversion of GST and 27 % conversion protein G′.

In 2017, the Pokorski group used PLP to create a wound patch conjugated with

epidermal growth factor (EGF).73 The patch, made of the flexible and biocompatible

polymer poly(ε-caprolactone) (PCL), was designed to release EGF in response to the

presence of matrix metalloproteinase-9 (MMP-9), an enzyme that is active during the

early stages of the wound-healing cascade.74 EGF is a small 6 kDa protein that

stimulates cell growth. The PCL patch was made to present surface hydroxylamine groups, facilitating the conjugation of ketone-containing molecules. The N-terminus of the protein was modified to express an MMP-9-cleavable octapeptide sequence followed by a PLP-modifiable AKT terminal sequence. In this way, the recombinant EGF can be transaminated and subsequently bound to oxyamine-containing substrates and released via cleavage of the octapeptide sequence. The transamination procedure involved the reaction of 100 mM PLP with 5 mM recombinant EGF in PBS buffer titrated with 6N

NaOH to pH 6.5 and incubated at 37 °C for 1 hour. Following the reaction, excess PLP was removed from the system via centrifugal filtration. To assess the efficacy of the transamination reaction, the ketone-protein was exposed to aminooxy-5(6)-TAMRA dye in the presence of 10 mM of aniline at 37 °C for two hours. Following the reaction, excess TAMRA was removed via spin filtration. Protein conjugated with the TAMRA dye was analyzed with fast protein liquid chromatography (FPLC) monitoring at 555 nm for TAMRA and 260 and 280 nm for protein.

Rapoport’s Salt as a Transamination Reagent

Although the Francis group found great success with the N-terminal modification of peptides by PLP transamination, they claim that its reactivity often 25

varied batch-to-batch, though they do not report these results explicitly.75 Given this finding, they decided to screen for an alternative aldehyde reagent with a similar reactivity, but with more consistent repeatability. In 2013, the group combined the aldehyde screening methods used in their original 2006 paper 62 with the 8,000-member combinatorial peptide library outlined in their 2010 paper68 to assess new transamination reagents while simultaneously identifying their optimal N-terminal sequences. N- methylpyridinium-4-carboxaldehyde benzenesulfonate salt (Rapoport's salt, RS) was found to be a highly effective transamination agent when used with glutamate- terminated amino acid sequences, making it particularly well suited for use with antibodies. The N-terminal sequence EES was particularly reactive to RS, showing over

80% transamination and ~70% oxime formation. DES showed about a 50% oxime conversion rate, and AES about 40%. Proline showed no reactivity to RS when at the N- terminal position, and when proline is in the second position of an EPS sequence, the conversion is greatly reduced. Isoleucine was similarly unreactive. Using RS, they were able to modify the heavy chains of the wild-type monoclonal antibody Herceptin without affecting its binding affinity to the HER2 receptor. The wild-type heavy chain of this antibody (N-terminal EVQ) underwent 67% conversion to the oxime product, in which 15% also included the RS adduct. No modification of the light chain was seen, likely because of steric hindrance and the n-terminal sequence (DIQ). In modifying both chains to express the ideal EES, they found 56% and 68% conversions for the light and heavy chains, respectively. Again, they did not report the reproducibility for these reactions.76

Summary of Carbonyl Formation and Oxyamine Conjugation

A transamination and subsequent oxime-formation protocol has not been used 26

successfully on collagen. In fact, there is no evidence in the literature that any

extracellular matrix proteins have been transaminated using these protocols. The Francis

group and others have used PLP as a transamination agent to site-specifically modify

many different proteins, including streptavidin, IgG, and filamentous phage capsid

proteins59,61,62,69,70,75. This reaction shows high specificity to N-terminal amines, showing no conjugation to lysine side chains. The PLP installs a ketone functionality to the N-terminus of protein molecules, and these ketones are highly reactive to oxyamine functionalities. This reaction forms a stable oxime bond between the two molecules. In their extensive characterization of PLP reactivity, the Francis group found that some N- terminal amino acid residues produced high yields of N-terminal transamination and subsequent oxyamine-conjugation (above 80%), such as alanine, glycine, and aspartic acid. Other amino acid residues, however, showed little to no N-terminal conversion and oxyamine conjugation, such as isoleucine and histidine. Lightly pepsin digested type I collagen (atelocollagen) displays three different triplets on the three alpha chains making up the molecule: ISV, DEK, and DAK.77 Since two of the N-terminal residues

(those ending in D, a.k.a. aspartic acid) showed some reactivity to PLP, we decided that

collagen would be a suitable candidate for a PLP reaction.

1.7 Silane Surface Reactions

To make use of ketone-modified collagen molecules, we needed a method to bind oxyamine groups to the titanium surfaces. One way to bind ketone-reactive oxyamine groups to surfaces is to use an oxyamine-terminated silane. Silane coupling agents are synthetic hybrid inorganic-organic compounds used to promote adhesion between dissimilar materials and can be terminated with virtually any functional group.

Standard silanization procedures usually involve a hydrolysis step where the protective

hydrocarbon groups are removed from the reactive Si-O group. This hydrolysis opens

the molecules up to bulk gelation, which can be useful in the formation of aerogels, but

not so much for the formation of surface monolayers.78,79 Amine-terminated silanes provide an advantage over other silane molecules by using an amine-catalyzed reaction.

Amines can catalyze the reaction between meth- or ethoxysilanes and hydroxyl groups.

Figure 1.6 shows two methods for conjugating aminosilanes to silicon surfaces.80

The solution-phase chemistry shown in the upper section of the figure can be

done without special laboratory equipment and results in surfaces where the amine

groups are free to react with solution-phase constituents. Solution-phase chemistry can

be carried out as follows. The surfaces are first functionalized with hydroxyl groups

typically using a sulfuric acid/hydrogen peroxide wash (piranha solution) or room-

Figure 1.6: Amine-catalyzed silane surface reactions. (top) Solution phase reaction wherein the amine group interacts with the Si atom of the silane molecule to form a pentacoordinate intermediate that is highly reactive to nucleophiles like surface-bound hydroxyl groups. (bottom) gas-phase reaction where the amine group forms a coordination bond with surface-bound hydroxyl groups, causing a nucleophilic attack of the silane group and binding the molecule to the surface.

temperature plasma treatment.81,82 In an anhydrous alcohol solution, the carbon chain on

the aminosilane can bend, allowing the amine to form a coordination bond with the Si-

O, encouraging the hydrocarbon-alcohol to leave. This process results in a stable covalent bond between the titanium and the silane. Such a reaction requires that the solution be anhydrous because otherwise, silane molecules will bond together to form bulk gels. Gas-phase reactions avoid exposure to water vapor, but these protocols typically result in the amine group being H-bonded to surface hydroxyl groups. These bound amines are less likely to participate in other reactions and also lead to less-dense surface coverages.83

Chapter 2: Study Design

2.1 Study Goal

The complications associated with dental implants may be traced back to the

bone-implant interface. As such, the long-term interest of this research is to build

synthetic periodontal ligament on titanium dental implants. Mimicking the natural

interface between bone and teeth may alleviate dental implant complications by

providing a more biomimetic force transfer between the artificial tooth and the bone,

providing a barrier to bacterial infiltration, and allowing orthodontia to take place if the

dental implant moves out of place.29,33,84,85 The goal of the present study was to develop

a method to conjugate collagen molecules, and subsequently collagen fibrils, to a

titanium surface as a proof-of-concept for the goal of producing the main structural

component of the periodontal ligament: the Sharpey’s fibers. We chose the Sharpey’s

fibers as a starting point because they make up the majority of the periodontal ligament,

act as the main structural support for the ligament, and can act as a scaffold onto which

other biological factors could be added in the future.32,33

2.2 Experimental Design

We seek to mimic the structure of natural periodontal Sharpey’s fibers by conjugating collagen fibrils perpendicularly to a titanium surface. Our method is a bottom-up procedure, allowing us to create covalently bound, anisotropically aligned collagen fibrils on a titanium surface, similar to the structure of the natural Sharpey’s fibers. The titanium oxide surface functions as a replacement for the tooth enamel.

Titanium is chosen because it is the most common choice for commercially available dental implants.

The structure of the developed artificial Sharpey’s fibers can be simplified by looking at one single collagen fibril. Figure 2.1 shows a general overview of this structure. A titanium surface is first functionalized by silanization with a monolayer of oxyamine-terminated silanes. The silane bond is represented by a black dot. The surface is then coated with modified collagen molecules that have ketones, rather than amines, at their N-termini represented by the yellow dot. A site-specific reaction occurs between the oxyamine and the ketone producing a covalent oxime bond. The single bound collagen molecules then act as nucleation sites for the growth of self-assembled collagen

Figure 2.1: Simple overview of the structure of a single collagen fibril bound to a titanium surface.

fibrils.

Appendix A details all of the protocols developed to build collagen fibrils on

titanium surfaces. Section 2.3 describes the methods by which this protocol was

developed.

2.3 Developing the Experimental Protocol

We decided to use the Francis method as the basis of our experimental

protocol.57 In essence, we wanted to convert the terminal amines on collagen molecules into ketones, then react these ketone-collagen molecules with a surface decorated with oxyamine functionalities. From there, we wanted to use a fibrillogenesis method to grow

fibrils that stand perpendicularly from the surface. To begin, oxyamine functionalities

needed to be added to the titanium surfaces.

Step 1: Silanization of Titanium

The end-goal of this line of experimentation was to have a monolayer of

oxyamine-terminated silanes on the titanium surfaces, as this would facilitate an even

distribution of collagen and eventually fibrils. To determine the efficacy of the various

silane conjugation chemistries tested, we used X-ray photoelectron spectroscopy (XPS)

to study the atomic composition of titanium before and after silane functionalization.

We focused on nitrogen, carbon, and silicon because these elements exist in the silane

molecules but are not present in appreciable amounts on clean titanium surfaces, which

are mainly comprised of titanium and oxygen. The main peaks are oxygen (530 eV),

titanium (460 eV), nitrogen (400 eV), carbon (280 eV), and silicon (100 eV).

We received silicon wafers coated with 200 nm of titanium from NASA and

diced the wafers into 1 cm square titanium slides. The slides are covered with a rutile

titanium oxide. To functionalize the titanium slides, we decided to use a silanization 31

protocol as the protocols are well documented in the literature.79,81 The Francis protocol

calls for PLP-modified protein molecules to be reacted with oxyamine functionalities,

but there were no U.S. vendors with oxyamine-terminated silane molecules in stock.

While searching for a suitable oxyamine-terminated silane molecule, we moved forward

with a different silane molecule known as 3-(aminopropyl)triethoxysilane (APTES) to

develop the silanization procedure. APTES is an inexpensive, readily available amine-

terminated silane molecule that is well documented in the literature. APTES has been

used to make organosilane mono- and multi-layers on a large number of oxide surfaces

such as silicon crystals,86–90 glass,80,87,91,92 quartz, 93 and various forms of titanium

dioxide 94–96. We decided that we would start attempting our silanization procedure with

O NaBH3CN H N N + N H C C C

Figure 2.2: Amine-carbonyl reaction with cyanoborohydride reduction. Reversible iminium formation between carbonyl and amine groups followed by sodium cyanoborohydride reduction to form a stable C-N linkage. We originally tried to use this reaction scheme to bind ketone-collagen to APTES, an amine-terminated silane.

APTES for two reasons. One, we would not have to waste a more exotic oxyamine-

silane molecule when trying to develop the protocol. Two, carbonyl and primary amine

groups can be reacted to form covalent bonds. Carbonyls and amines react to form

unstable and reversible imine bonds. Sodium cyanoborohydride can then be used to

reduce this unstable double bond, stably linking the original carbonyl- and amine-

containing molecules. A diagram of this reaction scheme can be found in Figure 2.2.97 If

we could move forward using APTES as our silane molecule, it would be less

expensive. Since we did not have immediate access to the equipment needed to do gas-

phase silanization, we went along with solution-phase chemistry to develop the silanization protocol. 81

Before the silanization step can take place, the titanium slides need to be cleaned

and functionalized with hydroxyl groups. First, the titanium slides are cleaned via

ultrasonication in methanol, ethanol, isopropanol, and deionized (DI) water in that order.

In the first protocol tested, the slides were left to dry in the 37℃ incubator, but given the

appearance of dirty spots (non-reflective areas) on the slides, we started removing excess DI water from the slides using a strong stream of dry nitrogen before drying in

the incubator. Slides are dried with nitrogen run through a 7-micron at 10 kPa streamed

through the end of a P-1000 pipette tip for approximately 20 seconds until the majority

of the water has either left the slides or evaporated. This process removes most of the

water from the slides, so any dissolved constituents do not dry onto the slides, leaving

them visibly reflective and clean following this protocol. After an hour in the 37℃

incubator to ensure evaporation, the slides are treated with a humid argon plasma for 10

minutes at 50 watts of power. This plasma treatment removes any remaining

adventitious hydrocarbons left from the cleaning process and decorates the surface of

the titanium slides with hydroxyl groups (-OH). We did not use XPS to assess the

plasma treatment because XPS is unable to detect the presence of hydrogen on surfaces

and because the TiO2 oxide layer on the titanium already contains a large quantity of

oxygen. Instead, we did a basic hydrophilicity test to determine if the plasma

functionalization was successful. We expect that a successfully hydroxylated surface

would be more hydrophilic than a pre-treatment surface as the surface hydroxyl groups

should be charged in water. We did not have access to a contact angle measurement setup, so we moved forward by assessing the shape of water droplets on treated and untreated surfaces as outlined in Figure 2.3. Before the cleaning procedures, a drop of

water (50 µL) placed on the sides would not spread out and would have a visibly large

contact angle. This is indicative of a hydrophobic surface. After cleaning but before

plasma treatment, the slides would still show a large contact angle to the eye. Following

plasma treatment, a drop of water placed on the slides would spread to cover the entire

slide and show a visibly small contact angle, indicative of hydrophilicity. Though we

expect this result to appear due to hydroxylation, this change in hydrophilicity may be

due only to the removal of adventitious hydrocarbons dirtying the surfaces. Since we did

not assay this directly, we moved on to the silanization procedure. We suspected that a

successful silanization procedure would be indicative of a successful hydroxylation, as

hydroxylation is necessary for silanization.

We first tried a conventional silanization technique used by Golas et.al on the

recommendation of Prof. Chris Siedlecki.98 In this protocol, silanization is carried out in a petri dish by a 20 min reaction of clean glass (or a titanium-oxide slide) with 95:5 v/v ethanol-water solution with 5% APTES that had been hydrolyzed overnight in the ethanol-water solution, followed by copious rinsing with ethanol.97 The slides are then

dried under a strong stream of nitrogen followed by 2 hours incubation at 37℃ in

preparation for XPS. The resulting slides are matte and dirty-looking to the eye,

Figure 2.3: Assessing plasma treatment by hydrophilicity. (left) a titanium slide prior to humid argon plasma treatment. 50 µL of water does not spread on these slides and has a large contact angle with the surface. (middle) plasma treatment. (right) a titanium slide following plasma treatment. 50 µL of water spreads across the slides and has a small contact angle with the surface.

indicating that a thick layer of some substance had been deposited on the slides.

Additionally, the slides are hydrophobic, which is unexpected given that the amine functionalities should be charged in DI water. These slides also show large nitrogen and carbon Figure 2.4: XPS spectra of a titanium slide exposed to peaks under XPS, and these peak hydrolyzed APTES (top purple spectrum) and a titanium slide exposed to non-hydrolyzed APTES (bottom green spectrum. The bottom spectrum was taken on a visually “clean” area on heights and corresponding atomic the slide. The N1s peak near 400 eV binding energy and the C1s peak near 300 eV binding energy (both highlighted in yellow) percentages varied wildly between are substantially larger on the hydrolyzed APTES slide compared to non-hydrolyzed slide. batches. Typically, carbon and nitrogen had atomic percentages of ~40 and ~6, respectively. Carbon varied between 30 -45%, while nitrogen varied between 2-7%. An

example spectrum of these slides is shown as the top purple spectrum in Figure 2.4.

Given the appearance of the slides, we hypothesized that the hydrolysis step was causing bulk aggregation of the silane molecules which were then deposited on the surfaces of the slides, leading to these results. Thus, we decided that removing the hydrolysis step might lead to monolayer conjugation.

Next, we tried to take advantage of the amine-terminated silanes and performed the reaction without adding any water to the reaction mixture and omitting the 24-hour hydrolysis step. As shown in Figure 1.6, amine-terminated silanes can perform amine- catalyzed silanization reactions with hydroxyl groups. The slides are cleaned and

hydroxylated using the sonication and plasma procedures, placed in a petri dish, then

exposed to a fresh 5% solution of APTES in ethanol for 10 minutes.81 The slides are

then rinsed thoroughly with ethanol followed by DI water. Following this protocol,

slides are still matte, dirty looking, and hydrophobic. Though the slides did look cleaner

using this protocol than the last, it was again hypothesized that more than a monolayer

of substance had been deposited on the slides. Similar to the hydrolysis tests, these

slides also had varying XPS spectra between samples, but also had “clean” and “dirty”

areas spotted around the slides. Clean areas were low in both nitrogen and carbon, while

“dirty” areas had spectra similar to hydrolyzed slides. In Figure 2.4, the lower green curve shows an example of a “clean” spot’s spectra. The “dirty” spots had similar curves to the upper purple spectra seen on hydrolyzed surfaces. Both of these first protocols also made use of Petri dishes, which require the use of a large quantity of APTES and ethanol and also led to poor control over the experimental conditions, in that these slides were often being exposed to the ethanol-air interface, the air and to ambient humidity.

We hypothesized that exposure to air and humidity was causing bulk aggregation and

adhesion of the silane molecules to the surfaces. Additionally, adventitious

hydrocarbons might have been collecting on the surfaces, masking nitrogen that may

have been present from the silane molecules. Sample-to-sample and within-sample

variability were likely also due to this lack of control. Thus, I designed 3D-printed

polycarbonate holders that would allow for controlled fluid exchange during the

silanization process and any subsequent steps.

A 3D model of the holders along with an image of the holders in use can be

found in Figure 2.5 and more details, including an engineering drawing, can be found in

Appendix B. The 3D-printed holders can hold 2.3 mL of fluid. They have an indentation at the bottom that allows the titanium sides to remain under 300 µL of solution when 2 mL are removed. These holders allow for controlled fluid exchange so that the slides are never exposed to the surface of the ethanol solution nor are they exposed to the air.

Figure 2.5: 3D-printed fluid exchange holders. (left) CAD model of a holder on its side. The fluid chamber has an inset at the bottom to hold 1-cm square titanium slides. When fluid is removed from the large chamber, 300 µL of fluid will remain on the slide, preventing the surface from being exposed to air. (right) Image of a holder being held by tweezers over a beaker during a fluid exchange process. A piece of titanium is sitting in the inset. These holders are used during the entire chemical process of building a synthetic ligament on titanium.

A diagram showing the fluid exchange process can be found in Figure 2.6. The holder (yellow) can hold 2.3 mL of fluid (blue) over a titanium slide (gray). First, A titanium slide is placed in a divot at the bottom of the holder. 2.3 mL of fluid fills the holder. Second, A pipette tip is inserted into the fluid and rests just above the upper platform to withdraw fluid. Only 2 mL of fluid can be withdrawn from the holder when the tip is in this position. Third, 300 µL of fluid remains in the holders, covering the titanium slide. Fourth, A pipette tip is placed directly over the titanium slide and 2 mL of fluid is added into the holder. The surface can be thoroughly sprayed to facilitate mixing when the tip is in this position. This process can be repeated many times to facilitate a thorough rinsing.

Figure 2.6: Clockwise diagram showing the fluid exchange process with 3D-printed holder cross-sections. The holder (yellow) can hold 2.3 mL of fluid (blue) over a titanium slide (gray). 1: (Top left) A titanium slide is placed in a divot at the bottom of the holder. 2.3 mL of fluid fills the holder. 2: (Top right) A pipette tip is inserted into the fluid and rests just above the upper platform to withdraw fluid. Only 2 mL of fluid can be withdrawn from the holder when the tip is in this position. 3: (Bottom right) 300 µL of fluid remains in the holders, covering the titanium slide. 4: (Bottom left) A pipette tip is placed directly over the titanium slide and 2 mL of fluid is added into the holder. The surface can be thoroughly sprayed to facilitate mixing when the tip is in this position.

The holders are cleaned using a different procedure than the titanium slides.

They are cleaned by scrubbing with a plastic brush soaked with Alconox soapy water, and sonication in cold ethanol followed by DI water. The holders are subjected to the same nitrogen drying procedure as the Ti slides and are then placed in the 37℃ incubator for one hour. Polycarbonate is not stable in methanol at room temperature and is only stable in ethanol below 40° C. The sonicator heats up during the cleaning process, hence the omission of methanol and the use of cold ethanol. Slides treated with

5% APTES in ethanol using these holders did show, on average, 3% nitrogen peaks and

25% carbon under XPS, and the batch-to-batch variability decreased. However, these

slides were still hydrophobic. We hypothesize that the bottled ethanol we were using was contaminated with water, adventitious hydrocarbons, and potentially other “dirty” chemicals that deposited on the surfaces. These chemicals may have prevented silanization of the surfaces and/or masked some of the nitrogen that was being deposited on the surfaces. Thus, we switched to an anhydrous 200 proof ethanol with a syringe- accessible seal to reduce the possibility of bulk gelation and adventitious hydrocarbons dirtying the sites.

Following the same titanium cleaning and hydroxylation procedures initially developed, we exposed the slides to a fresh 5% v/v solution of APTES in

200 percent ethanol for 10 minutes, followed by a copious rinsing with ethanol. This final protocol is deemed successful. We obtain clear, repeatable, 5% nitrogen and 25% carbon peaks using XPS, the slides look reflective following silanization, and are hydrophilic. Figure 2.7: XPS spectra of APTES titanium slides with positive results. This XPS profile, compared to that XPS spectra of a clean titanium slide without surface modification (bottom blue spectrum) and a titanium slide modified by surface addition of APTES (top red spectrum) of a clean slide, can be seen in using 200-proof ethanol. The N1s peak near 400 eV binding energy and the Si2p peak near 100 eV binding energy (both Figure 2.7. We considered this to be highlighted in yellow) are substantially larger on the oxyamine-silane functionalized slide compared to the unmodified clean slide. The inset shows a magnified view of a successful protocol that would be the N1s peak. used on future APTES and oxyamine-silane slides.

We found a company in France called SikèMia that did have an 11-carbon oxyamine-silane, 11-(O-hydroxylamine)undecyltriethoxysilane, in stock. More information can be found in Appendix C. We used the APTES procedure using this oxyamine molecule and got successful results. We see clear, repeatable, 2% atomic percent nitrogen on the surfaces of these slides, and the slides are reflective and hydrophilic following the silanization step. An example

XPS spectrum of an oxyamine slide compared to that of a clean slide can be found in

Figure 2.8. We see larger carbon peaks for oxyamine slides compared to APTES slides, but this is expected because APTES is a three- carbon molecule while the

Figure 2.8: XPS spectra of a clean titanium slide without surface oxyamine-silane is an 11- modification (bottom blue spectrum) and a titanium slide modified by surface addition of oxyamine-silanes (top red spectrum). carbon molecule. The N1s peak near 400 eV binding energy and the Si2p peak near 100 eV binding energy (both highlighted in yellow) are substantially larger on the oxyamine-silane functionalized slide compared to the The N1s peak located at unmodified clean slide. The inset shows a magnified view of the N1s peak along with the best fit curve used to determine peak position 400.13 eV on the activated and area. surface is consistent with an oxyamine-containing molecule according to work by

Yoshida et. al. documented in the NIST X-ray Photoelectron Spectroscopy Database.99

Qualitatively, N1s and Si2p both show noticeable peaks after the conjugation reaction

while little to no intensity is seen in the control spectrum. The insert at the top right of

Figure 2.8 shows that the size of the nitrogen peak is larger for the oxyamine activated surface than it is for the clean titanium, further supporting the hypothesis that oxyamine functional groups were successfully conjugated to the titanium surface. Areas under the peaks were used to determine the relative atomic percent of each element. Nitrogen accounts for approximately 2% of the atoms on the oxyamine activated surface.

Assuming that the oxyamine molecules are evenly distributed across the titanium and that the titanium surface has a tetragonal rutile crystal structure, 2% nitrogen equates to

Figure 2.9: Scheme of oxyamine-silane surface chemistry. Binding oxyamine-terminated silane molecules to a titanium surface using a solution phase, amine-catalyzed reaction. The reaction is carried out using anhydrous 200-proof ethanol in unique fluid-exchange holders.

one oxyamine functional group every 4.6 nm on the surface of a (0,0,1) plane.100 Typical

reconstituted collagen fibrils are generally larger than 10 nm in diameter. Thus, this

spacing of collagen monomers is sufficient to allow for multiple covalently bound

monomers to be associated with each fibril grown on the surface. Figure 2.9 shows a

schematic of the oxyamine-silane conjugation reaction.

Step 2: Ketone-Collagen Production

Dr. Jon Pokorski provided us with our original PLP protocol, based on the

Francis protocol from 201059, as the Pokorski group had previously found success in

modifying EGF.73 The PLP protocol we used involves mixing the protein with 100 mM

PLP in pH 6.5 phosphate-buffered saline (PBS) and incubating at 37℃ for two hours,

followed by centrifugal filtration against PBS to remove the excess PLP. I used this protocol on 0.5 mg/mL of PureCol collagen. We used a UV-visible dye and UV-visible

spectroscopy (UV-vis) to assay the collagen molecules for ketone functionalities

following this PLP reaction. The UV-visible oxyamine dye, aminooxy-5(6)- tetramethylrhodamine (TAMRA), is expected to site-specifically react with ketones in a

“click chemistry” fashion. Following a 24-hour conjugation step in pH 5.5 acetate buffer, the unbound TAMRA dye was removed via extensive centrifugal filtration against PBS until no purple absorbance was detected in the flow-through. Ultraviolet- visible spectroscopy (UV-vis) was then used to ascertain the yield of the transamination chemistry performed on the collagen monomers. Collagen has an absorbance at around

230 nm, while TAMRA has an absorbance at both 230 and 560 nm. UV-absorbances of the TAMRA dye and the collagen molecules were both measured alongside known concentrations of each molecule allowing the measured sample absorbances to be

converted to concentrations. The TAMRA’s contribution to the 230 nm absorbance was

accounted for when calculating the collagen content of the sample solutions. More

information can be found in Appendix E. Based on the UV-vis data, we determined that

using this PLP protocol the first time, each collagen molecule was bound to an average

of 3.5 TAMRA dye molecules. At the time, we deemed this reaction a success, as we

wanted to see approximately three TAMRA dye molecules (and therefore ketones) per

collagen molecule given collagen’s three N-termini. However, when doing this first

reaction, we failed to take into account the N-terminal sequences of the collagen

molecules we were modifying. Since one of the three alpha chains of atelocollagen ends

in isoleucine (essentially non-reactive to PLP) while the other two end in aspartic acid

(reactive), we should have expected to see, at most, two ketones per collagen molecule as discussed in section 1.6.65

Though we can now speculate that there was an issue with the original PLP-

collagen reaction, at the time we were happy that we saw any TAMRA conjugated to the

collagen molecules. As such, we then attempted to conjugate the first-attempt PLP-

modified ketone-collagen to APTES surfaces using the sodium cyanoborohydride

procedure in Figure 2.2. To assess the efficacy of this amine-carbonyl reaction, we

initially used FTIR on the surfaces and looked for the amide II band as evidence for

collagen. The amide II band, between 1510 and 1580 cm-1, results from the N-H

bending vibration and from the C-N stretching vibration. Both of these bond types are highly prevalent in collagen.58 As shown in Figure 2.10, we were unable to see clear peaks within the amide bands (highlighted) in the IR spectra of slides exposed to PLP-

collagen compared to controls. To compare, a clear peak within the amide II band can

Figure 2.10: FTIR spectra of titanium slides. Slides were exposed to just our APTES silanization procedure (left) and to the APTES silanization followed by PLP-modified collagen. The amide II band is highlighted in both images. No amide II peaks could be seen on the surfaces exposed to ketone-collagen. be seen in Figure 2.11, which is the FTIR spectrum of a sample where PureCol collagen was left to dry on a titanium slide. We hypothesized that FTIR was not sensitive enough to pick up low surface concentrations of collagen and that even if we did see peaks after changing the protocol, we would not know if those peaks were indicative of collagen.

Thus, we decided that we needed a more collagen-sensitive analysis technique.

Following the inconclusive FTIR results, we decided to use antibody staining procedures to look for collagen on the sides. We use a primary anti-bovine type I collagen antibody followed by an FTIC-labeled secondary antibody. We expected that surfaces exposed to both the silane and collagen chemistry would fluoresce more brightly under 494 nm light than control slides (clean or APTES-only slides) after staining, but we were unable to see such a difference. All slides showed mean grayscale values of around 10-30, and the slides were visibly black. We hypothesized that the sodium cyanoborohydride reduction was not sufficient to link the collagen molecules to the APTES surfaces, indicating that we did need to use an oxyamine functionalized

silane group as the Francis articles

state.59,62 Then, we attempted a reaction between the previously made

PLP-modified collagen molecules and oxyamine functionalized surfaces. We still saw no appreciable fluorescence following antibody staining on these

slides compared to controls. After Figure 2.11: FTIR spectrum of a titanium surface with a dried spot of PureCol collagen. A 10 µL droplet of 3 mg/mL PureCol was left to dry at many attempts at conjugating the PLP- room temperature overnight. The peak labeled 6 within the highlighted Amide II band is indicative of protein on collagen to both APTES and oxyamine the Ti sample surface.

slides, I had run out of my stock solution of PLP-collagen. As such, I needed to make more.

Using the original PLP protocol again, I was unable to replicate the original results. I tried repeating the PLP reaction and the TAMRA dye process several times but found no evidence that the TAMRA was binding to the collagen molecules. We first used a plate reader to assay for the 560 nm absorbance of the TAMRA-dyed collagen solutions following an extensive centrifugal filtration process to remove any unbound

TAMRA. Unable to see any appreciable 560 nm absorbance, we moved to use the

NanoDrop 1000 UV-vis spectrophotometer. The NanoDrop requires only 2 µL of protein solution, whereas the plate reader requires 200 µL and requires the solution to be diluted to a protein concentration less than 7 mM (~2 mg/mL). Appendix E goes into more detail regarding this change and the general UV-vis procedure. Despite using the

NanoDrop, we still saw no 560 nm absorbance in TAMRA-treated sample solutions. I

then thought that if our reaction efficiency was very low, there may be very little bound

TAMRA in the system. Collagen has a molecular weight of 300 kDa, while the TAMRA dye molecule has a molecular weight of just 815.71 Da. I then concentrated the samples via lyophilization and rehydration in 10 µL of fluid, increasing the collagen concentration from 10 mM to 330 mM. If there was any appreciable TAMRA in the system I expected to see it after this 33x concentration, but still, no 560 nm absorbance could be found.

Upon closer examination of the PLP reaction solution, I found that a gel had been forming at some point during the PLP-collagen transamination reaction. Figure

2.12 shows an image of such a collagen gel being removed from a reaction vessel with a

Figure 2.12: PLP gel mass. Figure 2.13: PLP gels on a pipette tip. Collagen gel (circled in red) pulled Picture of a pipette tip after gentle use out of a PLP-PBS solution with a in a gelled PLP-Collagen solution. syringe following the original PLP Note the droplets of clear substance transamination procedure. on the tip. Those are droplets of collagen gel. syringe following a 1-hour incubation period. These delicate gels break apart with very little force, and as such agitating the fluid with a p-1000 or P-100 micropipette would cause the gel to fall apart into droplets. These droplets, shown on a pipette tip in Figure

2.13, could be mistaken as fluid thus disguising the issue in previous tests. Since we

wanted to conjugate individual collagen molecules to a surface, we needed all of the

collagen to be in solution. I hypothesized that this gelation was either preventing the

transamination reaction from happening by making most of the N-terminal amines

unavailable, or that the gelled collagen was undergoing the N-terminal transamination

but that these groups were made unavailable to the TAMRA dye by the gelation, leading

to the lack of 560 nm absorbances in previous solutions. Concerned that this collagen

gelation would interfere with the transamination procedure and subsequent surface-

conjugation reactions, we started to modify the original Francis procedure in an attempt

to replicate our original results and prevent gel formation.

Collagen is known to aggregate into fibrils in the presence of 137 mM sodium

chloride and 10 mM phosphate ions (such as in PBS) at pH 7 and 37℃, forming gel

masses.47. D-banded fibrils can also form with NaCl concentrations ranging from 50 to

500 mM50 and phosphate concentrations ranging from 10-50 mM,51,52 as discussed in

section 1.4. My first hypothesis was that the PBS buffer used during the transamination

reaction was causing collagen gelation similar to a fibrillogenesis process. We replaced

the PBS buffer with a 30 mM 2-(N-morpholino)ethanesulfonic acid (MES) buffer with

150 mM sodium chloride but still found that the collagen would gel in the 100 mM PLP

reaction solution. In fact, through this battery of tests, I found that the gels were forming

within 30 seconds of mixing the collagen with the reaction solution. We then tried

reducing the salt concentration, as salt causes charges on the protein to be screened so

that their forces of interaction fall off exponentially, rather than at 1/distance2. This

effect allows charged collagen molecules to get close enough to aggregate via hydrophobic interactions.101 I tested reaction solutions using 133, 100, 50, 10, and 0 mM

sodium chloride and continued to find that gels would form within 30 seconds of adding the collagen no matter the salt concentration. The MES is a salt as well and therefore could have been participating in the screening, but a buffer is necessary to maintain the pH of the reaction solution. I knew I needed to use a buffer, so I then suspected that the

MES buffer was inappropriate for use with collagen just as the PBS was. I tried similar buffers using Tris and HEPES that had also been used in PLP transamination procedures,62 but still found that the collagen would gel on-contact with the solution.

My next suspicion was that the PLP itself was causing this gelation. Figure 2.14 shows the chemical structure of PLP, a phosphate-containing molecule. I suspected that the PLP may have been participating in a fibrillogenesis process with the Figure 2.14: Chemical structure of pyridoxal 5'-phosphate (PLP). collagen molecules, acting as a phosphate ion. To test the idea, I made solutions with 30 mM MES buffer at pH 6.5, omitting the PLP, and found that the collagen gels would no longer form on-contact with the solution. This test strengthened my suspicion that the PLP itself was the blame for the gelation. I then tried many varying concentrations of PLP from 100 to 5 mM, below the normal fibrillogenesis phosphate concentration of 10 mM, but gels would still form quickly upon mixing. As discussed in Table 1 from section 1.4, there are a large number of solution conditions that lead to the formation of collagen aggregates that are not D- banded fibrils. For example, Uquillas et. al. found that fibrous collagen structures lacking D-banding would form in solutions containing about 6 mM of phosphate and 70 mM of NaCl.102,103 Though we omitted the NaCl in these reactions, the low 5 mM

concentration of PLP may have been participating in the gelation as free phosphate just

as Uquillas et. al. showed.

It is possible that the original positive TAMRA results we obtained using PLP

were the result of inadequately rinsing the excess TAMRA dye molecules from the

collagen solution, and that the PLP procedure did not result in N-terminal ketone

functionalization due to this gelation issue.

Given that the PLP seemed to be initiating

collagen gelation almost immediately on contact

with a PLP-transamination solution, I decided to

use N-methylpyridinium-4-carboxaldehyde

benzosulfate, otherwise known as Rapoport’s salt

(RS), as the transamination reagent. The chemical Figure 2.15: Chemical structure of N- methylpyridinium-4-carboxaldehyde structure of RS can be found in Figure 2.15, where benzosulfate salt, A.K.A. Rapoport's salt.

the N-methylpyridinium-4-carboxaldehyde

participates in the transamination reaction. The

Francis group originally introduced RS as an

alternative to PLP for the modification of

glutamate-terminated amino acid sequences,

particularly for IgG antibodies.75 In my case, I

decided to switch to RS because it has been used to

Figure 2.16: SDS-collagen gel. transaminate proteins with similar N-terminal Image of a dialysis cassette filled with a cloudy collagen gel in the lower corner. sequences as one of collagen’s alpha-I chains and This collagen solution was dialyzed against SDS following an RS i i i also because it does not contain a phosphate group

as PLP does. Lightly pepsin digested type I collagen displays three different triplets on

the three alpha chains making up the molecule: ISV, DEK, and DAK.77 Witus et al.

showed that groups with a terminal I, isoleucine, show almost no reactivity to

transamination.59 This allows us to rule out the chain ending in ISV as a transamination

target. They did not specifically test DEK or DAK. However, they did show that DES

(similar to collagen’s DEK) had a conversion efficiency near 50% 75,76 Combining 0.5 mg/mL of collagen molecules with a 100 mM solution of RS in 30 mM MES (pH 6.5)

and 37℃ for one hour, I did not find any more gels in the reaction vessel. Gels would not form within 30 seconds of the mixing process, nor would they form during a 2-hour incubation period. In fact, no gel would form during a 24- or 120-hour incubation period either. However, upon centrifugal filtration against 30 mM MES buffer, I found that the collagen would once again gel following a 1-hour RS reaction. I first suspected that the centrifugal filtration procedure used to remove excess RS was too aggressive, causing the collagen molecules to denature and gel.

I attempted to use a gentler 24-hour dialysis procedure as a replacement for centrifugal filtration but found that the collagen solution would still gel inside the

membrane during the RS removal process. Given that this gentler method did not prevent gelation, and that the collagen only gelled once the RS was removed from the system, I then suspected that the RS was somehow keeping the collagen

molecules in solution following the transamination

procedure. I continued to use dialysis, thinking that the Figure 2.17: Chemical structure of centrifugal filtration could still be playing a part. I 3-(1-pyridinio)-1-propanesulfonate, a “bulky” salt with a similar chemical structure to RS.

attempted to dialyze against 5 mM sodium dodecyl sulfate, thinking that a surfactant

would prevent the collagen molecules from aggregating, but gelation still occurred and

lead to more opaque gels (Figure 2.16). I think that the SDS caused the collagen to

denature. Then, thinking that the bulky ring structure on the RS was physically getting

in the way of collagen aggregation, I also tried to dialyze the collagen solution against a

100 mM bulky salt solution, 3-(1-pyridinio)-1-propanesulfonate (Figure 2.17). Gelation still occurred during the RS removal process using this bulky salt.

Finally, we tried a chaotropic agent. When reading the literature regarding the solubility of proteins, we found papers describing kosmotropic and chaotropic agents.

These are molecules that are known to strongly affect the water solubility of large molecules via their effect on the ordering of water molecules.104 Kosmotropes are

described as “water-structure makers”. They are small ions with a high charge density

that bind water molecules tightly and increase the surface tension of aqueous solutions.

These agents stabilize protein molecules and decrease their solubility, exhibiting a

- 2- phenomenon called salting-out. Examples of kosmotropes are F and SO4 . Conversely,

chaotropes (“chaos-makers”) are large ions with a low charge density that do not bind

water, but rather increase the mobility of nearby water molecules. In this way, chaotropes destabilize proteins and increase their solubility (salting-in). High concentrations of chaotropic molecules can cause proteins to denature, but smaller concentrations can increase the solubility of said proteins without destabilizing their 3D-

2+ - 2+ structure. Mg and ClO4 are examples of a chaotropes. We chose to try Mg in the

form of MgCl2 to prevent the aggregation of our ketone-collagen molecules following the removal of the RS from the reaction solution. We chose magnesium partly because

we had it on hand, but also because Mg2+ is a divalent cation, covered next.

The Douglas group found that when collagen monomers are put into a solution

containing a divalent cation, the isoelectric point (pI) of the monomers increases.105 For

example, the pI of collagen in 12 mM NaCl is 8.9. It shifted to 7.0 after adding 10 mM

Na2SO4 [divalent anion] and conversely shifted to 9.4 when after adding 10mM CaCl2

[divalent cation]. The isoelectric point of a protein refers to the pH where the net charge of the molecule becomes zero, thus reducing electrostatic repulsions among identical

proteins. As such, by the addition of a divalent cation, one can increase the pI of a

collagen molecule to well above the reaction conditions of pH 6.5, increasing

electrostatic repulsions among molecules and therefore preventing aggregation.

Additionally, Mg2+ has a weak binding ability to soluble collagen.106 This combination

of information suggests that Mg2+ binds with collagen monomers, making them more

positively charged at lower pH values than they would be otherwise. The Douglas group

found that 150 mM of MgCl2 completely inhibits fibril formation at pH 7.4 and 37℃,

105 while 350 mM of MgCl2 caused collagen denaturation. These effects may be due to

the combination of the chaotropic and divalent cation characteristics of Mg2+.

In our procedures, the addition of 150 mM MgCl₂ to the rinsing solution

prevented the formation of gels and therefore allowed for the production of solution-

phase ketone-collagen molecules using 100 mM RS. Figure 2.18 shows the reaction

scheme of this modified RS transamination procedure.

Figure 2.18: Conversion of N-terminal amines of collagen using N-Methylpyridinium-4-carboxaldehyde as a transamination agent.

We assessed the efficacy of this modified reaction by the same TAMRA analysis described at the beginning of this section, replacing all rinsing solutions with the aforementioned MgCl2 solution. Figure 2.19 shows the UV-spectra of unmodified and ketone-modified collagen molecules. The peak at around 560 nm in the modified collagen curve shows the existence of TAMRA bound to these molecules. The concentrations of TAMRA and collagen in the reacted solution gave a ratio of TAMRA to collagen of approximately 1 to 10. Some ambiguity remains regarding what percentage of the collagen monomers contain ketone end groups. Each collagen molecule consists of three alpha chains, each of which might have been converted to a ketone. Thus, each collagen molecule has the potential to bind up to three TAMRA molecules. This gives a range of converted collagen monomer efficiency somewhere between 3.3% (assuming all three alpha chains were converted) and 10% (assuming

only one alpha chain per collagen

molecule was converted). Based on

previous work, this range of efficiencies

can be reduced. Lightly pepsin digested

type I collagen displays three different

triplets on the three alpha chains making

up the molecule: ISV, DEK, and DAK.

Figure 2.19: UV-vis spectra of collagen solutions. Witus et al. showed that groups with a The TAMRA-labeled ketone-collagen solution (black) shows a peak at ~560 nm, indicative of TAMRA, and terminal I, isoleucine, show almost no ~230 nm, indicative of collagen. The unmodified collagen (red) only shows a peak at ~230 nm for collagen. reactivity to transamination.59 This allows

us to rule out the chain ending in ISV as having been converted to contain a ketone.

They did not specifically test DEK or DAK. However, they did show that DES had a

conversion efficiency near 50% while DIQ was near 0%. Thus, we cannot say whether

one or both of the two chains with a terminal D, aspartic acid, were converted to

ketones. Our conclusion, then, is that somewhere between 5% and 10% of the collagen

molecules in our initial solution were converted to ketone-collagen. The stock ketone-

collagen solutions are stored at a concentration of ~4 mg/mL in the -80℃ freezer to hinder collagen aggregation or denaturation and to retain the ketone functionalities.

Step 3: Binding Ketone-Collagen to Oxyamine-Silanized Titanium

Following the successes of the ketone-collagen production and oxyamine- silanization steps, we then moved on to combining these two components. We did not attempt a cyanoborohydride-reduction reaction between the ketone-collagen and the

APTES surfaces. Oxime bond formation between 0.4 mg/mL of modified collagen molecules and the oxyamine-functionalized surfaces is carried out in a 30 mM MES buffer at pH 5.5 for 17 hours, following the Francis procedure.75 Aggregation of the modified collagen molecules is prevented by the addition of 150 mM MgCl₂ and by the omission of PBS, as in the transamination procedures. We had little trouble getting this reaction to work once the silane-surface chemistry and collagen-modification chemistry had been developed. A stable oxime linkage is formed between the oxyamine functionality on the silane molecules and the ketone functionality on the collagen molecules. Figure 2.20 shows a diagram of this reaction and the final surface-bound collagen.

This reaction can be assayed by the use of collagen-specific FITC- tagged antibody staining procedures discussed previously, the results of Figure 2.20: Oxime linkage formation between surface-bound oxyamine- silane molecules and ketone-modified collagen N-terminus. which are outlined in Figure This process results in individual collagen molecules covalently bound to a titanium surface. 2.21. In the figure, the

topmost histogram comes from a titanium slide that only underwent oxyamine chemistry

and was not exposed to any collagen. The image of the slide, shown to the right of this

histogram, appeared black to the eye and quantitatively showed a peak grayscale value

of 14. The middle histogram comes from a titanium slide that underwent oxyamine

chemistry and was subsequently exposed to unmodified collagen molecules directly

from the PureCol solution. Similar to the surface exposed to no collagen, the slide

(shown to the right) appears black to the eye and displays a histogram with a low mean

grayscale value of 28. The bottommost histogram comes from a titanium slide that

underwent oxyamine

chemistry and was

subsequently exposed to

our RS-modified ketone-

collagen molecules. The

field of view looked

noticeably brighter with

a generally uniform

contrast, shown in the Figure 2.21: Histograms of immunofluorescence grayscale values for the detection of collagen molecules on activated titanium surfaces. (top) Titanium slide that only underwent oxyamine chemistry and was not image to the right of the exposed to any collagen. The image appeared black to the eye and quantitatively showed an intensity histogram with a peak at a grayscale value of 14. (middle) Titanium slide that underwent oxyamine chemistry and histogram, and had a was subsequently exposed to unmodified collagen molecules. Similar to the surface exposed to no collagen, the image appears black to the eye and mean grayscale value of displays a histogram with a very low mean grayscale value of 28. (bottom) Titanium slide that underwent oxyamine chemistry and was subsequently exposed to modified collagen molecules. The field of view looked noticeably 3 9.8×10 . This value is brighter with a generally uniform contrast. Scale bars are 5 microns long. Quantitatively, the histogram of image intensity gray scale values had a 3 nearly 1000X greater mean of 9.8*10 , nearly 1000X greater than the control surfaces. than the control surfaces. All images shown have scale bars that are 5 microns long and

have a window of 0-14,000 grayscale values.

Qualitatively, the control groups appear dark while the treated group is noticeably brighter. The contrast in the treated group looks generally uniform, indicating that surface coverage of the bound collagen molecules was uniform on a length scale commensurate with the optical images. Since very little antibody binding is seen on the unmodified-collagen control slides, it is unlikely that collagen attachment not specific to the oxyamine- (ketone-collagen) reaction occurred. Since the ketone should exist only at the terminus of the molecule75,76 and the oxyamine binding sites take up ~2% of the surface area, a simple steric argument suggests that, on average, the collagen molecular axis of symmetry projects away from the surface at some angle close to the surface normal. Assuming that the bound collagen molecules extend up from the surface at some appreciable angle, they should be available to act as nucleation sites for fibril formation. Despite the low percent nitrogen (2%) observed on the oxyamine-silane surfaces from step 1, the homogeneity and brightness of the antibody-stained collagen- molecule-coated slides suggest that significant decoration of the surface with collagen monomers has occurred. We did not directly assay for covalent bonding between the collagen molecules and the oxyamines. However, the chemical reaction between the terminal ketone and the oxyamine is well known to result in a covalent bond. Also, copious washing was performed after the conjugation chemistry, raising confidence that attachment of the molecules was robust.

While the direct characterization of the oxime bond was not performed, I did attempt to hydrolyze the oxime bonds formed on surfaces by acid exposure. Oxime bonds are stable in near-neutral pH solutions but are unstable in acidic conditions.107

The Pokorski group conjugated doxorubicin (DOX) to polymer scaffolds using an oxime linker, then exposed these surfaces to a pH 4.5 MES buffer at 37 °C for 48 hours.

They found that approximately 60% of the DOX had been released from the scaffold by the 30-minute mark, indicating that about 60% of the oxime bonds had been broken during that time. If the ketone-collagen + oxyamine-silane chemistry described above resulted in oxime bond formation, then I would expect that ketone-collagen surfaces would show collagen-release kinetics similar to the Pokorski group’s DOX experiments.

By that logic, ketone-collagen slides exposed to low pH conditions should have lower collagen content and therefore less antibody binding than those exposed to neutral pH conditions. To test this hypothesis, I exposed two sets of ketone-collagen titanium slides to a pH 4.5 acetate buffer and a pH 7.4 MES buffer at 37°C for one hour (three slides for each reaction condition). I also exposed a single slide to a pH 5.5 MES buffer at 37℃ for one hour. Following exposure, I rinsed the slides thoroughly and antibody stained the slides using the same protocol as above. I looked to see if the fluorescence intensity of the pH 4.5 slides was significantly less than that of the pH 7.4 or pH 5.5 slides.

However, there was no significant difference in the fluorescent intensity of the three pH

4.5 slides compared to the three pH 7.4 or 5.5 slides. Figure 2.22 shows the histograms of fluorescent intensity found on each of these slides. The top three blue graphs show the fluorescent intensity histograms of the pH 7.4 slides, the middle gray graph shows

the fluorescent intensity of the pH 5.5 slide, and the bottom three red graphs show the fluorescent intensity histograms of the pH 4.5 slides. Also displayed are the mean (µ) and standard deviation (σ) of the intensity of each slide, along with the number of pixels (n) used to generate the histograms.

These oxime bonds may have different reaction kinetics than the DOX experiment, and this is not conclusive evidence that no oxime bond was formed.

Figure 2.22: Histograms of fluorescent intensity found on the acid- This oxime bond formed hydrolyzed slides. The top three blue graphs show the fluorescent intensity histograms of here may be more stable in the pH 7.4 slides, the middle gray graph shows the fluorescent intensity of the pH 5.5 slide, and the bottom three red graphs show the fluorescent intensity histograms of the pH 4.5 slides. Also displayed are the mean (µ) acidic conditions than the and standard deviation (σ) of the intensity of that slide, along with the number of pixels (n) used to generate the histograms. No significant difference can be found in the mean fluorescent intensity of pH 7.4, pH oxime created in the 5.5, and pH 4.5-exposed slides.

Pokorski paper, and

therefore a longer reaction time is warranted. More work, potentially using FTIR or other analysis techniques, needs to be done to confirm that an oxime bond does exist between the ketone-collagen and the silane molecules.

Although this acid hydrolysis experiment did not yield the expected results, it may lend credibility to this chemistry in producing a synthetic ligament. According to a study done by Karpavicius et al. in 2019, the average pH value of the peri‐implant crevicular fluid around a fully healed two‐piece dental implant (implant with abutment) was 6.16 with a range of 5.6–6.6. 108 This environment is more acidic than the average pH value of the gingival crevicular fluid around healthy teeth, which was recorded as

6.64 with a range of 6.2–7.0. Given that my pH 5.5 and pH 4.5 slides did not show any signs of collagen detaching from the surfaces, a synthetic brush of Sharpey’s fibers formed on an implant may be able to withstand the slightly acidic conditions found around healthy dental implants. Given these findings, it would be reasonable to do longer-term in-vitro studies where collagen-functionalized slides are exposed to pH values in the range of 5.6-7.0 to determine the long-term impact of acidic conditions.

Step 4: Nucleation and Growth of Collagen Fibrils on Titanium

Following the conjugation of the modified collagen molecules with the oxyamine-silanes, a controlled nucleation-and-growth fibrillogenesis procedure can take place. Scanning electron microscopy (SEM) was used to determine if collagen fibrils were grown on the modified titanium surfaces. Samples were prepped with ethanol dehydration followed by either air drying or critical point drying. Ethanol dehydration followed by critical point drying increases the chances any fibril standing in aqueous conditions would remain standing once dry. Based on trying to mimic the nanostructure 60

of a biological ligament, our target is to have collagen fibrils 50-100 nm in diameter

with spaces of 10-20 nm between each fibril. A standard fibrillogenesis protocol used in

the Eppell lab is to make a 0.18 mg/mL collagen in cold, pH 7.4 PBS and incubate that

solution overnight at 37℃.47 This procedure results in thin, wispy gels floating in PBS.

At the micron scale, these gels are entangled mats of collagen fibrils. An SEM image of such a disorganized structure can be seen in Figure 2.23. To create a standing brush of artificial Sharpey’s fibers, more control over the fibrillogenesis process is required.

Collagen fibrillogenesis can be broken up into nucleation and growth steps. To generate long fibrils, stable aggregations of collagen molecules called nuclei must first form.45 Once the nuclei are formed, fibrils can then grow longitudinally off of these nuclei, as discussed in section 1.4. Figure 1.4 shows a standard turbidity curve for the self-assembly of collagen molecules into fibrils. We used this knowledge of collagen

Figure 2.23: SEM image of a collagen gel on a titanium slide formed using a standard fibrillogenesis procedure. Each rope-like structure is a bundle of collagen fibrils. This structure is not conducive to producing ligaments, as the fibrils are not anisotropically aligned, nor are they bound end-on to a surface.

turbidity profiles to develop a nucleation-and-growth protocol for building collagen

fibrils on surfaces.

As a starting point, we decided that the fibrillogenesis solution should be made

such that no bulk gels should be visible to the naked eye after a 24-hour incubation at

37℃. This would limit the number of fibrils that form in the bulk solution. A solution of

3 µg/mL of collagen in PBS satisfies this constraint. First, we tried to make collagen fibril surfaces using this low-concentration solution to see if any surface fibrils would form. Using only this low concentration, we only found one example of a standing fibril like the one shown in Figure 2.24C. We hypothesized that adding a nucleation step before the growth step would increase the number of viable fibril nuclei that could form on the surface-bound collagen molecules, thereby increasing the number of standing fibrils on the surfaces. We hypothesized that the nucleation step would require a relatively high concentration of collagen to encourage nuclei to form on the surface- bound collagen molecules, but that a low concentration could be used for the growth phase. In the low concentration solution, fibrils would already be nucleated on the surfaces and be able to readily grow, while nuclei in the bulk would be less likely to form.

The literature reports that the lag phase for type I collagen nucleation can last between three (1.0 mg/ml collagen)109 and ten minutes (0.2 mg/mL collagen)110 at 37℃ and pH 7.5. Our first nucleation protocol involved exposing the surfaces to five washes of a 0.18 mg/mL concentration of collagen molecules in 37℃ PBS. Each solution was

left on the collagen surfaces for five minutes and then promptly washed off with an excess of PBS before the next solution was added. We did multiple collagen-solution

exposures and rinses in an attempt to nucleate as many fibrils as possible without allowing any bulk gelation to settle on the slides. We hypothesized that this period (5-minutes) would result in the formation of few bulk collagen fibrils, given literature reports that 0.2 mg/mL collagen solutions had a 10-minute lag phase at

37℃.110 However, these experiments led to surfaces that looked much like the mat shown in

Figure 2.23. We then tried the same protocol but only exposed the collagen solution to the slides for one minute between rinses, but still saw Figure 2.24: SEM images of titanium surfaces. mat-covered slides. These results did not line up A: clean slide showing no fibril-like formations. B: net of collagen fibrils standing off the surface. C: single fibril standing upright. with the 10-minute lag phase we were expecting.

Upon further research, we found a great discrepancy in reported lag phase times, as each lab used collagen from different sources (rat vs fish vs bovine) with different protocols (warm start vs cold start) at different temperatures, collagen concentrations, and pHs.45,52,109–112 Given that the literature reported such a wide range of lag phase times, and that the literature did not include turbidity curves following our exact procedures, we decided that we needed to create our own turbidity curves to better understand our reaction kinetics. Angela Yu was in charge of this turbidity project.

Through her work, we found that the lag phase of our collagen solutions was typically

much shorter than that found in the literature; a 0.18 mg/mL collagen solution at 37℃ had a lag phase of 30 seconds. We wanted to rinse the slides of excess nucleation solution well before the predicted end of the lag phase to limit the amount of bulk gelation that formed.

Using the holders and the nucleation procedure, I was

Figure 2.25: SEM images of zoomed-in titanium surfaces. physically unable to rinse this The picture in the top right is a clean, unmodified surface. The rest of the image show fibrils making up fibril nets similar to the net in nucleation solution off of the Fig. 5B. Arrows show places where fibrils interface perpendicularly to the surface, indicating end-on attachment. all scale bars are 200 nm long. titanium in less than 30 seconds. I needed the lag phase to last longer than 30 seconds to practically carry out a nucleation procedure. Based on Angela’s research, we found that a 50 µg/mL solution of collagen in PBS at 30℃ had a lag phase of two minutes and 30 seconds. Thus, my new nucleation protocol involved exposing ketone-collagen surfaces to five washes of a 50

µg/mL of collagen solution in 30℃ PBS. The solutions are left on the slides for one minute before rinsing. Using this protocol, five standing fibrils like the one shown in

Figure 2.24C on three different titanium slides could be found under SEM. Much more common were net-like structures shown in Figure 2.24B. These net-like structures show evidence of end-on conjugation of collagen fibrils, shown with yellow arrows in Figure

2.25. I hypothesize that these nets are made of previously standing collagen fibrils that have collapsed onto one another, potentially during sample preparation. The SEM images are consistent with some collagen molecules self-assembling to produce fibrils oriented with their symmetry axis parallel to the titanium surface normal. Only surfaces that have gone through plasma treatment, oxyamine functionalization, ketone-collagen conjugation, and exposure to nucleation-and-growth procedures show such fibrils. All other surfaces look like the surface in Figure 2.24A. Since only our chemistry produces these fibrils, and the fibril ends shown in Figure 2.25 we assume that they are conjugated end-on to the surface.

Summary of Developed Protocol

Figure 2.26 gives a simple overview of the developed protocol and is organized as four steps. The full protocol can be found in Appendix A. In step 1, a plasma-treated titanium surface is functionalized with a monolayer of oxyamine-terminated alkylsilanes. In step 2, the terminal amines of a solution of collagen molecules are converted to ketones using Rapoport’s salt. In step 3, the resultant ketone-collagen is exposed to the oxyamine-functionalized titanium surface resulting in collagen molecules being firmly attached to the titanium. In step 4, collagen fibrils are nucleated around the covalently bonded collagen molecules. These nuclei are subsequently grown into single nanofibrils.

Activation of Titanium Surfaces 1 Activation of 2 Collagen Monomers Clean titanium 11-(O-hydroxylamine)undecyltrimethoxysilane RS O NH2 NH2 Plasma Treat Si O MgCl O 11 O 2 O NH2 NH2 OH OH OH O O Oxyamine bridge molecule Ketone functionality

Combine Activated Single end-on Fibrillogenesis on Titanium 4 Surface and fibril Collagen Monomers

Nucleation Single Collagen 3 Monomers on Ti

Growth

Figure 2.26: Overview of the chemical steps necessary to produce collagen fibrils conjugated end-on to a titanium surface. The chemical process can be broken into four distinct steps: 1) activation and silanization of the titanium surfaces, 2) activation of collagen monomers by n-terminal transamination, 3) conjugation of ketone-collagen monomers end-on to the silanized titanium, and 4) nucleation on the single ketone-collagen monomers followed by growth to produce fibrils.

Chapter 3: Manuscript

3.1 Preface

The following manuscript has been written and edited with the help of Dr.

Jonathan Pokorski and Dr. Steven Eppell. All experiments and analyses related to this

project were performed by me. Dr. Pokorski contributed to the experimental design of

the modified collagen molecules and to the chemistry used to connect these molecules to the oxyamine-functionalized titanium surfaces. Dr. Eppell contributed to the experimental design of the nucleation and growth portions of the fibrillogenesis procedure used to grow fibrils on the titanium surfaces. Angela Yu conducted collagen turbidity experiments to attain the proper nucleation and growth conditions for the fibrillogenesis process. Portions of the text and figures of this manuscript have been

repeated in chapters 1 and 2 of this thesis.

3.2 Manuscript

COLLAGEN NANOSTRUCTURES ON TITANIUM

3.2.1 Abstract

Proof-of-concept is presented for a method to create upright-standing collagen

nanofibrils covalently bonded to a titanium surface. The developed procedure starts with

activation of the titanium surface with a plasma discharge treatment followed by

functionalization with an oxyamine-terminated silane coupling molecule. Using a

Rapoport salt procedure, the N-termini of individual type I collagen monomers are

converted to ketones. When presented to the functionalized titanium surface, these

ketones are expected to form an oxime linkage, immobilizing the collagen. In a two-step

process, these bonded molecules act as sites for the formation of fibrils. Many fibril-

surface junctions were observed by scanning electron microscopy on three different surfaces. These findings set the stage for working toward a high surface density of such features which might act as a platform from which to build a synthetic ligament.

3.2.2 Introduction

It is recognized that biomimetic motifs centered on self-assembly, typically driven by weak interactions like hydrogen bonding and metal-coordination chemistry, produce useful nanoscale architectures.113 In tissue engineering, it is often important to control the presentation of proteins and molecular assemblies at the nanoscale to elicit desired cellular responses.114 Often, simply transferring a portion of a biological extracellular matrix to a treatment site is not sufficient. The natural matrix may possess poor mechanical properties and there may be problems with an immune response. Thus, it is common to design and synthesize artificial nanoscale patterned surfaces intended to produce clinically useful interfaces.115 This paper presents a new chemical approach designed to produce such a nanopatterned surface.

Most extracellular matrices are built upon a collagen scaffold. The two major collagen types used as substrates on which to build these scaffolds are type I and type

IV.116 In this paper, we focus on attaching type I collagen to a surface. Since many clinical applications use titanium implants interfacing with predominantly type I collagen tissues like bone, tendon, and ligament, we look at attaching type I collagen to a titanium surface.84 The chief novel aspect of this attachment is that we have used chemistry intended to predominantly form attachments only at the amino terminus of the collagen molecule. This has not been done previously.

Typically, “collagen-coated surface” refers to a plastic, glass, or metal oxide substrate exposed to a collagen-monomer solution such that the collagen non-covalently 68

adsorbs onto the surface.117–120 Most collagen molecules on such surfaces lie parallel to

the plane of the surface. To bind a collagen molecule end-on to a surface, the N- or C-

terminal ends must be targeted. EDC or EDC/NHS reactions are used to form peptide

bonds between carboxyl and primary-amine groups.54 However, amines and carboxyls

are not unique to the termini of proteins. There are only 3 terminal amines and 3

terminal carboxyls per collagen molecule, but around 240 residues containing

carboxylate side-groups (aspartic and glutamic acids) and 100 containing primary amine

side-groups (lysines).55,56 Since these side-groups are much more prevalent than the N-

or C- termini, carbodiimide-based procedures are insufficient to site-specifically

conjugate collagen end-on to a surface.

To overcome the non-specificity of carbodiimide crosslinking reactions, we

chose an alkoxyamine-based method. Oxyamine groups react specifically with aldehyde

or ketone groups to form oxime linkages 59. We converted the terminal amines on

collagen molecules into ketones using a procedure introduced by Francis, then reacted

these ketone-collagen molecules with a surface decorated with oxyamine-terminated

silanes. The Francis procedure involves using pyridoxal-5-phosphate (or a PLP analog)

to transaminate N-terminal amines into ketones. The pKa of N-terminal α-amines (pKa =

6–8) is lower than that of other aliphatic amines (pKa 10.5), allowing us to target the

N-terminal amines by carrying out this reaction at pH ∼6.5.61 Oxyamine-silane molecules can then be conjugated to a surface. Using this method, we site-specifically immobilize individual collagen molecules.

A nucleation-and-growth fibrillogenesis procedure47 is known to allow collagen

to self-assemble into fibrils via weak interactions under physiological conditions101. As

one application of our site-specific immobilization, we used the site-specifically immobilized monomers as nucleation sites around which to grow collagen fibrils. The idea was to coat a titanium surface with a monolayer of covalently-bound upright- standing collagen molecules, form fibril nuclei around these monomers,45 and finally grow the nuclei into upright-standing fibrils. This paper highlights the work done on producing monolayers of collagen molecules.

3.2.3 Results and Discussion

Our results are presented in an order that parallels the sequence of steps used to produce upright-standing fibrils on a titanium surface. Figure 3.1 summarizes these four steps. In step 1, a plasma-treated titanium surface is functionalized with a monolayer of oxyamine-terminated alkylsilanes. In step 2, the terminal amines of a solution of

Combine Activated Single end-on Fibrillogenesis on Titanium 4 Surface and fibril Collagen Monomers

Nucleation Single Collagen 3 Monomers on Ti

Growth

Figure 3.1: Overview of the chemical steps necessary to produce collagen fibrils conjugated end-on to a titanium surface. The chemical process can be broken into four distinct steps: 1) activation and silanization of the titanium surfaces, 2) activation of collagen monomers by n-terminal transamination, 3) conjugation of ketone-collagen monomers end-on to the silanized titanium, and 4) nucleation on the single ketone-collagen monomers followed by growth to produce fibrils.

collagen molecules are converted to ketones using Rapoport’s salt. In step 3, the

resultant ketone-collagen is exposed to the oxyamine-functionalized titanium surface

resulting in collagen molecules being firmly attached to the titanium. In step 4, collagen

fibrils are nucleated around the covalently bonded collagen molecules. These nuclei are

subsequently grown into single nanofibrils. In what follows, evidence is presented that

each of these steps results in the desired product.

Step 1: Oxyamine-Silanization of Titanium

To determine the efficacy of the oxyamine-silane-surface conjugation chemistry

performed in step 1, we used X-ray photoelectron spectroscopy (XPS) to study the

atomic composition of titanium before and after oxyamine-silane functionalization

(surface activation). We focused on nitrogen, carbon, and silicon because these elements

exist in the oxyamine-silane molecule but are not present in appreciable amounts on

clean titanium surfaces, which are mainly comprised of titanium and oxygen.

Figure 3.2 shows the typical XPS spectra of both clean and oxyamine-

functionalized titanium surfaces. The main peaks are oxygen (530 eV), titanium (460

eV), nitrogen (400 eV), carbon (280 eV), and silicon (100 eV). The N1s peak located at

400.13 eV on the activated surface is consistent with an oxyamine-containing molecule

according to work by Yoshida et. al. documented in the NIST X-ray Photoelectron

Spectroscopy Database.99 Qualitatively, N1s and Si2p both show noticeable peaks after the conjugation reaction while little to no intensity is seen in the control spectra. The insert at the top right of Figure 3.2 shows the size of the nitrogen peak is larger for the

oxyamine activated surface than it is for the clean titanium further supporting the hypothesis that oxyamine functional groups were successfully conjugated to the titanium surface. Areas under the peaks were used to determine the relative atomic percent of each element. Nitrogen accounts for

approximately 2% of the atoms Figure 3.2: XPS spectra of a clean titanium slide without surface modification (bottom blue spectrum) and a titanium slide modified on the oxyamine activated by surface addition of oxyamine-silanes (top red spectrum). The N1s peak near 400 eV binding energy and the Si2p peak near 100 eV binding energy (both highlighted in yellow) are surface. Assuming that the substantially larger on the oxyamine-silane functionalized slide compared to the unmodified clean slide. The inset shows a magnified view of the N1s peak along with the best fit curve used oxyamine molecules are evenly to determine peak position and area. distributed across the titanium and that the titanium surface has a tetragonal rutile crystal structure, 2% nitrogen equates to one oxyamine functional group every 4.6 nm on the surface of a (0,0,1) plane.100 Typical reconstituted collagen fibrils are generally larger than 10 nm in diameter. Thus, this spacing of collagen monomers is sufficient to allow for multiple covalently bound monomers to be associated with each fibril grown on the surface. Of course, if every oxyamine site nucleates an independent fibril, this could lead to problems in that only very thin fibrils could be grown prior to impinging on their nearest neighbor. It will be shown that we have the opposite problem of producing too few total fibrils. So future work will focus on increasing nucleated fibrils rather than decreasing this rate.

Step 2: Ketone-Collagen Production

In step 2, single collagen monomers are modified to present ketones rather than amines on one or more of the

N-termini of each of the three collagen alpha chains. These non-biologically

synthesized ketones are unique to the Figure 3.3: UV-vis spectra of collagen solutions. The TAMRA-labeled ketone-collagen solution (black) molecule, allowing molecular site-specific shows a peak at ~560 nm, indicative of TAMRA, and ~230 nm, indicative of collagen. The unmodified collagen (red) only shows a peak at ~230 nm for attachment to the activated titanium collagen. surface in step 3. We utilized a procedure that involves using a pyridoxal-5-phosphate,

PLP, analog to transaminate N-terminal amines into ketones. While PLP and other analogs have been used to modify the N-termini of many proteins such as streptavidin and IgG,59,61,69,70,121,122 no one has yet applied this reaction to collagen molecules. We found the traditional reaction conditions47 lead to aggregation of the collagen molecules.

This was likely because the reaction needs to be carried out at 37 °C near neutral pH and the PLP, which contains a phosphate group, is used at concentrations in excess of 5 mmol. These are well-known conditions that lead to collagen monomer aggregation via fibril formation. A divalent anion, typically phosphate ions, are necessary for collagen fibrillogenesis.53 For this reason, we modified the transamination procedure to prevent the collagen molecules from aggregating. We chose N-methylpyridinium benzosulfate as the PLP analog because it does not contain a phosphate group. In addition, it has been shown to work on proteins having a similar N-terminal sequence to one of the collagen

alpha chains.75 Using MES to buffer the solution to pH 6.5, we found no obvious

problems with the reaction until, at the end of the reaction, we attempted to exchange

the solution for one without the Rapoport’s salt. Upon this exchange, we saw visible

aggregates of collagen forming. We were able to prevent this aggregation by adding 150

mM MgCl2 to the exchange buffer. This may have worked due to an increased

electrostatic repulsion between collagen molecules.105

To determine the efficiency of the ketone conversion, a UV-active dye,

aminooxy-5(6)-tetramethylrhodamine (aminooxy-5(6)-TAMRA) was used. TAMRA

has an aminooxy group like the oxyamine-silane from step 1, allowing it to attach to the

ketone groups on the collagen monomers in an analogous fashion. Ultraviolet-visible

spectroscopy (UV-vis) was used to ascertain the yield of the transamination chemistry

performed on the collagen monomers in step 2. UV-absorbances of the TAMRA dye

and the collagen molecules were both measured alongside known concentrations of each

molecule allowing the measured sample absorbances to be converted to concentrations.

Collagen has an absorbance at around 230 nm, while TAMRA has an absorbance at both

230 and 560 nm. The small contribution TAMRA gives to the 230 nm absorbance was

accounted for when calculating the collagen content of the sample solutions. Figure 3.3 shows this data. The concentrations of TAMRA and collagen in the reacted solution gave a ratio of TAMRA to collagen of 1 to 10. Some ambiguity remains regarding what percentage of the collagen monomers contain ketone end groups.

Each collagen molecule consists of three alpha chains, each of which might have been converted to a ketone. Thus, each collagen molecule has the potential to bind up to three TAMRA molecules. This gives a range of converted collagen monomer efficiency

somewhere between 3.3% (assuming all three alpha chains were converted) and 10%

(assuming only one alpha chain per collagen molecule was converted). Based on previous work, this range of efficiencies can be reduced. Lightly pepsin digested type I collagen displays three different triplets on the three alpha chains making up the molecule: ISV, DEK, and DAK. Witus et al. showed that groups with a terminal I, isoleucine, show almost no reactivity to transamination.59 This allows us to rule out the chain ending in ISV as having been converted to contain a ketone. They did not specifically test DEK or DAK. However, they did show that DES had a conversion efficiency near 50% while DIQ was near 0%. Thus, we cannot say whether one or both of the two chains with a terminal D, aspartic acid, were converted to ketones. Our conclusion, then, is that somewhere between 5% and 10% of the collagen molecules in our initial solution were converted to ketone-collagens.

Step 3: Binding Ketone-Collagen to Silanized Titanium

To detect collagen molecules on the titanium surface after the ketone collagen solution is presented to the oxyamine functionalized titanium surface, fluorescent antibody labeling is used. Figure 3.4 shows characteristic images of the antibody-stained surfaces displayed to the right of the histograms. Care was taken to ensure the illumination and display settings were the same for each sample. The scale bar in each image is 5-microns. To the left, data from these images is quantified with corresponding histograms of their grayscale values. Antibody labeling was performed on oxyamine functionalized titanium surfaces exposed to one of three conditions: (top) only oxyamine chemistry; (middle) oxyamine chemistry followed by exposure to unmodified collagen

molecules; (bottom) oxyamine chemistry followed by exposure to modified (ketone-) collagen molecules. Quantitatively, the three treatment groups showed a mean brightness of 14, 28, and 9.8X103, respectively. The mean grayscale value for the treatment group is nearly 1000X greater than that of either control group indicating that steps 1-3 successfully functionalized titanium surfaces with ketone- collagen molecules.

Since the ketone exists only at the terminus of the molecule and the binding sites take up

only ~2% of the surface Figure 3.4: Histograms of immunofluorescence grayscale values for the detection of collagen molecules on activated titanium surfaces. (top) Titanium slide that only underwent oxyamine chemistry and was not area, a simple steric exposed to any collagen. The image appeared black to the eye and quantitatively showed an intensity histogram with a peak at a grayscale argument suggests that, value of 14. (middle) Titanium slide that underwent oxyamine chemistry and was subsequently exposed to unmodified collagen molecules. Similar to the surface exposed to no collagen, the image appears black to the eye on average, the collagen and displays a histogram with a very low mean grayscale value of 28. (bottom) Titanium slide that underwent oxyamine chemistry and was subsequently exposed to modified collagen molecules. The field of view molecular axis of looked noticeably brighter with a generally uniform contrast. Scale bars are 5 microns long. Quantitatively, the histogram of image intensity gray symmetry projects away scale values had a mean of 9.8*103, nearly 1000X greater than the control surfaces. from the surface at some angle close to the surface normal.

commensurate with the optical images. Since very little antibody binding is seen on the control slides, it is unlikely that collagen attachment not specific to the oxyamine-

(ketone-collagen) reaction occurred. Assuming that the bound collagen molecules extend up from the surface at some appreciable angle, they should be available to act as nucleation sites for fibril formation. Despite the low percent nitrogen observed on the oxyamine-silane surfaces from step 1, the homogeneity and brightness of the antibody- stained collagen-molecule-coated slides suggest that significant decoration of the surface with collagen monomers has occurred. We did not directly assay for covalent bonding between the collagen molecules and the oxyamines. However, the chemical reaction between the terminal ketone and the oxyamine is well known to result in a covalent bond. In addition, copious washing was performed several times after the conjugation chemistry so we are confident that attachment of the molecules was robust.

Step 4: Nucleation and Growth of Collagen Fibrils on Titanium

Scanning electron microscopy (SEM) was used to determine if collagen fibrils were grown on the modified titanium surfaces. We did not attempt to assay for nucleated microfibrils because these would have been near the resolution limit of our microscope. Figure 3.5 shows one clean surface (A) and two chemically modified surfaces (B & C). The surface shown in Figure 3.5A was a bare titanium surface that had not been exposed to any solvents. The surfaces shown in Figures Figure 3.5B and

Figure 3.5C were exposed to steps 1-3 as well as fibril nucleation and growth steps.

Briefly, the surface with bound collagen molecules was exposed to a high-concentration

(200 µg/ml) collagen monomer solution for one minute. This is hypothesized to have allowed nucleation to occur both on the surface and in the bulk solvent. Further fibrillogenesis was stopped by replacing the solution with one containing no free collagen. Thus, any solution- phase nuclei were removed while the surface- bound nuclei remained. Finally, the surfaces were exposed to a low collagen concentration solution (3 µg/ml) for 17 hours. This concentration is hypothesized to have been too low to allow for bulk nucleation but high Figure 3.5: SEM images of titanium surfaces. enough to allow for the growth of the surface- A: clean slide showing no fibril-like formations. B: net of collagen fibrils standing off the surface. C: single fibril standing bound nuclei. Figure 3.5C shows a collagen upright. fibril standing about 5 microns tall with an average diameter of 73 nanometers. Proof that this fibril was standing up from the surface was obtained when it was seen to fall over during imaging, likely due to the bombardment of the electron beam (video in supplemental data). Five such fibrils were imaged on three different modified titanium surfaces. While such free-standing fibrils were rare in these experiments, net-like fibril structures like the one shown in Figure 3.5B were found on every slide having undergone steps 1-4. They were seen about 5X more often than the upright fibrils. At the base of these nets, individual fibrils can be seen attaching perpendicularly to the

titanium surfaces.

Figure 3.6 shows images of these end-on attachments with several of them highlighted by arrows.

These image features are consistent with the procedure outlined in

Figure 3.1 producing end-on conjugated

collagen fibrils on a Figure 3.6: SEM images of zoomed-in titanium surfaces. The picture in the top right is a clean, unmodified surface. The rest of the image show fibrils making up fibril nets similar to the net in Fig. 5B. Arrows surface. show places where fibrils interface perpendicularly to the surface, indicating end-on attachment. all scale bars are 200 nm long. The SEM images are consistent with some collagen molecules self-assembling to produce fibrils oriented with their symmetry axis parallel to the titanium surface normal. Only surfaces that have gone through plasma treatment, oxyamine functionalization, ketone-collagen conjugation, and exposure to nucleation and growth collagen solutions show such fibrils. All other surfaces look like surface A. Since only our chemistry produces these fibrils, we assume that they are conjugated end-on to the surface, given the assumptions garnered from the results of steps 1-3.

3.2.4 Conclusions

A proof-of-concept was presented showing it is possible to produce titanium 79

surfaces with collagen fibrils bound roughly perpendicular to the surface. Data was

presented showing that each of the four steps needed to obtain this result can be

accomplished. First, XPS showed plasma treatment with argon gas bubbled through

water followed by exposure to an oxyamine terminated silane results in a surface

containing about 2% nitrogen. Chemical shift data from the XPS spectra is consistent

with these nitrogens being present as oxyamines. Second, UV-vis spectroscopy shows

that a procedure using Rapoport's salt in a non-phosphate containing buffer is capable of

modifying single collagen molecules to express ketones. Consistent with previously

published work, these ketones are likely to present only at the amino termini of the

collagen monomers. Third, fluorescently tagged antibody labeling shows that exposure

of the oxyamine functionalized titanium surface to the modified collagen monomers

results in attachment of the collagen monomers to the titanium. The uniform brightness

of these fluorescently labeled surfaces is consistent with the collagen monomers being

evenly distributed on the titanium surface at the micron scale. Fourth, SEM images

show that a two-step procedure involving exposing the collagen monomers attached to

the surface to high concentrations of soluble collagen for short periods followed by a

lower concentration for 24 hours resulted in 50-100 nm diameter collagen fibrils attached to the titanium surfaces. Some of these fibrils are seen attaching nearly perpendicularly to the surfaces.

3.2.5 Methods

Figure 3.1 provides an overview of the procedures used to create collagen fibrils on titanium surfaces.

Production of titanium-coated silicon surfaces: Six-inch silicon wafers were

titanium-sputtered using an MRC sputtering tool in argon at 200 WRF, 8 mTorr, and at a rate of 66.6 Å/min. Wafers were coated with 200 nm of titanium and then diced into 1 cm squares using the Disco DAD3350 Dicing Saw.

Activation of titanium surface to produce surface hydroxyls: 1 cm square

titanium wafers are cleaned by sonication with ethanol followed by distilled water.

Wafers are then exposed for 15 minutes to a plasma of argon bubbled through DI water

at 50 watts of RF power (Branson/IPC model PM 118 500 W, Hayward, CA).

Activation of hydroxylated titanium surface to produce surface oxyamines:

The hydroxylated titanium surfaces are placed in rapid-prototyped polycarbonate

holders able to hold 2 ml of fluid and allowing withdrawal of all but 100 µl of a solution

in a manner that keeps the titanium surface wet at all times. The surfaces are then

immersed in a 5% solution of 11-(O-hydroxylamine)undecyltriethoxysilane in 200 proof

ethanol for 10 minutes at room temperature. The surfaces are then rinsed profusely with

ethanol then DI water within the holders.

Activation of collagen to produce ketone-collagen: A 50 mM solution of type

I bovine dermis atelocollagen (PureCol, Advanced BioMatrix, San Diego, CA) is

incubated at pH 6.5 and 37°C for one hour with 100 mM Rappaport's Salt (RS, N-

Methylpyridinium-4-carboxaldehyde benzenesulfonate) in a buffer containing 30 mM 2-

(N-morpholino)ethanesulfonic acid (MES), 100 mM sodium chloride, and 150 mM

magnesium chloride. The ketone-collagen solution is then concentrated and buffer-

exchanged using 0.5 mL,100 kDa MWCO centrifugal filters (Millipore Amicon Ultra,

Cork, Ireland) spun at 12g for 15 minutes. The sample solution is concentrated to 200

µL and then rinsed six times with 150 mM MgCl2. The final solution is stored at 4 mg/mL collagen at -80°C.

Reacting ketone-collagen with the activated titanium surfaces: The oxyamine functionalized titanium surfaces are reacted with 40 µg/mL of ketone-collagen in 100

mM MES buffer at pH 5 for 17 hours at room temperature. Following the reaction, the

slides are rinsed with MES buffer and subsequently PBS.

Growing fibrils: Using the conjugated ketone-collagen molecules as nucleation

sites, a nucleation solution containing 200 µg/mL of free, unmodified collagen

monomers in 30°C PBS is used to create fibril nuclei. The nucleation solution is left on

the surface for 1 minute and subsequently rinsed thoroughly with PBS. This process is

repeated twice. The surfaces are then exposed to a growth solution of 3 µg/mL collagen

in PBS for 17 hours at 37° C then rinsed with PBS.

X-ray Photoelectron Spectroscopy: A Phi Versaprobe 5000 spectrometer is

used to collect survey scans. The area beneath the nitrogen 1s and silicon 2p peaks are

reported as a fraction of the area under all peaks. Each peak was corrected for x-ray

sensitivity using the built-in Phi software.

UV-Visible Spectroscopy: The ketone-collagen molecules are reacted with aminooxy-(5,6)-TAMRA at a molar ratio of 1:10 collagen to TAMRA. The reaction proceeds for 24 hours at room temperature. The solution is then purified via centrifugal spin filtration using 30-minute intervals at 14,000 RCF until no purple absorbance is observed in the flow-through. The final solution's absorbance spectra analyzed using

UV-Visible spectroscopy (NanoDrop 1000 Spectrophotometer, Thermo Fisher

Scientific, Wilmington, DE). More information on the absorbance analysis can be found

in the supplemental information.

Fluorescence Microscopy: Surfaces are exposed to a rabbit anti-bovine collagen type I antibody (BIO-RAD Laboratories, Hercules, California). Secondary staining is done with anti-rabbit IgG-FTIC (Sigma-Aldrich Corporation, Saint Louis,

MO). Samples are imaged with an upright light microscope (Nikon Eclipse E600FN) equipped with a digital camera (Blackfly S USB3, FLIR, Arlington, VA) and Spinnaker

SDK imaging software. Four images per sample are obtained. Histograms of the grayscale values of each sample are created using MATLAB and ImageJ, where the images are run through a Matlab program to calculate the average brightness and standard deviation of each sample. For viewing purposes, characteristic images of each sample are set to the same window of grayscale values, 0-14,000, using ImageJ.

Scanning Electron Microscopy: Samples are ethanol-dehydrated and critical- point dried (Baltec CPD 030). They are then sputter-coated with 20 nm of palladium

(Denton DESK IV, Denton Vacuum, Moorestown, NJ) and imaged with an FEI Helios

Nanolab 650. Images shown are collected at a tilt angle of 40°.

Chapter 4: Conclusions and Future Work

4.1 Conclusions

The clinical pathologies experienced around dental implants may be derived from the a-biomimetic bone-implant interface. Implementing a biomimetic artificial periodontal ligament at this interface may address the peri-implant complications which remove from the quality of life that a healthy dental implant is meant to ensure. In this thesis, a proof-of-concept was presented showing it is possible to produce titanium surfaces with collagen fibrils bound roughly perpendicular to the surfaces. These surfaces lay the groundwork to create synthetic brushes of Sharpey’s fibers. Data was presented showing that each of the four steps needed to obtain this result can be accomplished. First, XPS showed plasma treatment with argon gas bubbled through water followed by exposure to an oxyamine terminated silane results in a surface containing about 2% nitrogen. Chemical shift data from the XPS spectra is consistent with these nitrogens being present as oxyamines. Second, UV-vis spectroscopy shows that a procedure using Rapoport's salt in a non-phosphate containing buffer is capable of modifying single collagen molecules to express ketones. Consistent with previously published work, these ketones are likely to present only at the amino termini of the collagen monomers. Third, fluorescently tagged antibody labeling shows that exposure of the oxyamine functionalized titanium surface to the modified collagen monomers results in attachment of the collagen monomers to the titanium. The uniform brightness of these fluorescently labeled surfaces is consistent with the collagen monomers being evenly distributed on the titanium surface at the micron scale. Fourth, SEM images show that a two-step procedure involving exposing the collagen monomers attached to

the surface to high concentrations of soluble collagen for short periods followed by a

lower concentration for 24 hours resulted in 50-100 nm diameter collagen fibrils

attached to the titanium surfaces. Some of these fibrils are seen attaching nearly

perpendicularly to the surfaces.

4.2 Future Work

While we have shown proof-of-concept that a few collagen fibrils can be grown perpendicular to a surface, there is still the need to increase the surface density of these features substantially before any significant mechanical load could be transferred through this surface. There are two approaches to increasing this feature density. First, it is necessary to determine if the net-like structures seen in Figure 3.5B and Figure 3.6 are actually upright standing fibrils in solution or if these nets represent a surface area that is not available for ligament-like fibril occupation. Other sample preparation techniques such as collagen crosslinking, uranyl acetate stabilization, and freeze-drying may answer

this question. Second, the density of upright standing fibrils needs to be increased.

Several strategies might impact this. Adding an amine catalyst such as ethylenediamine

(EDA) to the silanization mixture could increase the density of amine functional groups

on the titanium surfaces.80 Additionally, gaining better control over the nucleation and

growth conditions would also allow for a rational approach to modifying the conditions

in step 4 to optimize for fibril density. Further turbidity measurements could be used to

find solution conditions and reaction times that will optimize the kinetics.

It may even be worthwhile to investigate a new collagen-surface conjugation

strategy altogether to achieve a desirable fibril density for mimicking Sharpey’s fibers.

In recent publications, the Francis group has found success in conjugating extracellular

matrix proteins to surfaces using a one-step site-specific protein modification

chemistry.123 In their original 2006 screening for aldehyde-reagents capable of

transaminating proteins, they also found that some of these substrates formed 1:1

condensation products with the peptides.62 2-Pyridinecarboxaldehyde (2-PCA) seemed to be particularly reactive to the n-terminal aspartic acid of angiotensin 1, forming a condensation product with no evidence for transamination, for conjugation to ɛ-amino groups of lysine residues, or for crosslinking between peptides. In the newer 2015 paper, the Francis group then modified the 2-PCA to express a secondary amine for functionalization and a tertiary amine for increased water solubility.123 The amine can be

used with NHS chemistry to attach the 2-PCA to a hydroxylated substrate, such as

hydroxylated titanium. For their extracellular matrix protein experiments, the group

formed 2-PCA functionalized polyacrylamide hydrogels and incubated these substrates

directly with 400 μL of 50 μg/mL collagen.124 The proteins used were type I bovine

collagen (PureCol), human plasma fibronectin, and mouse laminin. This process does

not require N-terminal modification of the proteins as the PLP protocol does, thus

removing the need for anti-gelation reagents like magnesium chloride. This surface

modification and subsequent collagen reaction may be a good alternative to the protocol

outlined in this thesis if consistent fibrillar surfaces cannot be obtained.

Further, more work, potentially using FTIR or other analysis techniques, needs

to be done to confirm that an oxime bond does exist between the ketone-collagen (and

therefore the fibrils) and the silane molecules. It would also be reasonable to do longer-

term in-vitro studies where collagen-functionalized slides are exposed to pH values in

the range of 5.6-7.0 to determine the long-term impact of the acidic conditions found

around dental implants.

Loading Fibril Surfaces

Once a consistent, dense surface of individual standing fibrils can be produced, it

will be necessary to do various in-vitro tests to assess the strength of the bound fibrils

before starting animal studies. To act as a brush of Sharpey’s fibers, the collagen-fibril

coating on the dental implant will need to withstand at least 120N of axial force as teeth

during normal chewing.23 Calculating the tensile rupture force of the bonds in the

synthetic ligament system will help determine the density of fibrils needed to withstand

the forces of mastication.

Most of the

force experienced by

periodontal ligament is

tensile. There are

several bonds in this

system as seen in

Figure 4.1, but

Grandbois et. al.

suggests that the C-Si

bond within the silane Figure 4.1: Chemical structure of surface-bound ketone collagen molecules. molecules would be A silanized titanium surface (top left) is exposed to ketone-modified collagen molecules (top right) resulting in an oxime bond between the collagen and the silane molecule. The final structure (bottom) shows all of the covalent bonds in the weakest. The ratio the system used to determine the tensile strength of bound fibrils.

of bond energy to bond length can be indicative of the theoretical strength of a bond.125

All considered bonds in the system are laid out in Figure 4.1.126–128

Table 4: Bond energies, lengths, and energy-to-length ratios of covalent bonds in the synthetic ligament system

Bond Bond Energy, D Bond Length, r D/r

(kJ/mol) (pm)

Si-O 452 163 2.77

C-Si 381 185 1.89

C-C 384 154 2.49

C-O 358 143 2.50

O-N 201 140 1.55

N=C 651 129 5.05

The Si-O bond has a D/r ratio of 2.77 and is not a candidate for the weakest link

because the triethoxy silane used forms three parallel Si-O bonds with the surface. The

C-Si bond has a bond energy (D) of 381 kJ/mol and a bond length (r) of 202 pm, giving

a D/r ratio of 1.89. While the bond strength of the O-N bond (D/r = 1.55) should

theoretically be the weakest bond, I was only able to find rupture force data for the

second weakest bond, C-Si. As such, the Si-C bond will be used to calculate the

maximum tensile force the ligament could withstand.

The Grandbois group used atomic force microscopy (AFM) to measure the

rupture force of various bonds including Si-C, Si-O, and C-N. Si-C was found to be the

weakest with an experimental rupture force of 2.0 ± 0.3 nN using force-loading rates of

10 nN per second. To compare, the C-N bond had a rupture force of about 4 nN.125

Assuming the Si-C bond is the weak link, each collagen fibril is loaded equally in

tension, and each collagen fibril is attached to only one silane molecule, an implant ligament would require 6×1010 fibrils to withstand 120N of tensile force. The popular 88

Brånemark System implant has a potential bone/implant contact area of 138 mm2.129

Such a system would require a fibril density of 435 fibrils per nanometer. Given that a

collagen fibril has an average diameter of 75 nm, this density is impossible to achieve.

To compare, in designing this system we assumed that a density of 1 fibril per 300 nm

would be sufficient to prevent the fibrils from falling over, and as such is our benchmark

for “dense” ligament. Using this density, an implant with a surface area of 138 mm2

would have only 4.6×10 fibrils and would be able to withstand just 920 µN of force. 5 Such force would be greatly exceeded during normal chewing behaviors and as such the

ligament would fail.

Beyond increasing the fibril density, the strength of the ligament could be

improved by bonding each fibril to more than one silane molecule; each fibril would

need to contain more than one ketone-collagen molecule from the titanium surface. If each collagen fibril is bound to the surface in more than one spot, the strength of each fibril is expected to increase. Additionally, a ligament is not made entirely of collagen fibrils; there are various other proteoglycans, matrix molecules, and blood vessels that

contribute to the strength of a natural ligament. Tissue engineering using the developed

technology as a scaffold may be necessary to make a functional artificial periodontal

ligament. In addition to improving mechanical properties, such a collagenous surface decorating the implant and abutment could provide a platform from which to deliver bioactive factors to treat infection, inflammation, and promote soft tissue adhesion.

Animal Modeling

Once a desirable density is achieved, simple animal models could then be used to assay for a biological response. Rat models can be used to assay the

biocompatibility/safety of the functionalized implant.130 For example, tests for

genotoxicity, carcinogenicity, and reproductive toxicity, for interactions with blood, and

in-vitro cytotoxicity could all be done using cells and fluid derived from rats. Following

in-vitro cell studies, functionalized implants could be placed in in-vivo rat models. Rat

models typically have an implant healing period of 8 weeks.49,131 A more complex

animal model may involve pigs or dogs. Dogs are often used for the assessment of

dental implants, partially due to their susceptibility to biofilm accumulation and spontaneous periodontitis formation, but also because larger dogs have jaws that can

accommodate human-sized dental implants.130

If through the animal studies, we find that the artificial Sharpey’s fibers on the

implant do not behave as desired, we could conjugate growth factors and/or non-

collagenous proteins to further imitate the natural ligament. For example, we wish to

mimic the biological mineralized dentin-unmineralized ligament-mineralized bone interface found in a natural ligament, meaning that the free ends of the fibers are expected to mineralize into the jaw bone but a portion of the fibers near the titanium is expected to remain unmineralized. The bone-fiber interface may not form correctly on healing. If the bone does not grow around the free ends of the artificial Sharpey’s fibers as they do in a natural ligament, BMP-2 (Bone Morphogenetic Protein 2) could be added to the bone-ends of the fibrous matrix to encourage bone infiltration and mineralization.49. At the same time, a portion of the Sharpey’s fibers must remain

unmineralized to carry out the structural functions of the natural periodontal ligament.

As discussed in section 1.2, fibroblasts play an important role in preventing bone from

infiltrating into the non-mineralized portion of the periodontal ligament.32 Human

periodontal ligament fibroblasts have been shown to inhibit bone nodule formation in- vitro.132 One chemical responsible for this action is Fibroblast Growth Factor 2 (FGF2), which inhibits alkaline phosphatase (ALP) activity, collagen synthesis, and matrix mineralization and while increasing cell proliferation. The addition of FGF2 to the inner portions of the artificial Sharpey’s fibers may help to maintain an unmineralized, ligament-like section of tissue between the implant and the bone. Further, the strength and durability of the ligament must also be able to withstand the normal occlusal forces experienced while chewing. If it seems that the artificial Sharpey’s fibers alone are not sufficient, more of the non-collagenous extracellular matrix components (such as elastin and fibronectin) found in natural ligament could be introduced to the scaffold. It may also be desirable to add recombinant human amelogenin (rHAM) to the matrix, as rHAM encourages the healing of tooth-supporting tissues around natural teeth.133

4.3 Other Potential Applications

Space-safe collagen adhesive

This idea was generated by Dr. Eppell before I entered the lab. In space, such as on the

ISS or in travel to Mars, NASA has proposed that there is a need to stably connect titanium pieces Figure 4.2: Diagram of bio-inspired adhesive using two titanium pieces without the use of heat, organic solvents, or other functionalized with collagen fibrils. hazardous chemicals. They want to build titanium structures and machines in space as safely and cost-effectively as possible. One solution to this need is an application of the technology described in this paper; two titanium pieces functionalized with standing collagen fibrils can be glued together with the addition of a simple sugar solution. 91

Collagen fibrils naturally crosslink in the presence of glucose, forming strong covalent bonds. Figure 4.2 shows a diagram of this design. This new adhesive would be repairable, more resilient and safer for astronauts than cyanoacrylates, and would allow for the adhesion of rough surfaces without a need for hazardous chemicals. Solution- phase collagen molecules added to the bonding sugar solution could be used to repair any broken fibrils if the interface is damaged. The material could also be released by using an enzyme near-neutral pH, eliminating the need for permanent complex adhesives and allowing robotic parts to be re-aligned if needed.

Dynamic Testing of Fibrils Synthesized In-vitro

Our lab is also concerned with the dynamic testing of individual collagen fibrils.

There is a wide range of hypotheses about the mechanisms of tendon injury at the nanoscale. These hypotheses are being explored by Christopher Slater by visualizing damage at the fibril and collagen molecule scale. Understanding the mechanism of tendon damage at this scale could improve treatment strategies or lead to better imaging of damage before it becomes irreversible. To that end, our lab has developed a microelectrical mechanical system (MEMS) device that has been used to strain and measure the strength and stiffness of single fibrils extracted from rat-tail tendons. To visualize damage, the fibrils are stained before and after straining with a collagen hybridizing peptide that only binds to denatured collagen monomers. These fibrils are made by the animal and are highly cross-linked in-vivo. If in-vitro-synthesized fibrils could also be tested with the MEMS device, the properties of the synthesized fibrils could be compared to that of the natural fibrils, giving an estimate of the functionality of synthetic fibril structures when used in-vivo. Unfortunately, fibril gels made in-vitro are typically so woven and chaotic that it is impossible to separate intact fibrils from the 92

mass. To get around that issue, the protocols described in this thesis could be used to build synthetic fibrils directly on the MEMS devices. Using a thiol functionality instead of a silane, collagen fibrils could be built on gold nanoparticles placed in controlled locations on the silicon MEMS devices.129 These fibrils could then be stretched across the MEMS devices and tested using the current MEMS protocol.

Protocols

This appendix details all of the tools and processes necessary to produce

collagen fibrils upright on a titanium surface. These are the steps that Dr. Steven Eppell,

Eloise Miller, and Aidan DiSanto have developed in Dr. Eppell’s laboratory.

First, this appendix contains a detailed protocol of how to obtain the titanium and

cleaning procedures for the necessary equipment.

Next, this appendix includes protocols for every step necessary to produce

collagen fibrils on a titanium surface in chronological order. Each protocol starts with

the Necessary Components, a section that outlines all of the tools, equipment, and

chemicals needed to follow the specified protocol.

Finally, this appendix outlines procedures that allow one to determine the

efficacy of each chemical step in the protocol, including how to prepare fibril-

functionalized samples for SEM imaging.

Titanium Slides

The square titanium slides are in a plastic container in drawer 7. They are stuck to a sheet of parafilm within the plastic container. We obtained these squares from

NASA. They are silicon wafers coated with 2000 angstroms of titanium. It is easy to chip the silicon and titanium, so be very careful when removing individual squares from the parafilm. They are located in drawer 7, marked “Aidan/Ellie”, below the pH meter

Necessary Components

• Gloves

• Teflon tweezers

• Clean Petri dish

• Patience

Protocol for obtaining titanium squares:

1. Put on gloves

2. Open the plastic container

3. Lift the parafilm with the gloved hand

4. Gently, put the thumb of the same hand on top of one of the squares, so you’re

gripping it from both sides of the parafilm

5. Gently stretch the parafilm out from underneath the titanium square, being

careful not to tilt it so that it chips on its neighbors

6. When the parafilm stretches, it should release the titanium

7. It may take many minutes to carefully remove the squares, so be patient

8. Place the titanium squares in a petri dish

To remove titanium from holders

1. Put the sharp end of a pair of metal tweezers in the divot beside the titanium

2. Gently pry the titanium upward

3. Grab the titanium with Teflon tweezers and transfer

Cleaning Procedures

The titanium surfaces and plastic holders must be thoroughly cleaned before chemistry begins. The holders and titanium squares are cleaned with the sonicator and thoroughly dried. When working with the titanium slides, do your best to only touch the edges or corners with the Teflon tweezers. This ensures that the middle of the slide is as 95

clean and un-scratched as possible. Place no more than three 20 mL beakers in the large

500 mL beakers when sonicating or the 20 mL beakers will become stuck and may break.

Set up the Sonicator

1. Fill the sonicator about halfway up with tap water

2. Turn on the sonicator

3. Place the acrylic sheet with two holes on top of the sonicator

4. Set the power to 99%

5. Set the time to 1 minute

6. Set the temperature to --, the lowest setting

Sonicating Titanium

7. Put on clean gloves

8. Fill solvent squeeze bottles

a. Methanol

b. Ethanol

c. Distilled water

9. Using Teflon tweezers, place one 1 cm titanium slide into a 20 mL beaker shiny-

side-up

10. Fill said 20 mL beaker with ~10 mL of methanol and set aside

11. Repeat with remaining titanium slides

12. Set aside

13. Fill a 500 mL beaker with 20 mL of DI water

14. Place the 500 mL beaker into one of the holes in the acrylic sheet, submerging

the bottom of the beaker in the sonicator water bath

15. Using long metal tweezers, place up to three 20 mL beakers into the 500 mL

beaker

16. Press the “start” button on the sonicator, wait for 1 minute

17. After sonication, remove each 20 mL beaker individually, pour out used

methanol into a waste container, and refill with ~10 mL of methanol. Set aside

18. Repeat with remaining 20 mL beakers

19. Repeat methanol sonication 3 times for all slides

20. Repeat sonication with ethanol followed by DI water, filling the beakers with

each solvent 3 times

21. After the final DI water sonication, remove 20 mL beakers from sonicator and

set aside

22. Get a clean petri dish and place one kimwipe, folded in half, in the bottom dish

23. Using Teflon tweezers, lift a titanium slide out of the 20 mL beaker

24. Spray the titanium slide liberally with DI water

25. Place the titanium slide shiny-side-up onto the kimwipe in the petri dish

26. Repeat with remaining slides

27. Take the petri dish with the slides over to the nitrogen tank by the fume hood, set

on the counter

28. The nitrogen tank is connected to a long tube with a metal 4-micron filter and

ends in a blue pipette tip – pick up the pipette tip like a pencil

29. Turn on the nitrogen tank, allow a gentle flow of nitrogen out of the pipette tip

(feel on your arm)

30. Blow all the water off the sides of the titanium with the clean nitrogen

a. Hold the titanium down with the Teflon tweezers if needed

b. The kimwipe sucks up the water

31. Transfer the dried titanium slides from the kimwipe onto the petri dish, cover,

and place in the 37C incubator to dry completely

Washing holders with soapy water

1. Put on clean gloves

2. Pick up a holder

3. Spray some soapy water into the holder

4. Scrub inside and outside of holder with a cleaning brush for 30 seconds

5. After washing, hold under the tap and rinse thoroughly for 1 minute, turning the

holder to remove all of the soap

a. Any soap left in the holder can hinder the reactions done later. Rinse the

holders very thoroughly!!

6. Place holder into a 20 mL beaker and fill with tap water, set aside for sonication

7. Repeat with the desired number of holders

Sonicating Holders

The protocol for sonicating the holders is similar to that of the titanium but with a few very important differences. The sonicator water bath must be below 35C when sonicating the holders in ethanol to prevent the breakdown of the polycarbonate. To be safe, add ice cubes to the water bath before/during ethanol sonication

1. Put on clean gloves

2. Fill solvent squeeze bottles

a. Tap water

b. Ethanol

c. Distilled water

3. Fill a 500 mL beaker with 20 mL of DI water

4. Place the 500 mL beaker into one of the holes in the acrylic sheet, submerging

the bottom of the beaker in the sonicator water bath

5. You should have the holders in 20 mL beakers filled with and tap water from the

previous cleaning step

6. Using long metal tweezers, place up to three 20 mL beakers into the 500 mL

beaker

7. Repeat with remaining holder beakers

8. Press the “start” button on the sonicator, wait for 1 minute

9. After sonication, remove each 20 mL beaker individually, pour out used tap

water into the sink, and refill with tap water. Set aside

10. Repeat with remaining 20 mL beakers

11. Repeat tap water sonication 3 times for all holders

12. Add ice from the freezer into the water bath to cool before ethanol sonication

13. Repeat sonication with ethanol, ensuring the water bath says cold by adding ice

as needed, 3 times for each holder

14. Repeat sonication with DI water, replacing DI water 3 times

15. After the final DI water sonication, remove 20 mL beakers from sonicator and

set aside

16. Get a clean petri dish and place one kimwipe, folded in half, in the bottom dish

17. Using tweezers, lift a holder out of the 20 mL beaker

18. Spray the holder liberally with DI water

19. Place the holder hole-side-down onto the kimwipe in the petri dish

20. Repeat with remaining holders

21. Take the petri dish with the holders over to the nitrogen tank by the fume hood,

set on the counter

22. Pick up a holder, still upside-down with your gloved fingers and bang it against

the kimwipe to knock out as much water as possible. Repeat with all holders

23. The nitrogen tank is connected to a long tube with a metal 4-micron filter and

ends in a blue pipette tip – pick up the pipette tip like a pencil

24. Turn on the nitrogen tank, allow a gentle flow of nitrogen out of the pipette tip

(feel on your arm)

25. Blow all the water out of the inside of the holders with the clean nitrogen

26. Transfer the dried holders from the kimwipe onto the petri dish, cover, and place

in the 37C incubator for at least an hour to dry completely.

Protocols – Chemical Steps to Produce Fibrils on a Titanium Surface

Ketone-Collagen

This step happens before any of the chemistry on the titanium.

The N-termini of the collagen molecules in this step are changed from an amine to a ketone. The ketone will react selectively with the oxyamine described in the next step. Many experiments worth of ketone-collagen is created in this step. The following procedure was developed by the Francis group at UC Berkley and was modified for use 100

with collagen in our lab.76

The RS may degrade in solution and therefore the RS solution should be made

fresh every time this reaction is done.

There is no need to adjust the pH of the RS solution. The RS reactant does not

significantly affect the pH of the buffer solution after its addition, so the pH of the

buffer (6.5) is consistent.

This recipe produces about 0.75 mg of ketone collagen at a concentration of

about 2 mg/mL.

Start this reaction as early as possible in the morning. Isolating the ketone-

collagen molecules by centrifugal filtration can take in excess of 4 hours. Overall, this procedure can take all day. This reaction and isolation must all happen in one day, as far as we know – I wouldn’t risk it.

Necessary Components:

• Hardware

o Gloves

o 15 mL falcon tube

o 2 mL microcentrifuge tube

o Surgical mask to prevent inhalation of RS particulate

o Weigh paper (for weighing RS)

o White Mettler Toledo analytical scale (for weighing RS)

o 37℃ incubator

o P-1000 micropipette and tips

o 100 kDa cutoff Amicon Ultra-0.5 mL Centrifugal Filters w/ two

101

included microcentrifuge tubes

o Blue Gusto/NASA centrifuge

o Waste beaker

• Chemicals

o pH 6.5 Transamination MES Buffer (see section “buffers and rinsing solutions”)

o N-methylpyridinium-4-carboxaldehyde benzenesulfonate salt (RS)

o 3 mg/mL PureCol Bovine Type I Collagen

o KeCol Rinsing Solution (see section “buffers and rinsing solutions”) Protocol:

1. Prepare a 100 Mm RS solution in pH 6.5 Transamination MES Buffer

(~0.5 hr.)

a. Ingredients

i. 4 mL pH 6.5 Transamination MES buffer

ii. 0.1118 g RS

• You will need 1.25 mL of RS solution for every 0.75 mg

of collagen (0.25 mL of 3 mg/mL PureCol Bovine Type I

Collagen) that is transaminated. You can make up to 3

vials of ketone collagen with this 4 mL volume.

b. Procedure

i. Pipette 4 mL of MES buffer into a 15 mL falcon tube

ii. Crease a square of weighing paper such that, after weighing, you

can easily dump your reactant into the solution.

102

1. Fold/crease a piece of weighing paper by the diagonal in

half to make a triangle. Open up the weigh paper back up

and fold/crease it across the other diagonal. Open up the

weigh paper. There should be a creased X on the paper

through the diagonals

2. Place the open and creased weigh paper cupped upward

onto the scale and zero it

iii. Put on a surgical mask to prevent inhalation of RS particulate

iv. Using the big white scale, measure out 0.1118 g of RS into the

creased weighing paper.

v. Dump the RS into the falcon tube, replace the cap, and shake until

dissolved

2. Make a 1.5 mL solution of PureCol collagen in the RS solution and perform the

transamination reaction (0.5 mg/mL of collagen)

(~1.5 hr.)

a. Ingredients

i. 1.25 mL of RS solution from part 1

ii. 0.25 mL of 3 mg/mL PureCol Bovine Type I Collagen (0.75 mg)

b. Procedure

i. Pipette 1.25 mL of RS solution into a 2 mL microcentrifuge tube

ii. Add 0.25 mL of PureCol collagen

iii. To mix, use a 1 mL pipette to gently draw up and push out the

103

solution into the tube

1. Vigorous mixing can cause the collagen to gel

iv. Place in the incubator at 37C for 1 hour

3. Removal of Excess RS - This step is particularly important because any RS that

remains mixed with the protein in the next step will quench your reaction by

reacting with your oxyamine compound, making it unable to react with the

ketone collagen. See the instruction manual for the 10kDa or 100kDa

microcentrifuge filters in the pink box.

(~4 hr.)

a. Ingredients

i. 1.5 mL of RS-collagen solution

• 1.5 mL of RS-collagen solution can fit into one centrifugal

filter. Many 1.5 mL volumes of RS-collagen solution can

be filtered at once by using many centrifugal filters.

ii. At least 20 mL of KeCol Rinsing Solution: 150 mM MgCl2

• You want to have enough buffer to thoroughly rinse all of

your samples. In a perfect world, you’d only need 3 mL

for each RS-collagen solution, but have more on hand.

b. Procedure

i. Add all your RS-collagen solution into the centrifugal filters by

concentration

1. Put a centrifuge filter into the provided centrifuge tubes

104

according to the box instructions

2. Place 500µL of collagen solution into the filter tube

3. Place in the centrifuge and balance

4. Run at 9,000 rpm (in the blue Gusto/NASA centrifuge) for

20-30 minutes until the solution is concentrated to a

volume of 100-200 µL

a. The filters say run at 14000 rcf, which is a measure

of the g-force the tubes will experience and can be

converted to RPM by knowing the diameter of the

centrifuge, but this centrifuge cannot handle

running at 25,000 RPM. This works just fine.

5. Dump the filtrate into a waste container. Don’t dump your

collagen!

6. Place another 300-400µL of collagen solution into the

filter

7. Repeat until all your solution is in the filters

ii. Fill the filter tubes with 300-400µL of KeCol Rinsing Solution

iii. Run for 30 minutes at 9,000 rpm to rinse out all of the RS

iv. Dump out the filtrate

v. Repeat rinses 6 times, more if the filtrate is still pink after the 6th

rinse.

4. Retrieval of ketone-collagen solution. When you have rinsed out all the RS and

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concentrated both filters down to 100-200 µL of ketone collagen solution, you

have to get all the solution out of the filters.

(~10 min)

a. Get out a clean centrifuge tube from the box

b. Remove the filter with the collagen solution from its used centrifuge tube

c. Carefully turn the filter over into the clean centrifuge tube so it is now

upside down

d. Place the tube into the centrifuge with the lid pointing away from the

center of the centrifuge

e. Balance the centrifuge with other tubes of the same weight

f. Spin at 4000 rpm for 3 minutes to collect the solution

g. Label the solution and store in the fridge*

i. *Aliquot?

1. The ketone collagen solution may be made into 10 µL

aliquots and frozen at -80C for storage. I’ve found that the

collagen clumps together within 2 months of being made

if kept in the fridge.

Plasma Treatment

See Appendix H for a detailed protocol on how to hydroxylate titanium slides.

In order to attach the silane-end of the oxyamine molecule, the titanium molecule must have –OH groups distributed on the surface. The plasma cleaner is in

Wickenden 215 by the blue fume hood. The instructions are also on a paper on top of the plasma cleaner. 106

Run the plasma cleaner according to the instructions in the document, treating for 15 minutes.

Oxyamine Surfaces

During this step, the oxyamine connector molecules are attached to the titanium surface. From now on, the titanium surface must not come into contact with the air!

Keep track of how much oxyamine in mL is used so it can be disposed of properly.

Normally, the “cleaning”, “plasma treatment”, “oxyamine surface” and subsequent “oxyamine-ketone-collagen slides” steps are done in one day. Start cleaning the titanium and holders as soon as you come in in the morning.

The oxyamine and ethanol used in this reaction need to be extracted from sure- seal protected bottles with a syringe. The syringes must be filled with an appropriate amount of dry nitrogen before extracting these substances so the pressure inside the container can be equalized following liquid extraction. The amount of each liquid extracted by the syringe is not accurate. The syringes should be emptied into labeled containers (centrifuge tubes or falcon tubes depending on the volume), and the final reaction solution should be prepared in a third container using micropipettes. The excess ethanol should be disposed of, but the excess oxyamine should be stored. The tube containing the excess oxyamine should be flooded with dry nitrogen, capped, wrapped in parafilm, and labeled with the date. The tube can then be put into the glass container in the fridge containing the rest of the oxyamine bottles and desiccant.

After the slides and holders have been cleaned and dried, this step takes about

1.5 hours overall. Start this reaction at about noon, finishing around 2 pm, so the next step is completed by 9 am. 107

Necessary Components:

• Hardware

a. Gloves

b. Goggles

c. Lab coat

d. Petri dish

e. Fluid exchange holders (Appendix B)

f. Curved Kelly forceps

g. Teflon tweezers

h. P-200 pipette and tips

i. P-1000 pipette and tips

j. Three microcentrifuge tubes

k. Centrifuge tube holder

l. Two 1 ml syringes

m. Two 3 inch needles

n. Two empty waste beakers

o. Nitrogen tank fitted with a filter and small nozzle tube

• Chemicals

a. Plasma-treated titanium slides

b. 200-proof ethanol

c. Oxyamine-silane molecules

d. Reagent-grade ethanol in a squeeze bottle

e. DI water in a squeeze bottle

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Protocol:

1. Put on gloves

2. Put on goggles and lab coat

3. Place the clean and dry holders upright in a clean Petri dish.

a. If there is any water left in the holders, the oxyamine will react with the

water rather than the titanium surface. Make sure they’re completely dry!

4. Carefully place the plasma cleaned titanium in the smallest part of the holder

using Teflon tweezers

5. Bring the holders in the Petri dish, the oxyamine bottle, ethanol, 3 centrifuge

tubes, centrifuge tube holder, two 1 mL syringes, two 3 inch needles, and two

empty waste beakers to the fume hood

6. Using a sharpie, label the three centrifuge tubes with “E” for ethanol, “O” for

oxyamine, and “mix” for the solution of ethanol and oxyamine. The syringes

extract an approximate amount of ethanol and oxyamine from their respective

bottles, but not an exact amount, so the correct amounts will need to be pipetted

into the “mix” container

7. Make a 5% solution of oxyamine in 200-proof ethanol in a 2 mL tube in the

fume hood, making 300 µL of solution for each holder (15µL of oxyamine in

285µL of ethanol per titanium slide)

a. Extract 285 µL of 200 proof ethanol per titanium slide out of the

membrane-covered 200 proof ethanol bottle

i. Put a syringe and needle together, leaving the protective plastic

tube on the needle

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ii. Go over to the nitrogen tank, open the tank with a steady flow of

clean nitrogen

iii. Flow nitrogen into the protective plastic tube on the needle and

fill the syringe with clean nitrogen, being careful not to touch the

needle to anything

iv. Walk back to the fume hood, remove the protective tube, and put

the needle through the protective membrane on the ethanol

container

v. Push the same volume of nitrogen out of the syringe that you

intend to remove (this equalizes the pressure inside the bottle after

you take out the ethanol, so no room-air is sucked into the

container)

vi. Extract the desired volume of ethanol

vii. Place extracted ethanol into the “E” microcentrifuge tube b. Using the p-1000 pipette, remove the desired amount of ethanol from the

“E” container and put it into the “mix” container and close the cap c. Use a 1 mL syringe and a long needle to extract the oxyamine through

the protective membrane

i. Use the same nitrogen-filling and fluid-extraction protocol used

for the 200 proof ethanol

ii. Place extracted oxyamine into the “O” microcentrifuge tube d. Using the p-100 pipette, remove the desired amount of oxyamine from

the “O” container and inject the oxyamine into the “mix” container under

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the surface of the ethanol to limit air exposure

e. Replace the cap on the glass oxyamine bottle. Wrap in parafilm. Place

back in the jar in the refrigerator.

f. The tube containing the excess oxyamine should be flooded with dry

nitrogen, capped, wrapped in parafilm, and labeled with the date. The

tube can then be put into the glass container in the fridge containing the

rest of the oxyamine bottles and desiccant and reused later.

g. Gently pipette the oxyamine solution up and down to mix evenly

8. Aliquot 300µL of oxyamine solution to each holder using a p-1000 micropipette

9. Let the solution sit on the titanium slides for 10 minutes

10. Rinse out the oxyamine – do NOT expose the surfaces to air!

a. Pick up the holder using the curved Kelly forceps

b. Hold the holder over an empty beaker to collect the waste ethanol

c. Using a squeeze bottle, carefully fill the holder to the top with ethanol

d. Overflow the holders with ethanol, letting the excess fall into the waste

beaker

e. Flow solution off the top for 1 minute so many dilutions have occurred

f. Spray the ethanol directly over the titanium surface to ensure that all of

the oxyamine is gone

g. Using the p-1000 pipette, suck out all but ~300 µL of ethanol to leave the

slide covered

h. Carefully repeat steps a-f with deionized water into a different waste

container

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i. Dispose of the ethanol-oxyamine solution in the labeled waste bottle

11. Place the holders back into the Petri dish

12. Using the p-1000 pipette, suck out some of the water, leaving 1 mL of water in

the holder

Oxyamine-Ketone Collagen Surfaces (oxime linkage reaction)

This section outlines how to conjugate the ketone collagens to the oxyamine surface.

Start the reaction at 4 pm! This reaction goes for 17 hours. You will rinse at 9 am the following morning.

Necessary Components:

• Hardware

o Gloves

o Paper towel

o Glass, high-walled Petri dish

o Domed glass cover for the Petri dish

o P-1000 pipette and tips

o P-10 pipette and tips

o Parafilm

• Chemicals

o Silanized titanium slides in fluid exchange holders under 1 mL of DI water

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o Oxime Linkage MES buffer (see section “buffers and rinsing solutions”)

o 2 mg/ml ketone-collagen stock solution (stored in -80℃ freezer)

o DI water in a squeeze bottle Protocol:

1. Put on gloves

2. Place the holders in a paper towel-lined glass Petri dish

3. Using the p-1000 pipette, rinse oxyamine-titanium in holders with Oxime

Linkage MES buffer at pH 5.5.

a. Suck excess water out of the holder, leaving the titanium surface safely

covered with water; approximately 300µL

b. Do not let the surface of the titanium touch air

c. Refill the holder with 1 mL buffer

d. Suck out 1 mL of buffer

e. Repeat steps b-d 6 times, so the fluid above the titanium is mostly buffer

4. Leave approximately 1000µL of buffer in each holder

5. Using the p-10 micropipette, inject 10µL of ~2 mg/ml ketone-collagen stock per

holder using a micropipette

a. The ketone-collagen stock is stored in the -80 C freezer in the Senyo lab

6. Using a squeeze bottle, Soak the towels lining the dish in DI water to maintain a

vapor pressure on the holders

7. Place the large round glass dome over the petri dish

8. Very carefully wrap the edges of the dish in parafilm

9. Leave at room temperature for 17 hours (overnight)

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10. Uncover the Petri dish

11. Rinse holders 6 times with DI water as described in step 2

12. Leave approximately 1000µL of water in each holder

Nucleation and Growth of fibrils (fibrillogenesis)

Necessary Components:

• Hardware

o Gloves

o Curved Kelly forceps

o 37℃ incubator (white)

o 30℃ incubator (gray)

o Petri dish

o 2 mL microcentrifuge tubes

o Stopwatch or other timing device, set to 1 minute

o P-1000 pipette and tips

o P-100 pipette and tips

o P-10 pipette and tips

• Chemicals

o Ketone-collagen-coated titanium slides in fluid exchange holders under 1 mL of DI water

o Cold PBS from the fridge (see section “Buffers and Rinsing Solutions”)

o Warm PBS in a squeeze bottle

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. Leave ~400 mL of PBS in a squeeze bottle the 30℃ incubator for

one hour

o 3 mg/mL PureCol collagen solution Protocol:

1. Put on gloves

2. Set up all necessary components by the gray 30C incubator

3. Rinse titanium in holders 6 times with room temperature 1x PBS

4. Remove most of the fluid in the holders, leaving about 300 µL of PBS in the

holder

5. Nucleation

a. Rinse the titanium in the holders for 20 seconds with the warm PBS in

the squeeze bottle.

b. Place holders back into Petri dish and put the squeeze bottle back into the

30C incubator

c. Remove excess PBS with the P-1000 pipette such that about 1 mL of

solution is left in the holders

d. Using the P-100 pipette, add 60 µL of PureCol to the holder solution and

mix gently

e. Start 1-minute timer

f. Put the lid back on the petri dish and place the holders and petri dish into

the 30C incubator

g. After 1 minute, remove holders from the incubator and rinse thoroughly

with the warm PBS in the squeeze bottle

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h. Repeat nucleation procedure 5 times

i. Finally, rinse thoroughly with warm PBS and set aside

6. Growth

a. Using the p-1000 pipette, rinse the holder 6 times with cold PBS

b. Remove excess PBS with the P-1000 pipette such that about 1 mL of

solution is left in the holders

c. Using the p-10 pipette, add 1 µL of PureCol to each holder

d. Mix gently with the p-1000 pipette

7. Place the holders in Petri dish lined with paper towels

8. Wet towels with DI water from a squeeze bottle

9. Place domed glass top on the petri dish

10. Wrap edges with parafilm

11. Let sit for 17 hours in the 37-degree Celsius incubator

12. Again, flow out the solution with DI water in a squeeze bottle

a. Flow until many dilutions have occurred

b. Spray the stream directly onto the surface to promote mixing

Buffers and Rinsing Solutions

Safety

General safety precautions for making buffer solutions

1. Before handling the acetic acid, put on the following

a. Splash goggles

b. Large nitrile gloves, located in cabinet 22

c. Synthetic apron 116

d. Lab coat

2. When working with the acid, work in the fume hood

3. NEVER add water to acid

4. ALWAYS add acid to water

a. The thermal coefficient of water prevents the solution from boiling and

splashing

5. Storage

a. Always label buffer solutions.

b. The label should have your name, the date, the chemicals, their respective

concentrations, and the pH written on it.

c. Optionally, the buffer could also have a label denoting its function (e.g.

Transamination MES Buffer)

Transamination MES Buffer: pH 6.5 with NaCl

This buffer is used when making the ketone collagen molecules with the RS reagent.

This recipe makes a buffer with the following characteristics:

• 30 mM MES

• 100 mM NaCl

• pH 6.5

Ingredients:

a. 100 mL DI water

b. 0.586 g MES free-acid (MW: 195.2 g/mol)

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c. 0.584 g NaCl (MW: 58.44 g/mol)

Protocol:

1. Fill 100 mL glass bottle with 100mL of deionized water and add stir bar

2. While stirring, add 0.586g MES

3. When dissolved, add 0.584 g NaCl

4. Adjust the pH using the pH meter and NaOH to pH 6.5.

5. Cap, label, and store the buffer in the refrigerator. The label should have your

name, the date, the chemical, its concentration, and the pH written on it.

6. Test the pH of the buffer before use every day.

Oxime Linkage MES Buffer: pH 5.5 with MgCl2

This buffer is used when conjugating the ketone-collagens to the oxyamine

surfaces. It has 100 mM of MgCl2 to prevent aggregation of the ketone collagen

molecules.

This recipe makes a buffer with the following characteristics:

• 30 mM MES

• 100 mM MgCl2

• pH 5.5

Ingredients:

a. 100 mL DI water

b. 0.586 g MES free-acid (MW: 195.2 g/mol)

c. 0.952 g MgCl2 (MW: 95.21 g/mol)

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Protocol:

1. Fill 100 mL glass bottle with 100mL of deionized water and add stir bar

2. While stirring, add 0.586g MES

3. When dissolved, add 0.952 g MgCl2

4. Adjust the pH using the pH meter and NaOH to pH 5.5.

5. Cap, label, and store the buffer in the refrigerator. The label should have your

name, the date, the chemical, its concentration, and the pH written on it.

6. Test the pH of the buffer before use every day.

KeCol Rinsing Solution: 150 mM MgCl2 (“ketone-collagen”)

Use this solution to rinse out the RS after the ketone collagen reaction. The 150 mM of MgCl2 prevents the aggregation of the collagen molecules. This is not a buffer, it

is just a salt solution, so the pH is not characterized.

This recipe makes a solution with the following characteristics:

• 150 mM MgCl2

Ingredients:

a. 100 mL DI water

b. 1.428 g of MgCl2 (MW: 95.21 g/mol)

Protocol:

1. Fill 100 mL glass bottle with 100mL of deionized water and add stir bar

2. While stirring, add 1.428 g MgCl2

3. Cap, label, and store the solution in the refrigerator. The label should have your

name, the date, the chemical, and its concentration written on it.

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PBS: Phosphate Buffered Saline

This buffer is used when growing fibrils on the titanium surfaces. The 10x PBS

solution by the incubator needs to be diluted 10x to make a 1x PBS solution. Whatever

volume you make, make sure the 10x PBS is diluted to 1x.

This recipe makes a buffer with the following characteristics:

• pH 7.4

• 10 mM phosphate

• 133 mM NaCl

Ingredients:

a. 45 mL DI water

b. 5 mL 10x PBS solution

Protocol:

1. Fill a 100 mL glass bottle with 5 mL of 10x PBS solution add stir bar

2. While stirring, add 40 mL of DI water

3. Measure and adjust pH to 7.4

4. Add DI water to fill beaker to 50 mL

5. Mix for 2 minutes

6. Cap, label, and store buffer in the refrigerator.

Determining Efficacy

UV-Vis Spectroscopy: Oxyamine-TAMRA reaction and analysis

This procedure is used to determine if the N-termini of the collagen molecules in the ketone-collagen step have been changed from an amine to a ketone. 120

The ketone collagens are dyed with the UV-active dye TAMRA. In this reaction,

an oxime linkage is formed between aminooxy-5(6)-TAMRA and the ketone on the collagen molecules.

TAMRA conjugation protocol:

1. Create ketone collagen, following the Ketone Collagen reaction procedure

through step 2 in Appendix A

2. Rinse out the excess RS using pH 5.5 Oxime Linkage MES buffer

a. Follow step 3 in the Ketone Collagen procedure, but rather than rinsing

with a 150 mM solution of MgCl2 in DI water, rinse with the pH 5.5

Oxime Linkage MES buffer

3. Retrieve the solution as stated in step 3 of the Ketone Collagen procedure

4. Add a 10x molar excess of aminooxy-5(6)-TAMRA to the collagen solution

a. Following the Ketone Collagen procedure, there should be ~ 0.75 mg

(2.5 nmol) of collagen in the sample solution

i. MW collagen ≈ 300,000 g/mol, or 300 kDa

b. The TAMRA-DMSO solution has a concentration of 5 mg/mL (612.96

µmol/L or 612.96 µM or 6.1296 mM)

i. MW aminooxy-5(6)-TAMRA = 815.71 g/mol

c. Need a 10x molar excess of TAMRA = 25 nmol

i. 2.5 nmol collagen*10 = 25 nmol of TAMRA dye needed

d. = 4.08 10 = 40 µ . −9 25 𝑚𝑚𝑚𝑚𝑚𝑚 𝑇𝑇𝑇𝑇𝑇𝑇𝑇𝑇𝑇𝑇 10 −5 𝑚𝑚𝑚𝑚𝑚𝑚 −6 612 96 𝐿𝐿 𝑇𝑇𝑇𝑇𝑇𝑇𝑇𝑇𝑇𝑇 ∗ 10 ∗ 𝐿𝐿 𝑇𝑇𝑇𝑇𝑇𝑇𝑇𝑇𝑇𝑇 𝐿𝐿 𝑜𝑜𝑜𝑜 𝑇𝑇𝑇𝑇𝑇𝑇𝑇𝑇𝑇𝑇 5. Gently pipette the solution up and down to mix

6. Incubate at room temperature for 17 hours (overnight)

121

5. Removal of Excess TAMRA dye - This step is particularly important because

any free TAMRA left in the solution will show up in your results. See the

instruction manual for the 10kDa or 100kDa microcentrifuge filters in the pink

box.

(~4 hr.)

a. Ingredients

i. TAMRA-collagen solution

ii. At least 50 mL of KeCol Rinsing Solution: 150 mM MgCl2

• You want to have enough buffer to thoroughly rinse all of

your samples. The DMSO is difficult to remove from the

solution

b. Procedure

i. Add all your TAMRA-collagen solution into the centrifugal filter

ii. Fill the filter tubes with 300-400µL of KeCol Rinsing Solution

iii. Run for 30 minutes at 9,000 rpm to rinse out all of the TAMRA

iv. Dump out the filtrate

v. Repeat rinses 10 times until the rinsate is no longer visibly

red/purple

vi. Collect rinsate from the 10th run, take to NanoDrop and measure

the 560 absorbances and compare it to DI water

1. Add 5 µL of DI water to stage

2. Collect 3 absorbance measurements

3. Wipe down the stage with a kimwipe

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4. Add another aliquot of DI water

5. Repeat 3 times

6. Repeat 3 times with rinsate

7. Wash stage with DI water

vii. If there is any 560 absorbance in the rinsate compared to the DI

water, rinse with the rinsing solution 2 more times and repeat

rinsate measurements

viii. Once there is no 560 nm absorbance, collect the TAMRA-

collagen for analysis

6. Retrieval of TAMRA-collagen solution. When you have rinsed out all the

TAMRA and concentrated the solution down to 100-200 µL, you have to get all

the solution out of the filters.

(~10 min)

a. Get out a clean centrifuge tube from the box

b. Remove the filter with the TAMRA-collagen solution from its used

centrifuge tube

c. Carefully turn the filter over into the clean centrifuge tube so it is now

upside down

d. Place the tube into the centrifuge with the lid pointing away from the

center of the centrifuge

e. Balance the centrifuge with other tubes of the same weight

f. Spin at 4000 rpm for 3 minutes to collect the solution

123

g. Analyze the solution using the UV-vis spectroscope, the NanoDrop 100

i. The process for analyzing the data using the NanoDrop can be

found in Appendix E

XPS

This machine is used to determine if the oxyamine molecules have been successfully conjugated with the titanium surface in the oxyamine functionalization step. Up to 5 slides can be analyzed during one appointment.

Schedule XPS time a few days before you plan to do the chemistry. Do not let the slides sit for more than 24 hours before using the XPS. The sooner you use the XPS after the slides have been prepared, the better. The XPS is extremely sensitive, so any dirt or dust that falls on the slides will also show up in the final chemical composition.

The XPS is in SCSAM on the first (lowest) floor of the Glennan building. Go to the lowest floor, then look for the large glass doors beside the staircase. Go through the doors and ring the doorbell labeled “XPS”.

To prepare samples for the XPS:

1. Follow the procedure in Appendix A until the oxyamine chemistry has been

performed on as many slides as desired

2. Prepare a clean titanium slide as a control

3. Rinse the slides in the holders with DI water

4. Using the sharp ended metal tweezers described in the Appendix E section

“Titanium”, place the sharp end of the tweezers in the small circular cut next to 124

the titanium slide in the holder

5. Gently lift the titanium piece with the sharp tweezers

6. Using the Teflon tweezers, gently grab the titanium slide

7. Place the slide on a clean Petri dish shiny side up and cover

8. Place in the 37-degree Celsius incubator until completely dry, approximately 15

minutes

9. Do not let the slides sit for more than 24 hours before using the XPS

10. 5 minutes before your appointment, carefully transport your slides to Glennan

a. Bring the slides in their Petri dishes, the Teflon tweezers, and your lab

notebook

b. Do not let the slides slide around in the Petri dish. They should not be

allowed to touch the sides or top of the Petri dish to ensure they stay as

clean as possible before doing XPS

11. Ring the doorbell labeled “XPS”

12. The technician will help you transfer the slides to the metal disk that will enter

the XPS, or do it himself

a. The slides will be stuck to a metal disk using copper tape and placed in

the vacuum chamber. Once the necessary pressure is reached, the disk

will be slid into the main XPS chamber.

13. Focus on looking for nitrogen, oxygen, and titanium. If the percent of the atoms

on the surface that are nitrogen increases significantly, then that suggests that the

chemistry was successful.

14. Look at the control slide first so you can compare the test slides to it

125

15. Ask to do a survey scan

a. A survey scan will give you the total atomic composition of a specific

spot on the titanium slide. It outputs percents, such as 5% nitrogen. If

there is a negligible amount of nitrogen on that spot, <1%, choose a new

spot on the surface and do a new survey scan.

16. If the slides show a significant amount of nitrogen in comparison to the control,

<2% more, then do a targeted scan

a. This scan will help you determine what kind of nitrogen compound is on

the surface based on the chemical shift. For example, you may see a peak

that correlates with an –NO group and a peak that correlates with a –NH2

group. The oxyamine has an amine, -NH2 on the end, so that is the kind

of shift we are concerned with

b. Later, analyze the peaks

c. NIST has tables at https://srdata.nist.gov/xps/EnergyTypeValSrch.aspx.

Under “choose energy type” select binding energy. Find where the peak

exists, such as 498, and look at what kinds of nitrogen compounds would

produce that peak. Any compound that ends in –NH2 is acceptable. The

NIST tables are long and must be carefully sorted through, so plan some

time for analysis

17. On a slide that gives good results, such as >3% nitrogen, check at least 3 spots to

ensure that the whole slide was successful.

Immunostaining: Immunofluorescent Imaging

Antibody staining is used to determine the efficacy of the ketone-collagen- 126

surface step above. To stain collagen, two types of antibodies are necessary. The primary antibody is specific to bovine type 1 collagen. The secondary antibody is attached to a fluorophore. The collagen-conjugated surfaces are exposed to the primary and then secondary antibodies and then observed with a fluorescence microscope in

Wickenden room 139, our lab downstairs. The primary antibodies are stored in the freezer in micro-centrifuge tubes in 5µL quantities. The secondary antibodies are in the refrigerator. Up to five slides can be stained at a time. The slides are incubated with the antibody for 60 minutes.

Staining Protocol:

1. Prepare slides conjugated with ketone-collagen and a control slide. Either:

a. Treat the control slide the same as the ketone-collagen slides, omitting

the ketone-collagen

b. Use a sonicated titanium slide

c. Prepare a titanium slide

2. Up to five slides can be stained using one microcentrifuge tube of primary

antibody, as there are 5µL of antibody solution in the tube to begin with

3. Set the slides, holders, and all equipment somewhere that doesn’t need to be

moved for an hour

a. Once the antibodies are on the slides, do not move the slides at all

4. Take one microcentrifuge tube containing primary antibody out of the freezer

5. Roll the tube around in your fingers to thaw the solution

6. Dilute primary antibody 1:80 in 1x PBS

127

a. Pipette 400µL of PBS into the microcentrifuge tube containing the

antibodies

b. Mix the fluid gently with the micropipette

c. Set aside briefly

7. Remove the slides from their holders and place upright in a clean Petri dish

a. In order to use the antibodies, the slides cannot be in the holders. The

volume of fluid is too small, and due to water adhesion the antibody

solution will move under the slide and the antibodies will not attach to

the surface.

8. Gently pipette 81µL of antibody solution onto each slide

a. Slowly push the fluid out of the pipette tip

b. The slides should be hydrophobic, so a bubble of fluid should form on

the slide

c. Do not allow the fluid to reach the edge of the slides, and do not touch

the slides until the experiment is over.

9. Incubate for 60 minutes at room temperature

10. Rinse 3 times with PBS

11. Dilute secondary antibody 1:80 in 1x PBS

a. Pipette 400µL of PBS into a clean microcentrifuge tube

b. Remove 5µL of secondary antibody solution

c. Mix the fluid gently with the micropipette

12. Gently pipette 81µL of antibody solution onto each slide

a. Slowly push the fluid out of the pipette tip

128

b. The slides should be hydrophobic, so a bubble of fluid should form on

the slide

c. Do not allow the fluid to reach the edge of the slides, and do not touch

the slides until the experiment is over.

13. Incubate for 30 minutes at room temperature

14. Rinse 3 times with PBS

15. Rinse 3 times with DI water

16. Ready to image

a. Place the slides in a Lab Snacks box to keep in the dark and to transport

downstairs

17. The labeled slides can be frozen for future imaging, but it is best to image when

fresh

Immunofluorescence Imaging:

The microscope is in Wickenden room 139. Use the 60x water-immersion lens.

The antibodies we use are conjugated with FITC. More information can be found in

Appendicies F and G.

129

1. Plug the camera into

2. Turn on the computer beside the microscope

3. Place the slide you wish to image on a glass coverslip

4. Place the coverslip under the lens

5. Place about 80µL of DI water on the slide

6. Lower the lens into the water droplet

SEM

Critical Point Drying:

1. A protocol for critical point drying is located on a binder beside the critical point

dryer

SEM Imaging:

We use the SEM to look for fully-formed collagen fibrils standing upright on the surface. Set up SEM time with SCSAM.

130

1. Schedule SEM time first

2. Start making the slides a few days before SEM

a. Prepare multiple slides, as getting the SEM ready takes a long time and

SEM time is expensive

b. The slides must go through cleansing procedures, all the chemistry, and

critical point drying in the days before SEM. Allow enough time to

prepare the slides.

3. Take the slides to White building

4. Sputter the slides with 15 nm of palladium

5. Stick the slides to posts with carbon tape

6. Put the slides in the Helios

7. After the operator calibrates the SEM, begin looking for fibrils

a. The fibrils will be many micrometers in length and a few nanometers in

diameter. See images below for fibrils under SEM.

b. Upright fibrils are brighter in color and will seem to “shine” on the

surface

c. Search around at low magnification, many micrometers to look for things

that look like they might be fibrils

d. If you find a patch, zoom in. Take pictures if the image looks like a fibril

upright or on its side

131

e. Upright fibrils will likely fall over as you view them due to the

bombardment of electrons. Take images quickly.

132

3D-Printed Holders

After the titanium has been plasma cleaned, all chemical steps detained in

Appendix A occur on the titanium inside these plastic holders. The holders allow for

fluid exchange, so the surface of the titanium never meets the fluid-air boundary, keeping the surface clean. Many pictures of the holders can be seen below

The holders are made of polycarbonate plastic, are white, and hold about 2 mL of fluid. The holders consist of two concentric wells. The smaller well at the bottom of the holder is just large enough to hold the titanium (1.1 cm2) and has a round cutout that allows the titanium to be removed with a sharp pair of tweezers.

The Solidworks file can be found under Z:\Ellie\The Good Stuff\3D Printing with the filename Holders_Solidworks_V3_LargerTiHole.SLDPRT

133

Solidworks drawing of fluid exchange holder

Images of holders in use

134

Ordering Oxyamine Molecule

The oxyamine molecule is 11-(O-hydroxylamine)undecyltriethoxysilane. We obtained this chemical from SikéMia in France. Here the original quotation.

135

Use of Selected Tools

This appendix outlines the purpose of and how to use some of the tools in the

Necessary Components section, including the scale and sonicator.

Using the Scale

Necessary Components:

1. Weighing boats or paper

a. Located in cabinet 22 beside the gloves, and beside the scale on the

counter

2. Chemical of choice

3. Scooper for chemical

Protocol:

The scale found on the right side of the lab is accurate to four decimal places.

To use the scale, first place your weighing boats or paper on the square metal scale plate inside the machine. Then, wave your hand over the left sensor. This will close the box over the scale plate. This prevents air movement from affecting the apparent weight of the paper or weighing boats.

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Then, wave your hand over the right sensor. This will make the scale zero to the weight of the weighing boats or paper so that measuring out chemicals is accurate. Be patient, this process takes a while.

When the machine is zeroed, the box will slide back and reopen, allowing you to put your chemical into the weighing boats or paper. Weigh out the appropriate amount of chemical.

Using the pH Meter

The pH of the buffers used in this protocol must be strictly controlled. Every day, the meter must be calibrated before use. Then, test the pH of any buffer one intends

137

to use that day. The calibration procedure is listed in the white binder beside the pH meter. The standards must be replaced once every three months.

Calibration

1. There are three standards on the shelf behind the pH meter – pH 4 (red), pH 7

(yellow), and pH 10 (blue)

2. Remove the probe from the tube it is housed in, rinse with DI water, and gently

dab with filter paper before placing it in the holder above the stir plate

3. Pictures:

4. Check to see that the standards have been replaced within the last 3 months. If

not, replace the standards.

5. Calibrate first with 7, then 4, then 10. This may take a very long time, up to half

an hour

6. The meter will beep once it stabilizes on each standard

7. Place the first standard on the stir plate, turn on the stir plate, and gently lower

the probe into the standard solution

8. Between standards, rinse the probe with DI water from a squeeze bottle, then

gently dab the probe on filter paper to dry it off 138

9. After the third standard, the meter will beep and briefly show the results of

calibration

10. Record the temperature and percentage listed on the meter’s screen in the white

binder

a. If the calibration failed, that means that the slope it calculated is

unsatisfactory (e.g. less than 90%) and the pH that it will measure will be

inaccurate. Try calibrating it again, or check to see if the standards have

been replaced within the last three months in the white binder.

Use

1. After calibration, rinse the probe with DI water, dab with filter paper, and place

it in the holder above the stir plate

2. Place the buffer on the stir plate and turn on the stir plate to speed 5

3. Lower the probe into the buffer

4. Wait until the pH stabilizes

5. Use 1 molar/normal sodium hydroxide or hydrochloric acid to adjust the pH

a. Put on gloves, goggles, and a lab coat before handling acids or bases

b. The acid is located in the acids cabinet under the fume hood, and the base

is in the bases cabinet beside the acid cabinet

c. Both sodium hydroxide and hydrochloric acid are in small containers

d. Use glass pipettes to transfer the acid or base to the buffer solution, one

drop at a time

e. Allow the pH meter to stabilize before adding more acid or base

Squeeze Bottles 139

The squeeze bottles are used for cleaning the components before doing chemistry, and to rinse chemicals off the titanium between steps

Soap

1. The soap squeeze bottles have alconox soap in them. The alconox powder can be

found below the sink in a white carton. For every 100mL of water, add 1g of

soap powder. These instructions are found on the alconox container.

Alcohols – Methanol, ethanol, isopropanol

1. Found on the counter behind the sonicator. All are labeled.

2. Methanol – MeOH

3. Ethanol – EtOH

4. Isopropanol

5. Refills for the squeeze bottles can be found in the solvents cabinet below the

fume hood labeled “flammable”

Waste – How to dispose of used solvents and chemicals

Used solvents and chemicals must be placed in the appropriate containers. All containers have green waste tags where lab workers must write down how much of each component is in the container. Contact EHS so they can remove the waste containers.

Their website has more information, at https://case.edu/ehs/waste-disposal/chemical- waste/how-to-dispose-of-chemical-waste/

Alcohol used for cleaning

Alcohols used to clean glassware or during sonication should be poured into the two-liter plastic jug labeled “used alcohols” in sharpie with the original label scratched

140

out (the alcohols dissolve sharpie, so be careful)

Oxyamine-contaminated solvent

Any solvent contaminated with the oxyamine must be disposed of separately from the alcohol used for cleaning. There is a plastic jug labeled “oxyamine waste” in the waste box under the sink. After completing the chemical steps involving oxyamine, pour waste into this jug and write down the amount of oxyamine in mL on the green waste tag.

Acetic acid buffer, PBS, collagen, ketone collagen, soapy water, water

These may be poured down the sink

141

UV-Vis Spectroscopy, NanoDrop 1000

I want to measure the absorbance of a concentrated dye+protein solution. I have modified some collagen molecules to express ketones rather than amines on the N- termini. To assess the efficacy of this reaction, I have dyed these ketones with TAMRA dye and I want to know about how many TAMRA molecules per collagen molecule I have – This is indicative of how many ketones per collagen I have. I am only expecting to see 1 dye molecules for every 10 collagens, so the sample solution needs to be very concentrated so I can see the TAMRA’s absorbance. This ratio is based on previous experiments (Collagen ~300 kDa, TAMRA ~800 Da). I’m expecting to see about 30 µM of TAMRA and 330 µM of collagen in my concentrated samples.

I cannot reliably make a set of collagen standards greater than 10 µM, as that is the concentration of the PureCol collagen we buy. Additionally, the 230 absorbance of a concentrated collagen solution is quite unstable, with a set of measurements all taken on the same sample at the same time having a standard deviation typically 10-100x greater than those of the standards. If I were to dilute the concentrated sample 30x to get the collagen to a reasonable concentration (10uM), then the TAMRA concentration would be about 1 µM and below the detection limit of the NanoDrop (~8 µM). For the above reasons, I obtain the TAMRA absorbance of a sample at the high concentration, then dilute that sample by a known amount (30x) to obtain the collagen absorbance. The diluted concentration measured is multiplied by 30 to obtain the concentrated collagen concentration and determine the ratio of dye/collagen. I make standard curves using

TAMRA (0-60 µM) and Collagen (0-10 µM). Peak absorbances: Collagen ~230 nm,

TAMRA ~550 nm.

142

Why not use a Bradford Assay?

The Bradford assay must use plate reader – Bradford assay absorbance is reliant

on time, so all samples need to be read simultaneously, which is impossible with

NanoDrop. The plate reader requires at least 200 µL of fluid (NanoDrop uses 1-5 µL). I

make 10 µL of concentrated ketone-collagen-TAMRA at a time, 200 µL would be A

LOT to make. The detection limit of the plate reader is ~8 µM of TAMRA. In order to detect both the TAMRA and the collagen at the same time, solutions must be concentrated. Concentrated solutions must contain at least 8 µM of TAMRA (detection limit), and given the expected 10:1 ratio of collagen-to-TAMRA, the solutions will contain ~80 µM of collagen. Therefore, the Bradford assay is NOT sensitive to protein concentrations above ~6 µM! I also cannot reliably make collagen standards above 10

µM (PureCol concentration). Given these considerations, I need to use the NanoDrop

and take measurements at the concentrated (for TAMRA) and dilute (for Collagen)

conditions

Sample Preparation

Concentration for NanoDrop:

• The ketone-collagen-TAMRA MgCl2 solutions, approximately 100uL in

volume, are frozen at -80C

• Lyophilize for 24 hours

• Reconstitute in 10uL of DI water (this time) 4 days before analysis to allow the

collagen to fully come back into solution

Dilution for NanoDrop

• After analyzing a volume of the concentrated sample to obtain the concentrated

TAMRA absorbance, the sample is diluted 30x to bring the collagen 143

concentration into the calibrated range

• This time, the remaining 7 µL of the TAMRA-ketone-collagen was diluted to

210 µL with DI water

Using the Nanodrop 1000

144

145

146

147

148

149

ImageJ processing of Immunofluorescence images

Samples are imaged with an upright light microscope (Nikon Eclipse E600FN) equipped with a digital camera (Blackfly S USB3, FLIR, Arlington, VA) and Spinnaker

SDK imaging software. Images are taken with the following parameters:

• Acquisition mode: Single

• Exposure time: 5 seconds

• Gain: 13.6

• Gamma: 0.8

• 60x water immersion lens

• 488 nm filter cube

The camera and microscope setup used to collect grayscale images of antibody- stained slides resulted in only partially in-focus images. The stage is slightly tilted, the camera is slightly out of focus with the eyepiece, and the FITC fluorophores photobleach so quickly that the sample must be put in-focus under the “FITC-safe” fluorescence cube which is out of focus with the FITC cube. The setup could use improvement to fix these issues. Since parts of the image that are out of focus have different brightness values than the true image, the images needed to be processed to crop out all of the not-in-focus portions of the images. Multiple images were taken of each slide in various places to confirm homogeneity. Using ImageJ, I transferred the in- focus parts of each image from each slide onto one single 7000x7000 pixel image with a black background. An image of this process can be found below.

150

151

ImmunoFluorBrightness.m

This code is used to determine the mean brightness of antibody-labeled ketone- collagen slides following ImageJ processing. (Appendix F).

clear

AllSamples = ["Sample1", "Sample2", "Sample3"]; cutpct = 98; % percentile cutoff for the bright-spot filter bn = [10 10 10]; % pre-defined bin numbers for histograms

showLabels = 1; % If showLables = 1, the histogram graphs will be populated with annotations and titles. Otherwise, the histograms are plotted without labels to make figure-making easier

data = {1,3}; % Saves brightness values from each image

for j = 1:length(AllSamples) sample = char(AllSamples(j)); % sample name, such as "OxyOnly" or “sample1” is pulled in file0 = [sample '.tif']; % image name and filetype fileType = '.tif'; % filetype I0 = imread(file0); % Reading in the image

% turn the image into a double vector and remove zeroes from the background (cropped % images pasted onto a black background from ImageJ) V0 = double(I0(:)); Idata = nonzeros(V0);

% make a logical vector of filtered data -> get indices of acceptable pixels r = Idata < prctile(Idata,cutpct,'all'); range = find(double(r));

% use range to filter out unacceptable, bright pixels from vector Ifilt = Idata(range);

%building the figure figure(1)

152

subplot(length(AllSamples),1,j) histogram(Ifilt, bn(j), 'normalization', 'probability') title([sample ' Histogram'])

% removes x and y-axis numbers if showLabels ~= 1 set(gca,'YTickLabel',[]) set(gca,'Xticklabel',[]) end

if showLabels == 1 if j == length(AllSamples) xlabel('grayscale value') end ylabel('count') txt = {['\bf\mu: \rm' num2str(mean(Ifilt), '%.2g')],... ['\bf\sigma: \rm' num2str(std(Ifilt), '%.0f')],... ['n: \rm' num2str(length(Ifilt), '%.2g')]}; ylim([0,0.35]) xlim([0,2*10^4]) text(0.3*10^4, 0.2, txt) end

% removes x and y-axis numbers if showLabels ~= 1 set(gca,'YTickLabel',[]) set(gca,'Xticklabel',[]) end % % calculate the percent of bright pixels removed from the data RmPixelsPct(j) = 100 * (length(Idata) - length(Ifilt)) / length(Ifilt);

data{j} = Ifilt; end

153

Plasma Cleaner Operation

The plasma cleaner is in Wickenden 215 by the blue fume hood. The instructions below are also on a paper on top of the plasma cleaner. The machine is finicky and it takes practice to get it working consistently. Please read through the instructions before beginning.

Necessary Components:

1. Plasma Cleaner

2. Clean, inert, stable, rectangular platform (glass slide, Plexiglas rectangle, etc.)

• If you are very careful, a petri dish can be used

3. Sample (hard, non-volatile solid or powder)

4. Gloves

DANGER:

This machine operates on very high voltages. Ensure that the “RF

POWER” switch is flipped DOWN before opening the chamber at any time.

Notes:

• This process takes between 30 minutes and an hour depending on if the machine

works properly the first time

• Be gentle when turning nobs. The gas-line valve knobs should only be barely

154

finger tight. They are pin valves and may break.

• NO GLUES/TAPES/GELS/STICKY THINGS. Under the vacuum and plasma

treatment, these substances will vaporize and make the chamber and your sample

very dirty.

• NO LARGE METAL OBJECTS. Objects larger than a quarter get very hot

inside the chamber.

• Plasma treatment will reach all around objects

. The underside of glass coverslips will get coated with OH groups

• Plasma treat samples on a stable rectangular platform

. Petri dishes can rock and throw your samples around. If you are very

careful, a petri dish can be used.

• Notes for those treating small, delicate objects like AFM tips or powders:

. See Ellie (epm43) first

. If the vacuum chamber’s pressure changes quickly, your samples

WILL be thrown around.

. Take care to place the objects in/on a heavy, stable, inert platform

such as a glass slide or hard plastic rectangle. You cannot use tape or

gel to hold them down, as you will make the chamber and your

sample dirty.

. The slow vacuum is your best friend. Do not open purge below 2

Torr.

155

Before you begin:

1. Check that the bubbler lines are securely connected and that the water level in

the bubbler is about an inch above the white nub inside. Fill with DI water as

needed

2. Argon tank regulator is closed and all pressure gauges read zero

3. On the two BRANSON/IPC boxes

a. Gas line 2 switch off. Valve closed (fully in, clockwise)

b. Purge switch off. Purge valve closed (fully in, clockwise)

c. RF Power switch off.

d. Vacuum switch off.

e. Plug in the vacuum pump and plasma cleaner

i. Ignore the blue machine with the light switch. The vacuum pump

is the gray metal box labeled 5 on the bottom shelf. It is

controlled by the metal flip-switches on the metal cart.

4. DO NOT adjust

a. Vacuum valve knob

b. RF power knob

i. Is set to 50 watts of FWD power

156

Protocol:

1. Plug in pump and plasma cleaner

2. Turn system AC POWER on

3. Turn on vacuum pump (the switch on the cart below the yellow button).

4. Put samples in vacuum chamber

5. Close metal chamber door

6. Turn vacuum switch to slow

7. Wait ~15 seconds. Watch the pressure in the chamber slowly decrease

8. Turn vacuum to fast

9. Wait for pressure to reach 200 mTorr

10. Turn on Argon tank beside the plasma cleaner

a. Open the main tank valve

b. Slowly open the smaller regulator valve until the bubbler bubbles

i. Turn the valve. Once you feel resistance, go slowly. Bubbling

will start shortly after. It does not take much argon to run this

machine.

ii. The bubbles will slow down, that’s ok

c. Let the regulator pressure get to 10 kPa (the first black tick)

d. Check the pressure during cleaning to ensure it stays around 10 kPa

11. Turn on gas line 2, flip switch up

12. Turn the gas line 2 knob to flood the bubbler and chamber with argon

a. Let it reach 2 Torr

b. Wait for one minute.

157

c. Turn the gas knob again down to 200mTorr

d. Flood the chamber to 2 Torr again

e. Bring down to 200mTorr

f. Back to 2 Torr

g. Bring down to 500mTorr (0.5 Torr). The plasma cleaner operates best at

this pressure.

13. Check that the Argon regulator still reads 10 kPa (first black tick)

14. If the RF Power light has turned on, turn RF on. FWD power = 50 watts

15. Flip RF switch to REFL

16. Turn “coarse impedance” knob counterclockwise to bring the REFL gage close

to 0

17. Treat for 15 minutes

a. Monitor the pressure in the chamber and the Argon regulator while

treating

18. Turn RF off

19. Turn vacuum to slow. Wait until the pressure gauge is fairly stable

20. Turn vacuum off. Let the argon bring the pressure up to at least 2 Torr

21. Turn purge on and open the purge valve VERY slowly (tiny finger movements)

until the chamber is at atmospheric pressure

a. You will hear hissing while air enters the chamber

22. Remove samples

23. Close main argon valve, regulator open

24. Allow gas pressure to come to zero. Turn “gas line 2” valve up to let the argon

158

out. Do not leave until regulator gage reads zero

a. Don’t touch the smaller regulator valve to let more argon into the

bubbler! Leave it at the first black tick mark. The bubbler cannot handle

very much pressure and it may break

25. Turn vacuum pump off on the cart

26. Turn system power off

27. Leave it as you found it, see before you begin for reference

159

Troubleshooting:

Contact [email protected], Ellie Miller, with any questions.

• If the vacuum chamber is not glowing with plasma, turn the coarse knob

clockwise until the plasma strikes

• Flip to REFL and adjust coarse impedance knob until gage is as close to 0 as

possible, about 10 W at best

• Check if the plasma cleaner is working correctly by looking at the color of the

plasma. Below are examples of what the vacuum tube might look like. The color

of the plasma will change as the impedance is adjusted, as well. Depending on

the sample, the color changes as well. Inert materials such as clean titanium,

glass, and Petri dishes have the best chance of working properly.

a. Perfect – blue/purple plasma. This plasma color is produced by argon

with an inert sample. The chamber is mostly argon, there are no leaks,

and your sample isn’t gassing or a large metal object. The bubbler was

properly evacuated of nitrogen. below is a perfect example of the plasma

cleaner working correctly.

•

b. Acceptable – between blue and red. Images below. This is not perfect but 160

does work. The machine can be finicky.

i. c. Incorrect – red/orange plasma. Images below. This color may be

produced by your specific sample. If the sample is small and inert, then

this plasma color is produced by nitrogen, and it means that there is a

significant leak in the vacuum chamber or there is not enough argon in

the chamber.

161

d. If the plasma is intensely red/orange, like the images directly above,

i. Turn off the RF power switch

ii. Flood the chamber with argon, as in step 18, and try again e. If it still doesn’t work, follow the steps below (25 onward) to let the

chamber come back to atmospheric pressure. Then,

i. Check that the bubbler lines are secure

ii. Open plasma cleaner

iii. Adjust the faceplate of the vacuum chamber by lifting it slightly

and pushing it against the chamber. Hold the plate gently against

the chamber while turning the slow vacuum back on. Let go and

close the front plate

iv. Pump the machine down and try again

162

Bibliography:

(1) American Dental Association. ADA 2005-6 Survey of Dental Services Rendered;

2006.

(2) Lang, N. P.; Berglundh, T. Periimplant Diseases: Where Are We Now? -

Consensus of the Seventh European Workshop on Periodontology. In Journal of

Clinical Periodontology; 2011; Vol. 38, pp 178–181.

https://doi.org/10.1111/j.1600-051X.2010.01674.x.

(3) Davis, L. G.; Ashworth, P. D.; Spriggs, L. S. Psychological Effects of Aesthetic

Dental Treatment. J. Dent. 1998, 26 (7), 547–554. https://doi.org/10.1016/S0300-

5712(97)00031-6.

(4) Berglundh, T.; Persson, L.; Rn Klinge, B. A Systematic Review of the Incidence

of Biological and Technical Complications in Implant Dentistry Reported in

Prospective Longitudinal Studies of at Least 5 Years. J. Clin. Periodontol. 2002,

29 (s3), 197–212.

(5) Fransson, C.; Lekholm, U.; Jemt, T.; Berglundh, T. Prevalence of Subjects with

Progressive Bone Loss at Implants. Clin. Oral Implants Res. 2005, 16 (4), 440–

446. https://doi.org/10.1111/j.1600-0501.2005.01137.x.

(6) Roos-Jansaker, A. M.; Lindahl, C.; Renvert, H.; Renvert, S.; Roos-Jansåker, A.

M.; Lindahl, C.; Renvert, H.; Renvert, S.; Roos-Jansaker, A. M.; Lindahl, C.; et

al. Nine- to Fourteen-Year Follow-up of Implant Treatment. Part I: Implant Loss

and Associations to Various Factors. J. Clin. Periodontol. 2006, 33 (4), 283–289.

https://doi.org/10.1111/j.1600-051X.2006.00907.x.

(7) Roos-Jansåker, A. M.; Lindahl, C.; Renvert, H.; Renvert, S. Nine- to Fourteen-

163

Year Follow-up of Implant Treatment. Part II: Presence of Peri-Implant Lesions.

J. Clin. Periodontol. 2006, 33 (4), 290–295. https://doi.org/10.1111/j.1600-

051X.2006.00906.x.

(8) Roos-Jansåker, A.-M.; Renvert, H.; Lindahl, C.; Renvert, S. Nine- to Fourteen-

Year Follow-up of Implant Treatment. Part III: Factors Associated with Peri-

Implant Lesions. J. Clin. Periodontol. 2006, 33 (4), 296–301.

https://doi.org/10.1111/j.1600-051x.2006.00908.x.

(9) Cosyn, J.; Eghbali, A.; De Bruyn, H.; Collys, K.; Cleymaet, R.; De Rouck, T.

Immediate Single-Tooth Implants in the Anterior Maxilla: 3-Year Results of a

Case Series on Hard and Soft Tissue Response and Aesthetics. J. Clin.

Periodontol. 2011, 38 (8), 746–753. https://doi.org/10.1111/j.1600-

051X.2011.01748.x.

(10) Albrektsson, T.; Buser, D.; Sennerby, L. Crestal Bone Loss and Oral Implants.

Clin. Implant Dent. Relat. Res. 2012, 14 (6), 783–791.

https://doi.org/10.1111/cid.12013.

(11) Cho-Yan Lee, J.; Mattheos, N.; Nixon, K. C.; Ivanovski, S. Residual Periodontal

Pockets Are a Risk Indicator for Peri-Implantitis in Patients Treated for

Periodontitis. Clin. Oral Implants Res. 2012, 23 (3), 325–333.

https://doi.org/10.1111/j.1600-0501.2011.02264.x.

(12) Browaeys, H.; Defrancq, J.; Dierens, M. C. A. A.; Miremadi, R.; Vandeweghe,

S.; Van de Velde, T.; De Bruyn, H. A Retrospective Analysis of Early and

Immediately Loaded Osseotite Implants in Cross-Arch Rehabilitations in

Edentulous Maxillas and Mandibles up to 7 Years. Clin. Implant Dent. Relat. Res.

164

2013, 15 (3), 380–389. https://doi.org/10.1111/j.1708-8208.2011.00367.x.

(13) De Bruyn, H.; Bouvry, P.; Collaert, B.; De Clercq, C.; Persson, G. R.; Cosyn, J.

Long-Term Clinical, Microbiological, and Radiographic Outcomes of

BrånemarkTM Implants Installed in Augmented Maxillary Bone for Fixed Full-

Arch Rehabilitation. Clin. Implant Dent. Relat. Res. 2013, 15 (1), 73–82.

https://doi.org/10.1111/j.1708-8208.2011.00359.x.

(14) Faggion, C. M.; Listl, S.; Tu, Y. K. Assessment of Endpoints in Studies on Peri-

Implantitis Treatment - A Systematic Review. J. Dent. 2010, 38 (6), 443–450.

https://doi.org/10.1016/j.jdent.2010.03.003.

(15) Faggion, C. M.; Giannakopoulos, N. N. Quality of Reporting in Abstracts of

Randomized Controlled Trials Published in Leading Journals of Periodontology

and Implant Dentistry: A Survey. J. Periodontol. 2012, 83 (10), 1251–1256.

https://doi.org/10.1902/jop.2012.110609.

(16) Faggion, C. M.; Chambrone, L.; Listl, S.; Tu, Y.-K. Network Meta-Analysis for

Evaluating Interventions in Implant Dentistry: The Case of Peri-Implantitis

Treatment. Clin. Implant Dent. Relat. Res. 2013, 15 (4), 576–588.

https://doi.org/10.1111/j.1708-8208.2011.00384.x.

(17) Eke, P. I.; Dye, B. A.; Wei, L.; Thornton-Evans, G. O.; Genco, R. J. Prevalence

of Periodontitis in Adults in the United States: 2009 and 2010. J. Dent. Res. 2012,

91 (10), 914–920. https://doi.org/10.1177/0022034512457373.

(18) CDC: Half of American Adults Have Periodontal Disease | Perio.org

https://www.perio.org/consumer/cdc-study.htm (accessed Feb 20, 2020).

(19) Palomo, L.; Terézhalmy, G. T. Peri-Implant Disease: Pathogenesis, Risk Factors,

165

Diagnosis, Prevention and Treatment. Crest + Oral-B Dent. Contin. Educ.

Course 2014.

(20) Moraschini, V.; Poubel, L. A. D. C.; Ferreira, V. F.; Barboza, E. D. S. P. P.

Evaluation of Survival and Success Rates of Dental Implants Reported in

Longitudinal Studies with a Follow-up Period of at Least 10 Years: A Systematic

Review. Int. J. Oral Maxillofac. Surg. 2015, 44 (3), 377–388.

https://doi.org/10.1016/j.ijom.2014.10.023.

(21) Davies, J. E.; Lowenberg, B.; Shiga, A. The Bone-Titanium Interface in Vitro. J.

Biomed. Mater. Res. 1990, 24, 1289–1306.

(22) Noguchi, S.; Ukai, T.; Kuramoto, A.; Yoshinaga, Y.; Nakamura, H.; Takamori,

Y.; Yamashita, Y.; Hara, Y. The Histopathological Comparison on the

Destruction of the Periodontal Tissue between Normal Junctional Epithelium and

Long Junctional Epithelium. J. Periodontal Res. 2017, 52 (1), 74–82.

https://doi.org/10.1111/jre.12370.

(23) Richter, E. J. In Vivo Vertical Forces on Implants. Int. J. Oral Maxillofac.

Implants 10 (1), 99–108.

(24) Waltimo, A.; Könönen, M. A Novel Bite Force Recorder and Maximal Isometric

Bite Force Values for Healthy Young Adults. Eur. J. Oral Sci. 1993, 101 (3),

171–175. https://doi.org/10.1111/j.1600-0722.1993.tb01658.x.

(25) McCormack, S. W.; Witzel, U.; Watson, P. J.; Fagan, M. J.; Gröning, F. Inclusion

of Periodontal Ligament Fibres in Mandibular Finite Element Models Leads to an

Increase in Alveolar Bone Strains. PLoS One 2017, 12 (11), e0188707.

https://doi.org/10.1371/journal.pone.0188707.

166

(26) Behrents, R. Growth in the Aging Craniofacial Skeleton (Craniofacial Growth

Series- Monograph 17), 1st ed.; Center for Human Growth and Development,

University of Michigan School of Education: Ann Arbor, 1985.

(27) Sharma, P.; Arora, A.; Valiathan, A. Age Changes of Jaws and Soft Tissue

Profile. Scientific World Journal. Hindawi Publishing Corporation 2014.

https://doi.org/10.1155/2014/301501.

(28) Mohan, R.; Singh, A.; Gundappa, M. Three-Dimensional Imaging in Periodontal

Diagnosis - Utilization of Cone Beam Computed Tomography. J. Indian Soc.

Periodontol. 2011, 15 (1), 11–17. https://doi.org/10.4103/0972-124X.82256.

(29) Naveh, G. R. S. S.; Lev-Tov Chattah, N.; Zaslansky, P.; Shahar, R.; Weiner, S.

Tooth-PDL-Bone Complex: Response to Compressive Loads Encountered during

Mastication - A Review. Arch. Oral Biol. 2012, 57 (12), 1575–2584.

https://doi.org/10.1016/j.archoralbio.2012.07.006.

(30) Shoulders, M. D.; Raines, R. T. Collagen Structure and Stability. Annu. Rev.

Biochem. 2009, 78 (1), 929–958.

https://doi.org/10.1146/annurev.biochem.77.032207.120833.

(31) Marchesan, J. T.; Scanlon, C. S.; Soehren, S.; Matsuo, M.; Kapila, Y. L.

Implications of Cultured Periodontal Ligament Cells for the Clinical and

Experimental Setting: A Review. Archives of Oral Biology. Pergamon October 1,

2011, pp 933–943. https://doi.org/10.1016/j.archoralbio.2011.03.003.

(32) de Jong, T.; Bakker, A. D.; Everts, V.; Smit, T. H. The Intricate Anatomy of the

Periodontal Ligament and Its Development: Lessons for Periodontal

Regeneration. Journal of Periodontal Research. Blackwell Munksgaard

167

December 1, 2017, pp 965–974. https://doi.org/10.1111/jre.12477.

(33) Wagner, D.; El Hafci, H.; Bensidhoum, M.; Petite, H.; Bolender, Y.; Rémond, Y.;

George, D. Periodontal Ligament Histology for Orthodontic Bone Remodeling:

First Quantification. J. Cell. Immunother. 2018, 4 (1), 41–43.

https://doi.org/10.1016/j.jocit.2018.09.010.

(34) Ho, S. P.; Sulyanto, R. M.; Marshall, S. J.; Marshall, G. W. The Cementum-

Dentin Junction Also Contains Glycosaminoglycans and Collagen Fibrils. J.

Struct. Biol. 2005, 151 (1), 69–78. https://doi.org/10.1016/j.jsb.2005.05.003.

(35) Dean, R. The Periodontal Ligament : Development , Anatomy and Function. J.

Oral Heal. Dent. Manag. 2017, 16 (6), 1–7.

(36) Wesselink, P. R.; Beertsen, W. Ankylosis of the Mouse Molar after Systemic

Administration of 1‐hydroxyethylidene‐1,1‐bisphosphonate (HEBP). J. Clin.

Periodontol. 1994, 21 (7), 465–471. https://doi.org/10.1111/j.1600-

051X.1994.tb00409.x.

(37) Cho, M. Il; Garant, P. R. Development and General Structure of the

Periodontium. Periodontol. 2000 2000, 24 (1), 9–27.

https://doi.org/10.1034/j.1600-0757.2000.2240102.x.

(38) Wills, D. J.; Picton, D. C. A.; Davies, W. I. R. The Intrusion of the Tooth for

Different Loading Rates. J. Biomech. 1978, 11 (10–12), 429–434.

https://doi.org/10.1016/0021-9290(78)90054-4.

(39) Kuo, C. K.; Marturano, J. E.; Tuan, R. S. Novel Strategies in Tendon and

Ligament Tissue Engineering: Advanced Biomaterials and Regeneration Motifs.

Sport. Med. Arthrosc. Rehabil. Ther. Technol. 2010, 2 (1), 2–20.

168

https://doi.org/10.1186/1758-2555-2-20.

(40) Kew, S. J.; Gwynne, J. H.; Enea, D.; Abu-Rub, M.; Pandit, A.; Zeugolis, D.;

Brooks, R. A.; Rushton, N.; Best, S. M.; Cameron, R. E. Regeneration and Repair

of Tendon and Ligament Tissue Using Collagen Fibre Biomaterials. Acta

Biomater. 2011, 7 (9), 3237–3247. https://doi.org/10.1016/j.actbio.2011.06.002.

(41) Gross, J.; Highberger, J. H.; Schmitt, F. O. Extraction of Collagen from

Connective Tissue by Neutral Salt Solutions. Proc. Natl. Acad. Sci. 1955, 41 (1),

1–7. https://doi.org/10.1073/pnas.41.1.1.

(42) Paten, J. A.; Siadat, S. M.; Susilo, M. E.; Ismail, E. N.; Stoner, J. L.; Rothstein, J.

P.; Ruberti, J. W. Flow-Induced Crystallization of Collagen: A Potentially

Critical Mechanism in Early Tissue Formation. ACS Nano 2016, 10 (5), 5027–

5040. https://doi.org/10.1021/acsnano.5b07756.

(43) Cheng, X.; Gurkan, U. A.; Dehen, C. J.; Tate, M. P.; Hillhouse, H. W.; Simpson,

G. J.; Akkus, O. An Electrochemical Fabrication Process for the Assembly of

Anisotropically Oriented Collagen Bundles. Biomaterials 2008, 29 (22), 3278–

3288. https://doi.org/10.1016/j.biomaterials.2008.04.028.

(44) Gretzer, C.; Mörgelin, M. US9295531B2 - Collagen Coated Article, 2012.

(45) Farber, S.; Garg, A. K.; Birk, D. E.; Silver, F. H. Collagen Fibrillogenesis in

Vitro: Evidence for Pre-Nucleation and Nucleation Steps. Int. J. Biol. Macromol.

1986, 8 (1), 37–42. https://doi.org/10.1016/0141-8130(86)90069-3.

(46) Gelman, R.; Poppke, D.; Piez, K. Collagen Fibril Formation in Vitro. The Role of

the Nonhelical Terminal Regions. J. Biol. Chem. 1979, 254 (22), 11741–11745.

(47) Holmes, D. F.; Capaldi, M. J.; Chapman, J. A. Reconstitution of Collagen Fibrils

169

in Vitro; the Assembly Process Depends on the Initiating Procedure. Int. J. Biol.

Macromol. 1986, 8 (3), 161–166. https://doi.org/10.1016/0141-8130(86)90020-6.

(48) Phosphate-Buffered Saline (PBS). Cold Spring Harb. Protoc. 2006, 2006 (1),

pdb.rec8247. https://doi.org/10.1101/pdb.rec8247.

(49) Baskin, J. Z.; Soenjaya, Y.; McMasters, J.; Ko, A.; Vasanji, A.; Morris, N.;

Eppell, S. J. Nanophase Bone Substitute for Craniofacial Load Bearing

Application: Pilot Study in the Rodent. J. Biomed. Mater. Res. Part B Appl.

Biomater. 2018, 106 (2), 520–532. https://doi.org/10.1002/jbm.b.33857.

(50) Harris, J. R.; Soliakov, A.; Lewis, R. J. In Vitro Fibrillogenesis of Collagen Type

I in Varying Ionic and PH Conditions. Micron 2013, 49, 60–68.

https://doi.org/10.1016/j.micron.2013.03.004.

(51) Harris, J. R.; Reiber, A. Influence of Saline and PH on Collagen Type I

Fibrillogenesis in Vitro: Fibril Polymorphism and Colloidal Gold Labelling.

Micron 2007, 38 (5), 513–521. https://doi.org/10.1016/j.micron.2006.07.026.

(52) Gelman, R.; Williams, B. R.; Piez, K. A. Collagen Fibril Formation. Evidence for

a Multistep Process. J. Biol. Chem. 1979, 254 (1), 180–186.

(53) Williams, B. R.; Gelman, R. A.; Poppke, D. C.; Piez, K. Collagen Fibril

Formation. Optimal in Vitro Conditions and Preliminary Kinetic Results. J. Biol.

Chem. 1978, 253 (18), 6578–6585.

(54) Stephanopoulos, N.; Francis, M. B. Choosing an Effective Protein

Bioconjugation Strategy. Nature Chemical Biology. Nature Publishing Group

2011, pp 876–884. https://doi.org/10.1038/nchembio.720.

(55) ncbi.nlm.nih. RecName: Full=Collagen Alpha-1(I) Chain; AltName: Full=Alpha-

170

1 Type I - Protein - NCBI. 2010.

(56) Ncbi.nlm.nih. Pre-pro-Alpha-2 Type I Collagen [Homo Sapiens] - Protein -

NCBI. 2002.

(57) Witus, L. S.; Francis, M. B. Site-Specific Protein Bioconjugation via a Pyridoxal

5′- Phosphate-Mediated N-Terminal Transamination Reaction. Curr. Protoc.

Chem. Biol. 2010, 2 (2), 125–134.

https://doi.org/10.1002/9780470559277.ch100018.

(58) NCBI.NLM.NIH.gov. Collagen Type 1 Alpha-1 Collagen Amino Acid Sequence.

https://doi.org/10.5061/DRYAD.BP20K.

(59) Witus, L. S.; Francis, M. B. Site-Specific Protein Bioconjugation via a Pyridoxal

5′- Phosphate-Mediated N-Terminal Transamination Reaction. Curr. Protoc.

Chem. Biol. 2010, 2 (2), 125–134.

https://doi.org/10.1002/9780470559277.ch100018.

(60) Scheck, R. A.; Francis, M. B. Regioselective Labeling of Antibodies through N-

Terminal Transamination. ACS Chem. Biol. 2007, 2 (4), 247–251.

https://doi.org/10.1021/cb6003959.

(61) Rosen, C. B.; Francis, M. B. Targeting the N Terminus for Site-Selective Protein

Modification. Nature Chemical Biology. Nature Publishing Group July 1, 2017,

pp 697–705. https://doi.org/10.1038/nchembio.2416.

(62) Gilmore, J. M.; Scheck, R. A.; Esser-Kahn, A. P.; Joshi, N. S.; Francis, M. B. N-

Terminal Protein Modification through a Biomimetic Transamination Reaction.

Angew. Chemie - Int. Ed. 2006, 45 (32), 5307–5311.

https://doi.org/10.1002/anie.200600368.

171

(63) Scozzafava, A.; Mastrolorenzo, A.; Supuran, C. T. Agents That Target Cysteine

Residues of Biomolecules and Their Therapeutic Potential. Expert Opinion on

Therapeutic Patents. Taylor & Francis 2001, pp 765–787.

https://doi.org/10.1517/13543776.11.5.765.

(64) Dixon, H. B. F. N-Terminal Modification of Proteins-a Review. Journal of

Protein Chemistry. Kluwer Academic Publishers-Plenum Publishers February

1984, pp 99–108. https://doi.org/10.1007/BF01024840.

(65) Scheck, R. A.; Dedeo, M. T.; Iavarone, A. T.; Francis, M. B. Optimization of a

Biomimetic Transamination Reaction. J. Am. Chem. Soc. 2008, 130 (35), 11762–

11770. https://doi.org/10.1021/ja802495w.

(66) Esser-Kahn, A. P.; Francis, M. B. Protein-Cross-Linked Polymeric Materials

through Site-Selective Bioconjugation. Angew. Chemie - Int. Ed. 2008, 47 (20),

3751–3754. https://doi.org/10.1002/anie.200705564.

(67) Esser-Kahn, A. P.; Iavarone, A. T.; Francis, M. B. Metallothionein-Cross-Linked

Hydrogels for the Selective Removal of Heavy Metals from Water. J. Am. Chem.

Soc. 2008, 130 (47), 15820–15822. https://doi.org/10.1021/ja807095r.

(68) Witus, L. S.; Moore, T.; Thuronyi, B. W.; Esser-Kahn, A. P.; Scheck, R. A.;

Iavarone, A. T.; Francis, M. B. Identification of Highly Reactive Sequences for

PLP-Mediated Bioconjugation Using a Combinatorial Peptide Library. J. Am.

Chem. Soc. 2010, 132 (47), 16812–16817. https://doi.org/10.1021/ja105429n.

(69) Carrico, Z. M.; Farkas, M. E.; Zhou, Y.; Hsiao, S. C.; Marks, J. D.; Chokhawala,

H.; Clark, D. S.; Francis, M. B. N-Terminal Labeling of Filamentous Phage to

Create Cancer Marker Imaging Agents. ACS Nano 2012, 6 (8), 6675–6680.

172

https://doi.org/10.1021/nn301134z.

(70) Christman, K. L.; Broyer, R. M.; Tolstyka, Z. P.; Maynard, H. D. Site-Specific

Protein Immobilization through N-Terminal Oxime Linkages. J. Mater. Chem.

2007, 17 (19), 2021–2027. https://doi.org/10.1039/b618002g.

(71) Gao, W.; Liu, W.; Mackay, J. A.; Zalutsky, M. R.; Toone, E. J.; Chilkoti, A. In

Situ Growth of a Stoichiometric PEG-like Conjugate at a Protein’s N-Terminus

with Significantly Improved Pharmacokinetics. Proc. Natl. Acad. Sci. U. S. A.

2009, 106 (36), 15231–15236. https://doi.org/10.1073/pnas.0904378106.

(72) Lempens, E. H. M.; Helms, B. A.; Merkx, M.; Meijer, E. W. Efficient and

Chemoselective Surface Immobilization of Proteins by Using Aniline-Catalyzed

Oxime Chemistry. ChemBioChem 2009, 10 (4), 658–662.

https://doi.org/10.1002/cbic.200900028.

(73) Kim, S. E.; Lee, P. W.; Pokorski, J. K. Biologically Triggered Delivery of EGF

from Polymer Fiber Patches. ACS Macro Lett. 2017, 6 (6), 593–597.

https://doi.org/10.1021/acsmacrolett.7b00212.

(74) Caley, M. P.; Martins, V. L. C.; O’Toole, E. A. Metalloproteinases and Wound

Healing. Adv. Wound Care 2015, 4 (4), 225–234.

https://doi.org/10.1089/wound.2014.0581.

(75) Palla, K. S.; Witus, L. S.; Mackenzie, K. J.; Netirojjanakul, C.; Francis, M. B.

Optimization and Expansion of a Site-Selective N-Methylpyridinium-4-

Carboxaldehyde-Mediated Transamination for Bacterially Expressed Proteins. J.

Am. Chem. Soc. 2015, 137 (3), 1123–1129. https://doi.org/10.1021/ja509955n.

(76) Witus, L. S.; Netirojjanakul, C.; Palla, K. S.; Muehl, E. M.; Weng, C. H.;

173

Iavarone, A. T.; Francis, M. B. Site-Specific Protein Transamination Using N-

Methylpyridinium-4-Carboxaldehyde. J. Am. Chem. Soc. 2013, 135 (45), 17223–

17229. https://doi.org/10.1021/ja408868a.

(77) Advanced BioMatrix. Analysis of Type I Collagen Products; Carlsbad.

(78) Bertelsen, C. M.; Boerio, F. J. Linking Mechanical Properties of Silanes to Their

Chemical Structure: An Analytical Study of γ-GPS Solutions and Films. Prog.

Org. Coatings 2001, 41 (4), 239–246. https://doi.org/10.1016/S0300-

9440(01)00135-7.

(79) Tripp, C. P.; Hair, M. L. Reaction of Chloromethylsilanes with Silica: A Low-

Frequency Infrared Study. Langmuir 1991, 7 (5), 923–927.

https://doi.org/10.1021/la00053a019.

(80) Kanan, S. M.; Tze, W. T. Y.; Tripp, C. P. Method to Double the Surface

Concentration and Control the Orientation of Adsorbed (3-

Aminopropyl)Dimethylethoxysilane on Silica Powders and Glass Slides.

Langmuir 2002, 18 (17), 6623–6627. https://doi.org/10.1021/la0203133.

(81) Acres, R. G.; Ellis, A. V.; Alvino, J.; Lenahan, C. E.; Khodakov, D. A.; Metha,

G. F.; Andersson, G. G. Molecular Structure of 3-Aminopropyltriethoxysilane

Layers Formed on Silanol-Terminated Silicon Surfaces. J. Phys. Chem. C 2012,

116 (10), 6289–6297. https://doi.org/10.1021/jp212056s.

(82) McCafferty, E.; Wightman, J. P. Determination of the Concentration of Surface

Hydroxyl Groups on Metal Oxide Films by a Quantitative XPS Method. Surf.

Interface Anal. 1998, 26 (8), 549–564. https://doi.org/10.1002/(sici)1096-

9918(199807)26:8<549::aid-sia396>3.0.co;2-q.

174

(83) White, L. D.; Tripp, C. P. Reaction of (3-Aminopropyl)Dimethylethoxysilane

with Amine Catalysts on Silica Surfaces. J. Colloid Interface Sci. 2000, 232 (2),

400–407. https://doi.org/10.1006/jcis.2000.7224.

(84) Ho, S. P.; Marshall, S. J.; Ryder, M. I.; Marshall, G. W. The Tooth Attachment

Mechanism Defined by Structure, Chemical Composition and Mechanical

Properties of Collagen Fibers in the Periodontium. Biomaterials 2007, 28 (35),

5238–5245. https://doi.org/10.1016/j.biomaterials.2007.08.031.

(85) Alruwaithi, M.; Jumah, A.; Alsadoon, S. Tooth Ankylosis And Its Orthodontic

Implication. IOSR J. Dent. Med. Sci. 2017, 16 (2), 108–112.

https://doi.org/10.9790/0853-160201108112.

(86) Zeng, X.; Xu, G.; Gao, Y.; An, Y. Surface Wettability of (3-

Aminopropyl)Triethoxysilane Self-Assembled Monolayers. J. Phys. Chem. B

2011, 115 (3), 450–454. https://doi.org/10.1021/jp109259b.

(87) Jain, S. C.; Tanwar, V. K.; Dixit, V.; Verma, S. P.; Samanta, S. B. Surface Order

and Structure Studies of Polymer-Solid Interface. Appl. Surf. Sci. 2001, 182 (3–

4), 350–356. https://doi.org/10.1016/S0169-4332(01)00447-0.

(88) Heiney, P. A.; Grüneberg, K.; Fang, J.; Dulcey, C.; Shashidhar, R. Structure and

Growth of Chromophore-Functionalized (s-Aminopropyl)Triethoxysilane Self-

Assembled on Silicon. Langmuir 2000, 16 (6), 2651–2657.

https://doi.org/10.1021/la990557w.

(89) Haller, I. Covalently Attached Organic Monolayers on Semiconductor Surfaces1.

J. Am. Chem. Soc. 1978, 100 (26), 8050–8055.

https://doi.org/10.1021/ja00494a003.

175

(90) Wei, Z. Q.; Wang, C.; Zhu, C. F.; Zhou, C. Q.; Xu, B.; Bai, C. L. Study on

Single-Bond Interaction between Amino-Terminated Organosilane Self-

Assembled Monolayers by Atomic Force Microscopy. Surf. Sci. 2000, 459 (3),

401–412. https://doi.org/10.1016/S0039-6028(00)00474-X.

(91) Chatterjee, K.; Vogler, E. A.; Siedlecki, C. A. Procoagulant Activity of Surface-

Immobilized Hageman Factor. Biomaterials 2006, 27 (33), 5643–5650.

https://doi.org/10.1016/j.biomaterials.2006.07.025.

(92) Zhu, M.; Lerum, M. Z.; Chen, W. How to Prepare Reproducible, Homogeneous,

and Hydrolytically Stable Aminosilane-Derived Layers on Silica. Langmuir 2012,

28 (1), 416–423. https://doi.org/10.1021/la203638g.

(93) Kowalczyk, D.; Slomkowski, S.; Chehimi, M. M.; Delamar, M. Adsorption of

Aminopropyltriethoxy Silane on Quartz: An XPS and Contact Angle

Measurements Study. Int. J. Adhes. Adhes. 1996, 16 (4), 227–232.

https://doi.org/10.1016/0143-7496(95)00052-6.

(94) Meroni, D.; Lo Presti, L.; Di Liberto, G.; Ceotto, M.; Acres, R. G.; Prince, K. C.;

Bellani, R.; Soliveri, G.; Ardizzone, S. A Close Look at the Structure of the

TiO2-APTES Interface in Hybrid Nanomaterials and Its Degradation Pathway:

An Experimental and Theoretical Study. J. Phys. Chem. C 2017, 121 (1), 430–

440. https://doi.org/10.1021/acs.jpcc.6b10720.

(95) Song, Y. Y.; Hildebrand, H.; Schmuki, P. Photoinduced Release of Active

Proteins from TiO2 Surfaces. Electrochem. commun. 2009, 11 (7), 1429–1433.

https://doi.org/10.1016/j.elecom.2009.05.022.

(96) Song, Y. Y.; Roy, P.; Paramasivam, I.; Schmuki, P. Voltage-Induced Payload

176

Release and Wettability Control on TiO2 and TiO2 Nanotubes. Angew. Chemie -

Int. Ed. 2010, 49 (2), 351–354. https://doi.org/10.1002/anie.200905111.

(97) Lane, C. F. Sodium Cyanoborohydride - A Highly Selective Reducing Agent for

Organic Functional Groups. Synthesis (Germany). Georg Thieme Verlag March

1, 1975, pp 135–146. https://doi.org/10.1055/s-1975-23685.

(98) Golas, A.; Joshua Yeh, C. H.; Siedlecki, C. A.; Vogler, E. A. Amidolytic,

Procoagulant, and Activation-Suppressing Proteins Produced by Contact

Activation of Blood Factor XII in Buffer Solution. Biomaterials 2011, 32 (36),

9747–9757. https://doi.org/10.1016/j.biomaterials.2011.09.020.

(99) Yoshida, T. NIST XPS Database Detail Page Ns1 1,2-Cyclohexanedione

Dioxime. Bull. Chem. Soc. Jpn. 1978, pp 3257–3260.

(100) Diebold, U. The Surface Science of Titanium Dioxide. Surf. Sci. Rep. 2003, 48

(5–8), 53–229. https://doi.org/10.1016/s0167-5729(02)00100-0.

(101) Leikin, S.; Rau, D. C.; Parsegian, V. A. Temperature-Favoured Assembly of

Collagen Is Driven by Hydrophilic Not Hydrophobic Interactions. Nat. Struct.

Biol. 1995, 2 (3), 205–210. https://doi.org/10.1038/nsb0395-205.

(102) Hao, J.; Li, S. K. Mechanistic Study of Electroosmotic Transport across Hydrated

Nail Plates: Effects of PH and Ionic Strength. J. Pharm. Sci. 2008, 97 (12), 5186–

5197. https://doi.org/10.1002/jps.21368.

(103) Uquillas, J. A.; Kishore, V.; Akkus, O. Effects of Phosphate-Buffered Saline

Concentration and Incubation Time on the Mechanical and Structural Properties

of Electrochemically Aligned Collagen Threads. Biomed. Mater. 2011, 6 (3),

035008. https://doi.org/10.1088/1748-6041/6/3/035008.

177

(104) Jelesarov, I.; Dürr, E.; Thomas, R. M.; Bosshard, H. R. Salt Effects on

Hydrophobic Interaction and Charge Screening in the Folding of a Negatively

Charged Peptide to a Coiled Coil (Leucine Zipper). Biochemistry 1998, 37 (20),

7539–7550. https://doi.org/10.1021/bi972977v.

(105) Li, Y.; Douglas, E. P. Effects of Various Salts on Structural Polymorphism of

Reconstituted Type I Collagen Fibrils. Colloids Surfaces B Biointerfaces 2013,

112, 42–50. https://doi.org/10.1016/J.COLSURFB.2013.07.037.

(106) Weinstock, A.; King, P. C.; Wuthier, R. E. The Ion-Binding Characteristics of

Reconstituted Collagen. Biochem. J. 1967, 102 (3), 983–988.

https://doi.org/10.1042/bj1020983.

(107) Kim, S. E.; Wallat, J. D.; Harker, E. C.; Advincula, A. A.; Pokorski, J. K.

Multifunctional and Spatially Controlled Bioconjugation to Melt Coextruded

Nanofibers. Polym. Chem. 2015, 6 (31), 5683–5692.

https://doi.org/10.1039/c5py00282f.

(108) Karpavicius, D.; Stasikelyte, M.; Baseviciene, N.; Sakalauskaite, U.; Ratkute, S.;

Razukevicius, D. The Determination of PH of Peri-Implant Crevicular Fluid

around One-Piece and Two-Piece Dental Implants: A Pilot Study. Clin. Exp.

Dent. Res. 2019, 5 (3), 236–242. https://doi.org/10.1002/cre2.177.

(109) Zhu, J.; Kaufman, L. J. Collagen i Self-Assembly: Revealing the Developing

Structures That Generate Turbidity. Biophys. J. 2014, 106 (8), 1822–1831.

https://doi.org/10.1016/j.bpj.2014.03.011.

(110) Silver, F. H.; Birk, D. E. Kinetic Analysis of Collagen Fibrillogenesis: I. Use of

Turbidity-Time Data. Top. Catal. 1983, 3 (5), 393–405.

178

https://doi.org/10.1016/S0174-173X(83)80020-X.

(111) Comper, W. D.; Veis, A. The Mechanism of Nucleation for in Vitro Collagen

Fibril Formation. Biopolymers 1977, 16 (10), 2113–2131.

https://doi.org/10.1002/bip.1977.360161004.

(112) Zhang, J.; Zou, M.; Zhang, M.; Wei, B.; Xu, C.; Xie, D.; Wang, H. Impact of

Telopeptides on Self-Assembly Properties of Snakehead (Channa Argus) Skin

Collagen. Food Biophys. 2016, 11 (4), 380–387. https://doi.org/10.1007/s11483-

016-9452-5.

(113) Schmittel, M.; Kalsani, V. Functional, Discrete, Nanoscale Supramolecular

Assimblies. Top. Curr. Chem. 2005, 245, 1–53. https://doi.org/10.1007/b98165.

(114) Shekaran, A.; Garcia, A. J. Nanoscale Engineering of Extracellular Matrix-

Mimetic Bioadhesive Surfaces and Implants for Tissue Engineering. Biochimica

et Biophysica Acta - General Subjects. March 2011, pp 350–360.

https://doi.org/10.1016/j.bbagen.2010.04.006.

(115) Rahmany, M. B.; Van Dyke, M. Biomimetic Approaches to Modulate Cellular

Adhesion in Biomaterials: A Review. Acta Biomaterialia. March 2013, pp 5431–

5437. https://doi.org/10.1016/j.actbio.2012.11.019.

(116) Kielty, C. M.; Grant, M. E. The Collagen Family: Structure, Assembly, and

Organization in the Extracellular Matrix. In Connective Tissue and Its Heritable

Disorders; John Wiley & Sons, Inc.: Hoboken, NJ, USA, 2002; pp 159–221.

https://doi.org/10.1002/0471221929.ch2.

(117) Moroi, M.; Jung, S. M.; Shinmyozu, K.; Tomiyama, Y.; Ordinas, A.; Diaz-ricart,

M. Analysis of Platelet Adhesion to a Collagen-Coated Surface Under Flow

179

Conditions: The Involvement of Glycoprotein VI in the Platelet Adhesion. Blood

1996, 88 (6), 2081–2092.

(118) Samokhin, G. P.; Smirnov, M. D.; Muzykantov, V. R.; Domogatsky, S. P.;

Smirnov, V. N. Red Blood Cell Targeting to Collagen-Coated Surfaces. FEBS

Lett. 1983, 154 (2), 257–261. https://doi.org/10.1016/0014-5793(83)80160-4.

(119) Zhao, Y. L.; Liu, W. W.; Liu, W. W.; Lu, Z. Y.; Xuan, D. H.; Zhang, X.; Liu, X.

L.; Hayashi, T.; Yamato, M.; Ogura, T.; et al. Phorbol Ester (PMA)-Treated U937

Cells Cultured on Type I Collagen-Coated Dish Express a Lower Production of

pro-Inflammatory Cytokines through Lowered ROS Levels in Parallel with Cell

Aggregate Formation. Int. Immunopharmacol. 2018, 55 (December 2017), 158–

164. https://doi.org/10.1016/j.intimp.2017.12.013.

(120) Marisca, O.; Kantner, K.; Pfeiffer, C.; Zhang, Q.; Pelaz, B.; Leopold, N.; Parak,

W.; Rejman, J. Comparison of the in Vitro Uptake and Toxicity of Collagen- and

Synthetic Polymer-Coated Gold Nanoparticles. Nanomaterials 2015, 5 (3), 1418–

1430. https://doi.org/10.3390/nano5031418.

(121) Azuma, Y.; Edwardson, T. G. W. W.; Hilvert, D. Tailoring Lumazine Synthase

Assemblies for Bionanotechnology. Chemical Society Reviews. Royal Society of

Chemistry May 21, 2018, pp 3543–3557. https://doi.org/10.1039/c8cs00154e.

(122) Gordon, M. R.; Canakci, M.; Li, L.; Zhuang, J.; Osborne, B.; Thayumanavan, S.

Field Guide to Challenges and Opportunities in Antibody-Drug Conjugates for

Chemists. Bioconjugate Chemistry. American Chemical Society August 26, 2015,

pp 2198–2215. https://doi.org/10.1021/acs.bioconjchem.5b00399.

(123) Macdonald, J. I.; Munch, H. K.; Moore, T.; Francis, M. B. One-Step Site-Specific

180

Modification of Native Proteins with 2-Pyridinecarboxyaldehydes. Nat. Chem.

Biol. 2015, 11 (5), 326–331. https://doi.org/10.1038/nchembio.1792.

(124) Lee, J. P.; Kassianidou, E.; MacDonald, J. I.; Francis, M. B.; Kumar, S. N-

Terminal Specific Conjugation of Extracellular Matrix Proteins to 2-

Pyridinecarboxaldehyde Functionalized Polyacrylamide Hydrogels. Biomaterials

2016, 102, 268–276. https://doi.org/10.1016/j.biomaterials.2016.06.022.

(125) Grandbois, M.; Beyer, M.; Rief, M.; Clausen-Schaumann, H.; Gaub, H. E. How

Strong Is a Covalent Bond. Science (80-. ). 1999, 283 (5408), 1727–1730.

https://doi.org/10.1126/science.283.5408.1727.

(126) Darwent, B. deB. Bond Dissociation Energies in Simple Molecules, “National

Standard Reference Data Series.” National Standard Reference Data Series -

National Bureau of Standards 31: Washington D.C. 1970.

(127) Benson, S. W. Resource Papers - III Bond Energies. J. Chem. Educ. 1965, 42 (9),

502–515. https://doi.org/10.1021/ed042p502.

(128) Stevenson, D. P. The Strengths of Chemical Bonds. J. Am. Chem. Soc. 1955, 77

(8), 2350–2350. https://doi.org/10.1021/ja01613a116.

(129) Schicho, K.; Kastner, J.; Klingesberger, R.; Seemann, R.; Enislidis, G.; Undt, G.;

Wanschitz, F.; Figl, M.; Wagner, A.; Ewers, R. Surface Area Analysis of Dental

Implants Using Micro-Computed Tomography. Clin. Oral Implants Res. 2007, 18

(4), 459–464. https://doi.org/10.1111/j.1600-0501.2007.01338.x.

(130) Wancket, L. M. Animal Models for Evaluation of Bone Implants and Devices:

Comparative Bone Structure and Common Model Uses. Vet. Pathol. 2015, 52 (5),

842–850. https://doi.org/10.1177/0300985815593124.

181

(131) Duan, Y.; Ma, W.; Li, D.; Wang, T.; Liu, B. Enhanced Osseointegration of

Titanium Implants in a Rat Model of Osteoporosis Using Multilayer Bone

Mesenchymal Stem Cell Sheets. Exp. Ther. Med. 2017, 14 (6), 5717–5726.

https://doi.org/10.3892/etm.2017.5303.

(132) Ogiso, B.; Hughes, F. J.; Melcher, A. H.; McCulloch, C. A. G. Fibroblasts Inhibit

Mineralised Bone Nodule Formation by Rat Bone Marrow Stromal Cells in Vitro.

J. Cell. Physiol. 1991, 146 (3), 442–450. https://doi.org/10.1002/jcp.1041460315.

(133) Hanhan, S.; Ejzenberg, A.; Goren, K.; Saba, F.; Suki, Y.; Sharon, S.; Shilo, D.;

Waxman, J.; Spitzer, E.; Shahar, R.; et al. Skeletal Ligament Healing Using the

Recombinant Human Amelogenin Protein. J. Cell. Mol. Med. 2016, 20 (5), 815–

824. https://doi.org/10.1111/jcmm.12762.

182