Open Collections

UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Cellulose-based biosensors of human neutrophil elastase (HNE) toward chronic wound point-of-care diagnostics Saisuwan, Ravi 2020

Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.

Notice for Google Chrome users:
If you are having trouble viewing or searching the PDF with Google Chrome, please download it here instead.

Item Metadata

Download

Media
24-ubc_2020_may_saisuwan_ravi.pdf [ 7.32MB ]
Metadata
JSON: 24-1.0389685.json
JSON-LD: 24-1.0389685-ld.json
RDF/XML (Pretty): 24-1.0389685-rdf.xml
RDF/JSON: 24-1.0389685-rdf.json
Turtle: 24-1.0389685-turtle.txt
N-Triples: 24-1.0389685-rdf-ntriples.txt
Original Record: 24-1.0389685-source.json
Full Text
24-1.0389685-fulltext.txt
Citation
24-1.0389685.ris

Full Text

CELLULOSE-BASED BIOSENSORS OF HUMAN NEUTROPHIL ELASTASE (HNE) TOWARD CHRONIC WOUND POINT-OF-CARE DIAGNOSTICS by  Ravi Saisuwan  B.Eng., Thammasat University, 2010 M. Eng., University of Tokyo, 2012  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Chemistry)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  March 2020  © Ravi Saisuwan, 2020 ii  The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, a thesis/dissertation entitled:  Cellulose-based biosensors of human neutrophil elastase (HNE) toward chronic wound point-of-care diagnostics  submitted by Ravi Saisuwan in partial fulfillment of the requirements for the degree of Master of Science in Chemistry  Examining Committee: Prof. Harry Brumer, Chemistry and Michael Smith Laboratory Supervisor  Prof. Lawrence P. McIntosh, Chemistry Supervisory Committee Member  Prof. Mark MacLachlan, Chemistry Supervisory Committee Member  Additional Examiner  iii  Abstract Chronic wounds, which fail to heal or heal only very slowly, remain a major challenge in medical treatment and a burden to healthcare systems. Chronic wounds are exacerbated by bacterial infection and endogenous proteases.  In particular, increased levels of human neutrophil elastase (HNE) have been observed in chronic wound fluid, and has been utilized as a severity indicator via clinical assays. To facilitate chronic wound treatment, a facile, in situ, detection method for HNE would be advantageous. Here, cellulose-based analytical devices have been designed and produced as a proof-of-concept for chronic wound point-of-care diagnostics.  Specifically, two distinct fluorogenic HNE substrates amenable to click-chemistry were synthesized based on tetrapeptide (Ala4) conjugates to the rhodamine derivatives, carboxyrhodamine110-PEG3-azide (cRho110-PEG3-N3) and rhodamine 110 (Rho110).  Michaelis-Menten kinetics were used to demonstrate activity of HNE toward these compounds that was comparable to known chromogenic substrates.  Following attachment under mild aqueous conditions to alkyne-functionalized Whatman No. 1 filter paper, a pure cellulosic substrate and model for cotton gauze, HNE detection on a solid surface was demonstrated visually under specific illumination and was quantified with a fluorescence scanner.  These results validate the concept of in situ protease detection using modified cellulose surfaces to monitor chronic wounds toward improved treatment outcomes.  Furthermore, the modular design of the cellulose-based analytical devices presented here suggests a broader potential for the detection of specific protease activity in diverse applications. iv  Lay Summary Chronic wounds of the skin, which take months to years to heal (if ever), are a major problem in healthcare.  This research focuses on the fabrication of an analytical device to a protein-degrading enzyme, the protease human neutrophil elastase (HNE), which is over-produced in chronic wounds, degrades proteins in the skin, and delays healing. I have designed and synthesized two compounds with specially-designed chemical features to interact with HNE and produce a color change upon detection. These compounds also include a chemical group that enables attachment to wound dressings such as cotton gauze. The resulting dressing can then be used to monitor HNE levels, and therefore chronic wound status, directly at the point-of-care. v  Preface  This work presented in this thesis is an original and unpublished work by the author R. Saisuwan, which was supervised by Dr. Harry Brumer. All the experiments were performed by the author with technical assistance from post-doctoral scientists Dr. Changqing Wang (chemical synthesis) and Dr. Laleh Solhi (cellulose surface modification and enzyme detection) in the Brumer group. vi  Table of Contents  Abstract ......................................................................................................................................... iii Lay Summary ............................................................................................................................... iv Preface .............................................................................................................................................v Table of Contents ......................................................................................................................... vi List of Tables ..................................................................................................................................x List of Figures ............................................................................................................................... xi List of Schemes ........................................................................................................................... xiv List of Symbols and Abbreviations .......................................................................................... xvi Acknowledgements .................................................................................................................. xviii Dedication ................................................................................................................................... xix Chapter 1: Introduction ................................................................................................................1 1.1 Chronic wounds .............................................................................................................. 1 1.1.1 Anatomy of skin .......................................................................................................... 1 1.1.2 Ordinary wound healing mechanism .......................................................................... 2 1.1.3 Physiology of chronic wounds .................................................................................... 3 1.2 Proteases ......................................................................................................................... 4 1.2.1 Human Neutrophil Elastase (HNE) ............................................................................ 4 1.2.2 HNE Specificity .......................................................................................................... 6 1.2.3 HNE Fluorogenic/Chromogenic Substrates ................................................................ 7 1.3 Biosensors ....................................................................................................................... 9 1.3.1 Cellulose for Biomedical Application ........................................................................ 9 vii  1.3.2 Cellulose-based Biosensors for HNE ........................................................................ 10 1.3.3 A Cellulose-based Esterase Biosensor from Brumer Group..................................... 11 1.4 Thesis Objective - Design of an HNE-responsive cellulosic biosensor ....................... 13 1.4.1 Design of CuAAC Clickable HNE Fluorogenic Substrates ..................................... 16 1.4.2 Cellulose Surface Activation .................................................................................... 17 1.4.3 Copper(I)-catalyzed Azide-Alkyne Cycloaddition (CuAAC) .................................. 19 Chapter 2: Synthesis of HNE Artificial Substrates ..................................................................22 2.1 Background and synopsis ............................................................................................. 22 2.1.1 Proposed Synthesis of Bis-tetraalanine HNE Substrate (5) ...................................... 22 2.1.1.1 Amide Bond Formation (Peptide Coupling) ..................................................... 24 2.1.1.2 Boc as a Peptide Protecting Group ................................................................... 26 2.1.2 Proposed Synthesis of Mono-tetraalanine HNE Substrate (13) ................................ 28 2.2 Results and Discussions ................................................................................................ 30 2.2.1 Synthesis of the Bis-tetraalanine HNE substrate (5) ................................................. 30 2.2.1.1 Coupling Reaction with HATU and DIPEA ..................................................... 31 2.2.1.2 Boc Deprotection .............................................................................................. 31 2.2.1.3 Structural Validation of Compound 5 and Synthetic Intermediates by NMR .. 32 2.2.2 Synthesis of Mono-tetraalanine HNE Substrate (13) ................................................ 38 2.2.2.1 Selective Boc Protection of Rho110 ................................................................. 38 2.2.2.2 Coupling of the Linker with Compound 7 ........................................................ 39 2.2.2.3 Boc- Deprotection ............................................................................................. 39 2.2.2.4 Structural Validation of Compound 13 and Synthetic Intermediates by NMR 40 2.3 Summary ....................................................................................................................... 46 viii  2.4 Experimental Section .................................................................................................... 47 2.4.1 Materials ................................................................................................................... 47 2.4.2 Thin Layer Chromatography (TLC) ......................................................................... 47 2.4.3 1D NMR and 2D NMR ............................................................................................. 47 2.4.4 High-Resolution ESI MS .......................................................................................... 48 2.4.5 Synthesis Procedure of Compound 5 ........................................................................ 48 2.4.6 Synthesis Procedure of Compound 13 ...................................................................... 52 Chapter 3: Application of Fluorogenic Substrates for Human Neutrophil Elastase ............59 3.1 Background and Synopsis ............................................................................................. 59 3.1.1 Enzyme Kinetics ....................................................................................................... 59 3.1.2 Kinetic Model of Human Neutrophil Elastase .......................................................... 62 3.1.3 Spectrophotometry .................................................................................................... 63 3.2 Materials and Methods .................................................................................................. 65 3.2.1 Solution-phase Enzyme Kinetics .............................................................................. 65 3.2.1.1 Determination of Extinction Coefficients ......................................................... 65 3.2.1.2 Preliminary Verification of the Activity of HNE on Compound 5 ................... 66 3.2.1.3 Michaelis Menten Kinetics of HNE on Compound 5 and Compound 13 ........ 66 3.2.2 Activation of Cellulose Paper Surfaces and Attachment of Fluorophores ............... 67 3.2.3 CuAAC Optimization using cRho110-PEG3-N3 and Compound 9 .......................... 68 3.2.4 Calibration Curve by Absorption of cRho110-PEG3-N3 and Compound 9 .............. 69 3.2.5 Fluorescence Imaging ............................................................................................... 69 3.2.6 Analysis of HNE Activity on Modified Cellulose Surfaces ..................................... 69 3.3 Results and Discussions ................................................................................................ 70 ix  3.3.1 Validation of Compounds 5 and 13 as Substrates for Human Neutrophil Elastase .. 70 3.3.2 Production and Analysis of Paper-based Biosensors for HNE ................................. 74 3.3.2.1 Bis-tetraalanine-based HNE Biosensor ............................................................. 75 3.3.2.2 Mono-tetraalanine-based HNE Biosensor ........................................................ 79 3.4 Summary ....................................................................................................................... 83 Chapter 4: Conclusion and Future Works ................................................................................84 4.1 Conclusion .................................................................................................................... 84 4.2 Future Works ................................................................................................................ 85 4.2.1 Synthesis of HNE Substrate Library by SPOT Synthesis ......................................... 85 4.2.2 Cellulosic Surface Chemistry Exploration and Development .................................. 87 4.2.3 Fabrication an HNE Dual-Mode Bio-Responsive Surface ....................................... 88 Bibliography .................................................................................................................................92 Appendices ..................................................................................................................................101 Appendix A Selected NMR Spectrum .................................................................................... 101        x  List of Tables   Table 1.1 List of commercially available HNE artificial substrates with chemical information (structure, molecular weight, and spectral properties) .................................................................... 8 Table 3.1 Michaelis-Menten kinetic parameters of HNE substrates ............................................ 73 Table 3.2 CuAAC reactivity of free cRho110-PEG3-N3 on propargyl amine coupled filter paper....................................................................................................................................................... 75 Table 3.3 CuAAC reactivity of compound 9 on propargyl amine coupled filter paper ............... 80   xi  List of Figures  Figure 1.1 Skin layers: epidermis, dermis, and hypodermis ........................................................... 2 Figure 1.2 Molecular pathology of chronic wounds. Chronic wound shows hyperproliferative and nonmigratory epidermis, unresolved inflammation, presence of infection, and biofilm formation ......................................................................................................................................... 3 Figure 1.3 Mechanisms of HNE on cleavage of peptide bonds ...................................................... 5 Figure 1.4 Relative activity of each subsite of HNE toward natural amino acids .......................... 6 Figure 1.5 Chemical structures of natural and synthetic amino acids specific to each subsite of HNE ................................................................................................................................................ 7 Figure 1.6 Molecular structure of HNE artificial substrate functionalized onto cellulosic supports....................................................................................................................................................... 11 Figure 1.7 Cellulose-based enzyme sensor approaches (A) Surface linking with biomolecules, (B) Surface linking with fluorogenic moiety. ............................................................................... 12 Figure 1.8 Molecular structure of fluorogenic substrates of esterase artificial substrate having 5(6)-carboxyfluorescein-tetraethylene glycol (TEG)-azide (FTA) as a chromogenic core which offers two hydroxyl group for esterification. ................................................................................ 13 Figure 1.9 Schematic illustration of HNE bioactive cellulosic surface ........................................ 14 Figure 1.10 Schematic illustration of cellulose-based elastase biosensor by conjugation between azide-terminated fluorogenic substrates and alkyne-functionalized cellulosic surface via CuAAC click chemistry .............................................................................................................................. 15 Figure 1.11 Molecular design of (A) divalent fluorophores, and (B) monovalent fluorophores .. 16 xii  Figure 1.12 (red) TEMPO oxidized cellulosic surface, (blue) PDITC activated cellulose surface, and (green) DVS activated cellulose surface ................................................................................ 18 Figure 1.13 TEMPO oxidation mechanism on hexopyranose ...................................................... 19 Figure 1.14 Schematic illustration of copper(I)-catalyzed azide-alkyne cycloaddition reaction . 20 Figure 1.15 Mechanism of copper(I)-catalyzed azide-alkyne cycloaddition forming 1,4 disubstituted [1,2,3] triazoles ........................................................................................................ 20 Figure 2.1 Structure and position numbering of compound 5 and its intermediates .................... 33 Figure 2.2 1H-NMR spectrum of 5 in MeOD-d4........................................................................... 33 Figure 2.3 13C-NMR spectrum of 5 in MeOD-d4. ........................................................................ 34 Figure 2.4 2D-COSY spectrum of 5 in MeOD-d4. ....................................................................... 34 Figure 2.5 2D-HSQC spectrum of 5 in MeOD-d4. ....................................................................... 35 Figure 2.6 2D-HMBC spectrum of 5 in MeOD-d4. ...................................................................... 35 Figure 2.7 Close-up 1H-NMR spectrum of cRho110-PEG3-N3, compound 3 and compound 5 in MeOD-d4 ....................................................................................................................................... 37 Figure 2.8 Structure and position numbering of compound 13 and its intermediates .................. 40 Figure 2.9 1H-NMR spectrum of 13 in MeOD-d4......................................................................... 41 Figure 2.10 13C-NMR spectrum of 13 in MeOD-d4. .................................................................... 41 Figure 2.11 2D-COSY spectrum of 13 in MeOD-d4. ................................................................... 42 Figure 2.12 2D-HSQC spectrum of 13 in MeOD-d4. ................................................................... 42 Figure 2.13 2D-HMBC spectrum of 13 in MeOD-d4. .................................................................. 43 Figure 2.14 Close-up 1H-NMR spectrum of Rho110, compound 9, 11 and 13 in MeOD-d4 ....... 45 xiii  Figure 3.1 (Solid line) Product(P) progress curves of enzymatic reactions with various substrate (S) concentration, (Dash line) Tangent line, representing the reaction velocities, of progress curves at initial substrate concentrations ...................................................................................... 61 Figure 3.2 Michaelis Menten plot between initial velocity (V0) and substrate concentration ([S])....................................................................................................................................................... 62 Figure 3.3 (A) Ping-Pong Bi-Bi enzymatic mechanism of proteolytic reaction by HNE, (B) Uni Bi enzymatic mechanism .............................................................................................................. 63 Figure 3.4 Jablonski diagram-Energy diagram showing fluorescence and phosphorescence ...... 64 Figure 3.5 Graph illustrating Strokes’ law and Strokes’ shift ...................................................... 65 Figure 3.6 Control solution and the HNE treated compound 5 solution under ambient and UV light with the wavelength of 254 nm. ........................................................................................... 70 Figure 3.7 Extinction coefficients of fluorogenic substrates and their fluorescent precursors in dH2O ............................................................................................................................................. 71 Figure 3.8 Michaelis-Menten curve of compound 5 in PBS pH 7.6 at 37°C, in HEPES pH 7.4 at 37°C and compound 13 in HEPES pH 7.4 at 37°C toward HNE ................................................. 72 Figure 3.9 HNE’s catalytic triad (His57, Asp102, and Ser195) covalently inhibited by N-[4-[(4-morpholinyl)carbonyl]benzoyl]peptidyl pentafluoroethyl ketone ................................................ 74 Figure 3.10 cRho110-PEG3-N3 clicked onto propargyl amine coupled paper discs with various reacted cRho110-PEG3-N3 quantities ........................................................................................... 76 Figure 3.11 (A) cRho110-PEG3-N3 absorbed Whatman no.1 filter paper without PDMS coating, (B) with PDMS coating, (C & D) Different amount of  cRho110-PEG3-N3 absorbed paper discs visualized under (C) a laboratory epi-illuminator with orange filter, (D) Bio-rad ChemiDoc XRS with Alexa Fluor 488 setting......................................................................................................... 77 xiv  Figure 3.12 (A) Calibration curve of cRho110-PEG3-N3 absorption method, (B) Close-up calibration curve from cRho110-PEG3-N3 absorption method in the range of 0 to 0.65 nmol giving a straight line before saturation and quenching effect occur. ............................................ 77 Figure 3.13 Enzyme concentration and time-dependent enzymatic hydrolysis of compound 5 on paper surfaces................................................................................................................................ 78 Figure 3.14 (A) Calibration curves of compound-9 absorption method, (B) Close-up calibration curve from compound-9 absorption method in the range of 0 to 2.9 nmol giving a straight line before saturation............................................................................................................................ 80 Figure 3.15 Enzyme concentration and time-dependent enzymatic hydrolysis of compound 13 on paper surfaces................................................................................................................................ 82 Figure 4.1 (A) Molecular structure of free cRho110-PEG3-N3 functionalized as a 7-mm spot on a paper, (B) SPOT synthesis paper template that can vary 2 peptide variables at a time ................ 86 Figure 4.2 (A) Original divalent substrate and (B)modified divalent fluorogenic substrate with amine-terminated linker and succinylated tetra-alanine chains as HNE recognition modules ..... 87 Figure 4.3 (red) Amide bond coupling of HNE substrate directly toward TEMPO oxidized paper; (blue) Thiourea formation of HNE substrate using PDITC as a surface activator, and (green) Secondary amine formation of HNE substrate using DVS as a surface activator ........................ 88 Figure 4.4 Schematic illustration of dual-mode bio-responsive cellulosic surface ...................... 90   xv  List of Schemes  Scheme 2.1 Retrosynthesis analysis of HNE divalent fluorogenic substrate 5, PG = protecting group ............................................................................................................................................. 23 Scheme 2.2 Synthesis scheme of divalent fluorophore 5 ............................................................. 24 Scheme 2.3 Mechanism of amide bond formation by HATU ...................................................... 25 Scheme 2.4 Mechanism of amide bond formation by DIC .......................................................... 26 Scheme 2.5 N-terminal Boc protection ......................................................................................... 27 Scheme 2.6 Boc deprotection mechanism in trifluoroacetic acid (TFA)...................................... 28 Scheme 2.7 Retrosynthesis analysis of monovalent fluorogenic substrate 13 .............................. 29 Scheme 2.8 Synthesis scheme of monovalent fluorophore 13 ..................................................... 30 Scheme 2.9 Synthesis of Boc-tri-L-alanine-COOH ...................................................................... 50 Scheme 2.10 General synthesis of Linker N3-PEG3-CH2COOH ............................................... 53    xvi  List of Symbols and Abbreviations  %wt Weight percent °C  Degree celcius 1D One-dimensional 2D Two-dimensional a.u. Arbitrary unit cRho110-PEG3-N3 Carboxyrhodamine 110-PEG3-Azide AFC 7-amino-4 trifluoromethylcoumarin AMC  7-amino-4-methylcoumarin Boc Tert-Butyloxycarbonyl Boc2O Di-tert-butyl dicarbonate COSY correlation spectroscopy CuAAC Copper(I)-catalyzed azide-alkyne cycloaddition DCM Dichloromethane DEPT Distortionless Enhancement by Polarization Transfer  DI Deionized DIC Diisopropyl carbodiimide DIPEA N,N- diisopropyl-N-ethylamine  DMAP 4-Dimethylaminopyridine DMF Dimethylformamide DMSO Dimethylsulfoxide DVS Divinyl sulfone EC Enzyme Commission Number ECM Extracellular Matrix EDC 1-Ethyl-3-(3-dimethylaminopropyl)carbodiimide EtOAc Ethyl acetate Fmoc Fluorenylmethyloxycarbonyl g Gram(s) h Hour(s) HATU 2-(1H-7-azabenzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate methanaminium  HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid  HMBC Heteronuclear Multiple-Bond Coherence  HNE Human Neutrophil Elastase HOAt 1-Hydroxy-7-azabenzotriazole HOBt Hydroxybenzotriazole HRESI-MS High-resolution electron spray ionization mass spectroscopy xvii  HSQC Heteronuclear Single Quantum Coherence  kDa Kilodalton(s) M Molarity MeO Methoxy MeOD Deuterated methanol MeOH Methanol MeO-Suc- Methoxy-succinylated MES 2-(N-morpholino)ethanesulfonic acid mg Milligram(s) MHz Megahertz min minute(s) ml Millilitre(s) MM Michaelis-Menten mmol Millimol(s) N Normality NHS N-Hydroxysuccinimide nm nanometer(s) NMR Nuclear magnetic resonance PBS Phosphate buffer solution PDF Postdoctoral fellow PDITC 1,4-phenylenediisothiocyanate PDMS Polydimethylsiloxane pmol picomol pNA Para-nitroaniline ppm Parts per million Rho110 Rhodamine 110 rpm Revolutions per minute RT Room temperature Suc Succinylated TEMPO (2,2,6,6-Tetramethylpiperidin-1-yl)oxyl TFA Trifluoroacetic acid THF Tetrahydrofuran THPTA tris-hydroxypropyltriazolylmethylamine TLC Thin-layer chromatography U Unit UV Ultraviolet v/v Volume by volume Vis Visible light  xviii  Acknowledgements  To accomplish this project, I would like to express my appreciation to these people and organizations for helpful support and advice:   My supervisor, Prof. Harry Brumer for this opportunity, support and guidance along the way of this project.  UBC NMR facility staff, UBC MS laboratory staff, UBC shared instrument facility staff  All my friends: Dr. Julie Grondin, Dr. Changqing Wan, Dr. Laleh Solhi, Dr. Stephanie M. Forget, Dr. Gregory Arnal, Dr. Guillaume Dejean, Dr. Yann Mathieu, Dr. Jonathon Briggs, Dr. Walid Abdelmagid, Ms. Hila Behar, Mr. James Li, Ms. Maria Cleveland, Mr. Sean Patrick McDonald, Mr. Kazune Tamura, Ms. Namrata Jain, Mr. Fan (Roderick) Xia, Ms. Joyce Li, Ms. Ariel Zhang, Ms. Sina Barghahn, Mr. Konrad Subieta, Ms. Thana Juckmeta, Ms. Boonyanoot Chaiyosang, Dr. Nattinee Bumbudsanpharoke, Mr. Hyuk Joon Jung for help proofreading my thesis, and to comfort my journey in UBC.  NSERC and Canfor Pulp for funding this project  My family – Mr. Wanich and Mrs. Rujirawarn Saisuwan, Ms. Varitthipa, Ms. Patteera, and Ms. Valaisakorn Disyawanawat - No words can explain this warm feeling.  Mr. Boris Sin – for proofreading both in-line and between-the-lines  Myself – I appreciate who I am, who I was, who I shouldn’t have been, and who I might become.  xix  Dedication          From the unstoppably expanding universe ‘till the smallest dust in the ocean, I hope to find a peace in my mind somewhere between.1  Chapter 1: Introduction 1.1 Chronic Wounds 1.1.1 Anatomy of Skin Accounting for one sixth of the total body weight and utilizing one third of the resting cardiac output, the integument, or skin, is considered to be the largest organ system in human body.1 Spanning from the outermost surface to the inner tissues, the skin consists of distinct connective tissues, which combine to serve multiple functions, akin to an in vivo biocomposite. The three main layers of the skin, having different thicknesses and compositions, are shown in Figure 1.1.   The epidermis is defined as a densely-packed layer with cells forming protective sheets of cells (‘epithelia’). Under the epidermis lies a thicker layer, the dermis, where blood vessels, hair follicles and sweat glands are located. Capable of storing the highest amount of fat in the entire integumentary system, the adipose tissue acts as a supportive structure underneath the epidermis and dermis that insulates the body.2 Through this complex design, the human body is shielded against multiple biotic and abiotic assaults. At the same time, the skin is structurally vulnerable, in the sense that minor injuries such as cuts and grazes can introduce defects leading to detrimental effects to the body.       2   Figure 1.1 Layers of skin: epidermis, dermis, and hypodermis. Reprinted with permission from Springer Nature (3).  1.1.2 Ordinary Wound Healing Mechanism Following damage, the wound healing process occurs in four overlapping phases to restore  the skin to its original condition.1,2  First, clotting of platelets and red blood cells (coagulation) stops bleeding and, further, prevents pathogens from entering the body through the wound. Second, an inflammation phase results from the clearance of dead cells, pathogens, and debris by leukocytes of the immune system.  Following the release of platelet-derived growth factors, the third phase, proliferation results in cell migration and division, leading to wound closure and construction of new tissue, including blood vessels and epithelium. To complete the healing process, at the end of proliferation, extracellular matrixproteins are subject to rearrangement in order to regain full epithelial integrity. When a wound fails to follow this normal progression, depending on severity, it may persist as a chronic wound.1,2   3  1.1.3 Physiology of Chronic Wounds Chronic wounds exhibit a prolonged inflammatory phase,4 in which bacterial growth can lead to a persistant biofilm in the wounded area (Figure 1.2). The proliferation phase is also inhibited, due to dermal and epidermal cells becoming unresponsive toward stimuli, thus leaving the defect exposed to further pathogenic contamination.5 Consequently, protease secretion by leukocytes becomes highly upregulated to target these pathogens, with the detrimental side-effect of degrading the extracellular matrix (ECM) proteins such as fibronectin and tenascin. With the ECM weakened, wound closure and remodeling is challenged.  As such, control of protease activity is vital to addressing chronic wounds.4,5   Figure 1.2 Molecular pathology of chronic wounds. (A) Chronic wounds show hyperproliferative and nonmigratory epidermis, unresolved inflammation, presence of infection, and biofilm formation. Reprinted with permission from Elsevier (6).  4  1.2 Proteases Proteases (proteinases) catalyze the hydrolysis of amide bonds in polypeptide. The diverse proteases serve a multitude of biological roles in the body, including in protein maturation during biosynthesis, e.g. the generation of active enzymes and hormones from precursors, intracellular protein recycling, and protein recycling following cell death or during tissue remodeling (e.g., in wound repair).7 Aberrant protease activity, on the other hand, is associated with tumor invasion, rheumatoid arthritis, Alzheimer’s disease, and chronic wounds, among other diseases. Proteases can be broadly classified according to catalytic mechanism, into serine proteases, aspartyl proteases, and metalloproteases.7,8  In the context of chronic wounds, human neutrophil elastase (HNE) is a serine protease that is highly overproduced, leading to sustained tissue damage.    1.2.1 Human Neutrophil Elastase (HNE) HNE (EC 3.4.21.37) is a single-chain, glycosylated serine protease comprised of 218 amino acids and an approximate molecular weight of 30 kDa. Utilizing a canonical serine hydrolase mechanism (Figure 1.3), the catalytic triad of HNE is comprised of Ser195, Asp102, and His57.9  As it is a human enzyme, HNE has a temperature optimum of 37 °C and is active over the pH range 5.5 – 8. HNE is found within the azurophilic granules of polymorphonuclear leukocytes, and is responsible for matrix remodeling in the final step of skin wound healing.9 However, significant increase in HNE level is be observed in chronic wounds, and as such, HNE is utilized as an inflammatory biomarker and indication of chronic wound severity. 10 5   Figure 1.3 Mechanisms of HNE on cleavage of peptide bonds. Reprinted with permission from Taylor & Francis (9).        6  1.2.2 HNE Specificity  Detailed studies have elucidated the amino acid sidechain specificity of HNE, which is particularly relevant for the design of artificial substrates to monitor activity.  Hermiston et al. and Drag et al. tested a broad range of tetrapeptides to determine the amino acid preference of each of the four subsites, P1 to P4, in the catalytic pocket of HNE (Figure 1.4).11,12   In general, P1 prefers valine, alanine, threonine, and isoleucine; proline is preferred for P2;11 P3 prefers glutamine, glutamic acid, and methionine;11 and P4 prefers norleucine.11    Figure 1.4 Relative activity of each subsite of HNE toward natural amino acids. Reprinted with permission from O'Donoghue (11).   In order to gain a more detailed understanding of HNE subsite specificity, Drag and co-workers employed a wide range of natural and non-natural amino acids.12  Similar to previous results (Figure 1.5), these authors determined that P1 strongly prefers small aliphatic residues. However, compared to alanine, the incorporation of 2-aminobutanoic acid (Abu) at this position can boost  catalytic efficiency by 500%. Similarly, when valine is replaced with Norvaline (Nva), an increase in catalytic efficiency is observed. Substitution of P2 with Octahydro-1H-indole-2-carboxylic acid (Oic), 6-benzyloxy-L-norleucine [Nle(O-Bzl)], Nε-(2-chloro-Z)-L-lysine [Lys(2-Cl-Z)], L-3,4-difluorophenylalanine [Phe(3,4-F)], 3,4-dehydroproline (dhPro), and homocitrulline (hCit), further indicated that P2 is highly specific to bulky amino acids. On the other hand, a five-fold increase in activity was observed by replacing glutamine with methionine dioxide [Met(O)2] 7  in P3. P4 displays a preference for non-natural hydrophobic amino acids with bulky side chains, such as 4-benzoylphenylalanine (Bpa), Oic, and cyclohexylalanine (Cha).12 Altogether, these detailed analyses have provided crucial fundamental information HNE specificity, which significantly enables the development of artificial substrates as HNE probes.   Figure 1.5 Chemical structures of natural and synthetic amino acids specific to each subsite of HNE  1.2.3 HNE Fluorogenic/Chromogenic Substrates Building upon the established peptide specificity of HNE, several artificial chromogenic and fluorogenic substrates based on tetrapeptides have been developed (Table 1.2). Hydrolysis by HNE of the simplest of these, Glp-PV-pNA, results in cleavage of the terminal amide bond to release the pH-sensitive chromophore para-nitroaniline, enabling visual or spectrometric observation.10 Likewise, hydrolysis of the terminal amide in MeOSuc-AAPV-AFC releases the fluorophore 7-amino-4 trifluoromethylcoumarin (AFC).10 In contrast, FAM-AAPV-Dabcyl and EDANS-AAPV-Dabcyl utilize Förster resonance energy transfer (FRET) as a principle for spectroscopic analysis. The cleavage of these two 8  peptides by HNE enables independent diffusion of the fluorophores and a resulting shift in the fluorescence spectrum.  Together, the substrates shown in Table 1.1 provide a convenient means to monitor HNE activity in solution-based assays.  Table 1.1 Commercially available artificial substrates for HNE. Reprinted with permission from Springer Nature (10).   9  1.3 Biosensors Classically, biosensors electrochemically detects and quantifies a biomolecules, for instance, protein, carbohydrate and lipid.13,14  Biosensors are composed of a probe that specifically interacts with an analyte to produce a signal such as electrons, ions, or photons, and a signal transducer  One of the most well-known biosensors is the blood glucose meter used in diabetes monitoring. Glucose molecules in a drop of blood are oxidized by glucose oxidase, ultimately generating current via a redox mediator (ferricyanide) ,which can be quantified. 13  A key concern in the development of commercial biosenors for consumer use is the cost of manufacture and operation versus performance.  In an effort to reduce costs, including in the case of blood glucose monitoring, disposable strip-based detection systems on paper or modified plastic substrates has a long history of development.  1.3.1 Cellulose for Biomedical Application As a structural material, cellulose - in the form of gauze, cotton, and non-woven fabrics - has been used widely in medical applications, including wound dressings, medical implants, drug delivery, vascular grafts, and scaffolds for tissue engineering.15–20 Along with these conventional applications, due to its biocompatibility, excellent filtering properties and high cost-effectiveness, cellulose-based materials have been studied for the purpose of constructing low-cost biosensors in recent years.21,22  The numerous examples include two- and three-dimensional paper-based analytical devices (PADs), often incorporating flow via capillary action, to detect proteins, nucleic acids, and other analytes. 23–29  10  1.3.2 Cellulose-based biosensors for HNE Building upon their earlier work using polymer substrates,30 Edwards and co-workers  were the first group to develop a cellulose-based HNE biosensor, through the covalent attachment of commercially available substrates suc–AAPV–AMC, suc–APA–AMC, and suc-AAPA-pNA (Table 1.1, cf. Section 1.2.3, above) to cotton fibers, microfibrils, and nanofibrills. 24–26,31   The ability of the resulting biosensors to detect HNE has been compellingly demonstrated with both AMC and pNA leaving groups, with the fluorophore AMC having significantly greater sensitivitiy than pNA, which required the addition of a color-amplifying reagent.31,32   In the context of direct HNE sensing in chronic wounds, the biosensors devised by Edwards and colleagues have two potential drawbacks related to hydrolysis of AMC and pNA from the cellulosic surface: First, continuous liberation of the chromophore/fluorophore from the biosensor31 will result in a loss of signal over time. Second, there is significant concern about the potential toxicity of the released molecule to the wound tissue. These issues suggest that alternate strategies may be of value to advance the application cellulose-based biosensors for enzymes such as HNE. 11     Figure 1.6 Molecular structure of HNE artificial substrate functionalized onto cellulosic supports. Reprinted with permission from Elsevier (33).  1.3.3 A cellulose-based esterase biosensor from Brumer Group Inspired by the seminal work of Edwards and co-workers, the Brumer group recently advanced an alternate strategy for cellulose-based enzyme biosensors that overcomes issues related to fluorophore diffusion after substrate cleavage (Figure 1.6).28 In this approach, the position of the fluorophore and peptide are switched with respect to the cellulose-linking moiety (Figure 1.7B).  As such, the fluorophore remains attached to the cellulose surface, while the (non-chromophoric) biomolecular recognition element (peptide, lipid, or carbohydrate) is allowed to diffuse away to be ultimately metabolized.  12   Figure 1.7 Cellulose-based enzyme sensor approaches (A) Surface linking with biomolecules, (B) Surface linking with fluorogenic moiety. Reprinted with permission from American Chemical Society (28).  To prove this concept, an esterase was chosen as an exemplar enzyme analyte because of the comparative simplicity of the synthesis of the corresponding fluorescein di-acyl ester.28 The carboxyl group of fluorescein moiety further provided a convenient point of attachment for an azide-terminated tetraethylene glycol linker for copper(I)-catalyzed azide-alkyne cycloaddition (CuAAC) click chemistry. Alkyne-terminated xyloglucan was physically adsorbed onto a cotton fiber-based paper (Whatman No. 1) to provide a complementary clickable cellulosic surface ( Figure 1.8). The applicability of this biosensor to esterase detection was validated by measuring the time- and enzyme-dependent increase in fluorescence using epi-illumination with a forensic flashlight for visual analysis and a fluorescence scanner for quantitation.28 We readily envisioned that this fabrication approach could be further adapted to engineer paper-based biosensors specific for other potential enzymes, including proteases such as HNE, which forms the basis of this thesis work. 13   Figure 1.8 Molecular structure of fluorogenic substrates of esterase artificial substrate having 5(6)-carboxyfluorescein-tetraethylene glycol (TEG)-azide (FTA) as a chromogenic core (R = propyl). Reprinted with permission from  the American Chemical Society (28).   1.4 Thesis Objective - Design of an HNE-responsive cellulosic biosensor Building on the seminal work of Edwards and our own work on cellulosic enzyme biosensors, the overarching goal of this thesis is to develop an improved point-of-care diagnostic tool to effectively detect secreted human neutrophil elastase in chronic wound fluids to facilitate better wound management (Figure 1.9). To design the optimal route for the production of a cellulose-based biosensor for HNE, both the synthesis of the fluorogenic substrate and the method of cellulose surface activation were considered (Figure 1.10). Firstly, two new fluorogenic HNE substrates amenable to cellulose-surface attachment were designed, synthesized, and validated by kinetics measurements in the solution phase. Second, the method of attachment of the artificial substrates onto cellulosic surfaces, ultimately using TEMPO oxidation, amidation, and copper(I) alkyne-azide cycloaddition (CuAAC) click chemistry, was carefully considered in light of simplicity and broad applicability to different cellulose sources.  In this context, the following sections will elaborate on HNE substrate design and cellulose surface modification methods. 14   Figure 1.9 Schematic illustration of HNE bioactive cellulosic surface  On the other hand, we opted for direct chemical activation of Whatman No.1 filter paper through TEMPO oxidation followed by amidation to introduce alkyne groups for copper(I)-catalyzed alkyne-azide cycloaddition (CuAAC) to attach the fluorogen in the final step to create the biosensor. 15   Figure 1.10 Schematic illustration of cellulose-based elastase biosensor by conjugation between azide-terminated fluorogenic substrates and alkyne-functionalized cellulosic surface via click chemistry  16  1.4.1 Design of CuAAC Clickable HNE Fluorogenic Substrates Caged fluorophores (Figure 1.11) were designed as the detecting unit of the biosensor, which contain a tetrapeptide as a specificity element.  Here, tetraalanine was chosen over other possible substrates for HNE for synthetic simplicity. The fluorophore, carboxyrhodamine 110-PEG3-Azide (cRho110-PEG3-N3) provides amine groups suitable for peptide conjugation, by analogy both with the AFC leaving group of soluble HNE substrates (Section 1.2.3) and the fluorescein-based esterase substrate (Section 1.3.3).  Indeed, a bis-tetraalanine conjugate of the related Rhodamine 110 fluorophore has previously been developed as an elastase substrate.34  Additionally, cRho110-PEG3-N3 allows direct conjugation of a terminal azide-containing tetraethylene glycol linker via an available carboxylate group.  It was envisioned that this linker would distance the substrates from the cellulose surface thereby improving HNE accessibility.  The functional group is directly compatible with CuAAC click chemistry on alkyne-functionalized cellulose, for example cotton papers and gauze.    Figure 1.11 Molecular design of (A) divalent fluorogenic substrate, and (B) monovalent fluorogenic substrate   In analytical applications, bis-substituted fluorogenic substrates have well-known limitations. Cleavage of a single functional group typically results in a limited increase 17  fluorescence (ca. 10%), and double cleavage is required to achieve the full spectral properties of the parent fluorophore.35 During this process, the co-existence of fully-cleaved and partially-cleaved fluorophores results in deviations from a linear correlation between signal intensity and enzyme concentration. Therefore, inspired by the work of Raines and co-workers, a fluorogenic substrate was designed that possesses only one tetrapeptide chain (Figure 1.11B).  In this case, the azide-terminated tetraethylene glycol linker is attached via an amide bond to the second amino group on cRho110-PEG3-N3. 1.4.2 Cellulose Surface Activation One of the major challenges in the creation of paper-based biosensors is determination of the optimal surface activation procedure appropriate to enable functionalization. Therefore, cellulose surface modification routes were carefully considered to facilitate the preparation of cellulosic HNE biosensors.  Due to the inherently poor reactivity of cellulose hydroxyl groups, diverse chemical activation strategies have been pursued.  These include reaction with bis-functionalized electrophilic reagents such as divinyl sulfone (DVS)36,37 and p-phenylene diisothiocyanate (PDITC).27,29  The relatively small size and rigid structure of these two activating reagents prevents of crosslinking via reaction with two hydroxyl groups on the same or different polysaccharide chains, while at the same time providing an electrophilic site for secondary modification. Moreover, it promotes direct functionalization of the substrate onto the cellulose surface without the chemical modification steps of cellulose as well as oxidation using (2,2,6,6-tetramethylpiperidin-1-yl)oxyl (TEMPO).38    18   Figure 1.12 (red) TEMPO oxidized cellulosic surface, (blue) PDITC activated cellulose surface, and (green) DVS activated cellulose surface.  As an alternative, the catalytic system comprising (2,2,6,6-tetramethylpiperidin-1-yl)oxyl (TEMPO) with NaOCl as the final oxidant is particularly attractive, as the carboxyl groups resulting from C-6 oxidation of cellulose can be further derivatized through the formation of esters or amides.38,39 Amides, in particular, are generally stable to hydrolysis over a wide pH range, while amidation with propargylamine can be used to install an alkyne suitable for CuAAC click chemistry with HNE probes containing azido groups.  The mechanism of TEMPO oxidation is shown in Figure 1.13.38,39  The reaction is most efficient in the pH range 10.0 to 11.0 and NaBr is added as a co-catalyst that significantly increases the reaction rate. 19   Figure 1.13 TEMPO oxidation on hexopyranose. Reprinted with permission from the Royal Society of Chemistry (39).  1.4.3 Copper(I)-catalyzed Azide-Alkyne Cycloaddition (CuAAC) “Click chemistry”, as defined by Sharpless nearly 20 years ago40, enables the coupling of two molecules through a “facile, selective, [and] high-yield reaction under mild conditions with few or no byproducts”.41 As such, click chemistry has been widely deployed in diverse fields to create functional molecules and materials, including for solid-phase modification.41  Some well-known examples of click chemistry are Diels-Alder, Michael addition, strain-promoted azide-20  alkyne cycloaddition (SPAAC), and copper(I)-catalyzed azide-alkyne cycloaddition (CuAAC, Figure 1.14).41   Figure 1.14 Schematic illustration of copper(I)-catalyzed azide-alkyne cycloaddition reaction. Reprinted with permission from American Chemical Society (41).  Formally, CuAAC is a type of Huisgen 1,3-dipolar cycloaddition, involving “a [regioselective] reaction between terminal alkyne and aliphatic azide in order to form 1,4-disubstituted [1,2,3]-triazoles”.41 Stabilized by aromaticity, the [1,2,3]-triazole product is generally robust over a wide range of conditions, including to redox conditions. Shown in Figure 1.15, The CuAAC mechanism relies on the coordination of copper(I) with two ligands, alkyne and azide in steps I and II. Rearrangement of the metal complex occurs in step III before the release of the final product in step IV.   Figure 1.15 Mechanism of copper(I)-catalyzed azide-alkyne cycloaddition forming 1,4 disubstituted [1,2,3] triazoles. Reprinted with permission from American Chemical Society (41). 21  The following chapters describe combination of the above concepts, i.e., the chemical synthesis and validation of HNE substrates and their conjugation to a model cellulose substrate via click chemistry, toward the development of new point-of-care diagnostic devices to monitor chronic wounds.  22  Chapter 2: Synthesis of HNE Artificial Substrates 2.1 Background and Synopsis This chapter describes the synthesis of two new fluorogenic HNE substrates, featuring either a single- or bis-tetrapetide structure, as illustrated in Figure 1.10. In considering the optimum synthetic route to each molecule, retrosynthetic analyses were performed. In light of the size and molecular complexity of both probes, a combination of HRESI-MS and single- and two-dimensional 1H and 13C NMR methods were used to demonstrate the success of the syntheses.  2.1.1 Proposed synthesis of bis-tetraalanine HNE substrate (5) A retrosynthesis analysis for the bis-substituted molecule (5), comprising two tetraalanine units as the simplest HNE recognition element, is shown in Scheme 2.1. Notably, the alanine units can be introduced either through a convergent synthesis, or a linear sequence, where a peptide bond is formed to the fluorescent core. Starting from the desired final product, disconnection of the amide bonds between the xanthene module and the proximal alanine provides two synthons, a nucleophilic di-aniline (cRho110-PEG3-N3), and two identical electrophilic tetraalanine peptides. However, in previous analogous syntheses the reported yield when attaching two suitably N-protected tetrapeptides in one step was as low as 2%.42  On the other hand, serial additions of single alanine residues onto cRho110-PEG3-N3 would require multiple steps, with the corresponding work-up and purification resulting in a compounding loss of overall yield at each step. Anticipating that the initial coupling might be the most problematic due to reduced nucleophilicity of the rhodamine core versus the primary amine of an amino acid, a convergent approach was selected. Thus, the convergent synthetic route shown in Scheme 2.2. was selected, which utilizes a bis (mono-alanine) rhodamine synthon and N- protected trialanine peptide.  Both the rhodamine-azide linker conjugate and trialanine are commercially available, significantly 23  simplifying the syntheses.  Furthermore, the initial coupling step, involving only a single N-Boc-alanine is anticipated to be amenable to more facile reaction optimization to achieve high yields.  Indeed, N-Boc-alanine is a significantly cheaper reagent than the corresponding tetra- and tri-alanine peptides. The synthesis outlined in Scheme 2.2 thus represents a balance between anticipated reaction efficiency and the overall number of synthetic steps.  Reassuringly, although the synthesis of compound 2 has not been published to the best of my knowledge, there is significant literature precedent for the bis-coupling of amino acids to rhodamine 110 (Rho110), which is structurally analogous to 1, but lacks the azide-terminated tetraethylene glycol linker.35,42  Given that the resulting synthesis from known starting materials is thus reduced to amide bond formation and deprotection steps, some practical an mechanistic considerations will be introduced in the following sections.    Scheme 2.1 Retrosynthesis analysis of HNE divalent fluorogenic substrate (5), PG = protecting group    24   Scheme 2.2 Synthesis scheme of divalent fluorophore 5.  2.1.1.1 Amide Bond Formation (peptide coupling) Amide bond formation has been a topic of study for decades, which has its roots in 1955 with the success in utilizing dicyclohexylcarbodiimide as a coupling reagent. 43 Since then, keen interest in yield improvement, racemization control, and cyclic polypeptide synthesis for pharmaceutical applications has driven a proliferation of alternative coupling-reagents.43–45  Mechanistically, the objective of this reaction is to form a bond between an electrophilic carboxyl carbon atom and a nucleophilic amine nitrogen atom. Despite the high stability of the amide bond, due in part to ℼ-electron delocalization, activation of the OH group of the carboxylic acid in the starting material is required to formally eliminate water.43 Thus, coupling reagents form an ester linkage with the carboxylate under basic conditions in order to generate better leaving group following attack by the nitrogen atom of a free amine (Scheme 2.3). 43    25  A wide range of coupling reagents are commercially available nowadays, which vary in chemical reactivity and physical properties affecting solubility and purification of by-products.  Here, HATU (Scheme 2.3) was chosen as a coupling reagent for this project because of its high reactivity and widespread use in modern peptide syntheses.44 DIPEA was used as a sterically hindered based with limited nucleophilicity to deprotonate carboxylic acid.  Scheme 2.3 Mechanism of amide bond formation by HATU. Reprinted with permission from The Royal Society of Chemistry (43).    Due to the higher price per unit mass, HATU is appropriate for small-scale reactions. In the next stage of the project that requires amidation of propargyl amine and carboxylated cellulose, the traditional carbodiimide-based coupling reagents such as diisopropylcarbodiimide(DIC) and 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC) were proven to be sufficient to provide alkyne-terminated surface. The coupling mechanism of carbodiimides are illustrated in scheme 2.4. Forming the intermediate, O-acylurea, the reaction undergoes different pathway to yield different products. Direct coupling with amine and forming anhydride intermediate finally give amide product. However, O-acylurea is able to undergo a side reaction to form N-acylurea.43 26   Scheme 2.4 Mechanism of amide bond formation by DIC. Reprinted with permission from The Royal Society of Chemistry (43).   2.1.1.2 Boc as a peptide protecting group  The multifunctionality of amino acids is one of the main challenges in polypeptide synthesis. This has lead to the development of a range of protecting groups for the N-terminus, C-terminus, or side-chain functional groups (e.g., thiol, hydroxyl, amine) to prevent undesired side reactions.46,47  In the case of compound 5, the methyl sidechains of tetraalanine significantly simplify the synthesis, and only protection of the N-terminus is required.    In this case, the simpler tert-butyloxycarbonyl (Boc) protecting group was chosen over the other widely used alternative, fluorenylmethyloxycarbonyl (Fmoc), for the following reasons. Firstly, the choice of basic coupling conditions using the tertiary amine DIPEA increase the likelihood of side reactions in the case of Fmoc. Secondly, the use of the Boc protecting group simplifies NMR interpretation due to a single, up-field singlet (~δ 1.40). In contrast, the aromatic 27  protons of Fmoc are observed downfield (δ 6-8 ppm), potentially overlapping with those of the rhodamine core. Lastly, Boc deprotection, work-up, and purification, involving treatment with TFA:DCM (1:1 v/v),35,48 is generally simpler than for Fmoc, which requires a high boiling base and solvent (TEA or piperidine in DMF). 35,48   Boc protection is performed in highly polar solvents such as water, dioxane, or dimethyl formaldehyde due to the solubility of the starting materials. Typically, the amine starting material is mixed in basic solution, e.g., saturated sodium bicarbonate solution, together with tert-butyl dicarbonate (Scheme 2.5). The lone pair of the amine attacks one of the two carbonyl carbons, displacing t-butylcarbonate, which subsequently decarboxylates, releasing carbon dioxide and tert-butoxide that is protonated by water.  An excess of base deprotonates the corresponding intermediate during amide bond formation leading to the final product.       Scheme 2.5 N-terminal Boc protection  The acid-catalyzed Boc deprotection mechanism is shown is Scheme 2.6. Initial protonation of the carbamate results in solvolysis via the tert-butyl cation and subsequent liberation of carbon dioxide.  Since the build-up of CO2 may potentially pressurize the reaction vessel and shift the reaction equilibrium, an open system is suggested.49   28    Scheme 2.6 Boc deprotection mechanism in trifluoroacetic acid (TFA). Reprinted with permission from The Royal Society of Chemistry (49).    2.1.2 Proposed synthesis of mono-tetraalanine HNE substrate (13) The mono-tetraalanine compound 13 (Scheme 2.7) was proposed to avoid potential issues arising from non-linear hydrolysis kinetics of the bis-tetraalanine compound 5.  As with 5, 13 comprises an identical HNE specificity element and linker attached to a fluorogenic core.  However, the molecular asymmetry of 13 required a different synthetic approach. As shown in Scheme 2.7, the retrosynthetic analysis disconnects two distinct amide bonds from the symmetrical rhodamine110 (Rho110), the two amine groups of which are chemically equivalent.  Regardless of the coupling order of the linker and peptide arms, this necessitates careful control of reagent stoichiometry in the first coupling reaction to avoid bis-substitution. Otherwise, the stepwise assembly of the peptide arm is analogous to that of the bis-tetraalanine compound 5.  29   Scheme 2.7 Retrosynthesis analysis of monovalent fluorogenic substrate  The resulting proposed synthetic scheme of the mono-tetraalanine fluorogen is illustrated in Scheme 2.8. The synthesis begins with the mono-Boc- protection of Rho110 to give compound 5 via a published procedure.48 The selectively-protected Rho110 is then coupled with the azide-terminated linker, N3-PEG3-CH2COOH, with DIC as a coupling reagent, followed by Boc deprotection with TFA. Analogous to the synthesis of the bis-tetraalanine compound, a single Boc-L-alanine is then coupled to the core using HATU. Subsequent Boc deprotection, coupling of Boc-trialanine using HATU, and final deprotection yields the target compound.  30   Scheme 2.8 Synthesis scheme of monovalent fluorophore 13   2.2 Results and Discussion 2.2.1 Synthesis of the bis-tetraalanine HNE substrate The synthesis scheme of the bis-tetraalanine flurogenic HNE substrate is illustrated in Scheme 2.2. Within the first step, coupling between two commercially available starting materials, Boc-L-alanine and  cRho110-PEG3-N3, was executed using HATU as the coupling reagent50 to yield compound 2. The synthesis of compound 2, from  cRho110-PEG3-N3 has not been published previously, however analogous reactions involving the coupling of amino acids with rhodamine 110 (Rho110, which lacks the azide-terminated tetraethylene glycol linker) are known.35,42 Deprotection of the Boc protecting group was achieved using trifluoroacetic acid (TFA) prior to  31  HATU-mediated  coupling with N-Boc-trialanine, followed again by Boc deprotection under acidic conditions.  Some specific aspects of each of these steps are detailed in the following sections. 2.2.1.1 Coupling Reaction with HATU and DIPEA Peptide coupling was achieved by mixing of an excess of Boc-Ala-OH (or Boc-tri-Ala-OH), HATU, and DIPEA in DCM, after 20 minutes, followed by addition of the limiting reagent, cRho110-PEG3-N3. After 2 hours, the reddish reaction mixture has turned pale yellow, indicating the success of the coupling reaction by greatly reducing the fluorescence of the fluorophore. After leaving the reaction overnight, TLC indicated that all of the cRho110-PEG3-N3 had been consumed. Consistent with the lower polarity of amide product in comparison with the starting amine, the Rf of compound 2 (and 4) was higher than that of the starting material. Reaction work-up started with a two-phase separation with water to remove ionic by-products, such as DIPEA-PF6 complex, and excess Boc-Ala-OH. After concentration of the organic phase, flash chromatography significantly improved the purity of the product. The yield of this reaction was high (91.8% for compound 2, and 83.8% for compound 4).  2.2.1.2 Boc Deprotection TLC indicated that Boc deprotection proceeded rapidly. To purify the free amines 3 and 5, excess TFA (boiling point 72.4 °C) was eliminated by repeated evaporation under reduced pressure following the addition of chloroform.  Initial attempts to purify compounds 3 and 5 using DCM/MeOH 7:3 v/v as the mobile phase were unsuccessful due to high retention of the diamines on the acidic silica-gel stationary phase. To overcome this problem, the high polarity solvent MeOH was utilized, such that it was possible to purify compound 3 using DCM:MeOH (7:3). For compound 5, 0.1% v/v TEA was added to the eluent to neutralize the acidity of the column. Despite 32  numerous attempts to remove it by evaporation, TEA remained as a contaminant of the final product. Therefore, trituration with hexane and ethyl acetate was used instead of using flash chromatography to purify compound 5. Since it has been reported that TFA can result in by-products due to peptide hydrolysis in Boc deprotection51, to increase the yield of this reaction, milder alternative conditions could be explored, for example, in water at 100°C, 49 or with the assistance of catalysts such as FeCl3, InCl3, AuCl3, etc.52 2.2.1.3 Structural validation of Compound 5 and synthetic intermediates by NMR The target bis-peptide fluorogen 5 is comprised of 27 distinct protons and 36 distinct carbons (Figure 2.1).  The 600 MHz 1H NMR and 150 MHz 13C NMR spectra are shown in Figure 2.2 and 2.3, respectively. Due to the complex structure of the molecule and numerous overlapping signals, the assignment required additional two-dimension NMR techniques to identify proton-proton and proton-carbon correlations.  2D Correlation spectroscopy (COSY, Figure 2.4) was used to detect the magnetization transfer between adjacent protons (3J coupling).  1H-13C Heteronuclear Single Quantum Coherence (HSQC, Figure 2.5) NMR spectroscopy was used to correlate proton signals with carbon signals (1JC-H coupling). Notably, phase information can be obtained by HSQC, thereby eliminating the need to run a separate 13C Distortionless Enhancement by Polarization Transfer (DEPT) experiment.  Finally, Heteronuclear Multiple-Bond Coherence (HMBC, Figure 2.6) reveals short- and long-range coupling between a proton and multiple carbon atoms.   33    Figure 2.1 Atomic numbering of compound 5 for NMR assignment      Figure 2.2 1H-NMR spectrum of compound 5 in MeOD-d4.   34   Figure 2.3 13C-NMR spectrum of 5 in MeOD-d4.    Figure 2.4 2D-COSY spectrum of 5 in MeOD-d4.  35   Figure 2.5 2D-HSQC spectrum of 5 in MeOD-d4.  Carbon signals assigned as -CH and -CH3 – are shown in green - and those assigned as -CH2 – are shown in blue.   Figure 2.6 2D-HMBC spectrum of 5 in MeOD-d4.  36   It is important to note that the starting material, 1, and all resulting compounds are comprised of two structural isomers, in which the linker is attached at either C4´ or C5´ of the phenyl ring.  The existence of these two isomers especially complicates the proton decoupled 13C spectra, giving rise to more peaks than would be expected for a single isomer.  The corresponding 1D and 2D NMR spectra for each of the intermediates and the final product in Scheme 2.2 are presented in the Appendix, Figures A1 – A31.  In the subsequent paragraphs, the assignment of final compound 5 will be described in detail. In 1H NMR of 5, the α-protons of the tetra-alanine groups are observed at δ 4.45, 4.38, 4.32 and 3.96. Three peaks of β methyl protons are at δ 1.51, 1.45, and 1.39 with the integration ratio of 1:1:2 which implies the overlap between two distinct groups of β methyl protons. Among the α-protons, the most upfield proton is at δ 4.45 because of the deshielding effect of the primary amine group. In contrast, the most downfield α-proton is at δ 3.98, which is proximal to the xanthene moiety of cRho110-PEG3-N3. Depending on the compound, the exchangeable amide protons were observed with variable chemical shifts in the range δ 6-10 ppm in in low polarity solvents, e.g. chloroform-d1 and acetone-d6 around, and were not detected in MeOD-d4. Notably, in the proton spectrum of compounds 4 and 2, the 18 protons of the Boc protecting groups overlap with and prevent assignment of the β-methyl protons. The 16 protons of the methylene groups of the tetraethylene glycol linker, H-1’ to H-8’, appear as a multiplet at δ 3.8-3.6. The aromatic protons appear in the downfield region (δ 6.8-8.1). In the downfield region (δ 6.5-8.5) of the 1H NMR spectra of cRho110-PEG3-N3, compound 3, and compound 5, the aromatic signals of xanthene moieties and the phenyl rings are shown and assigned (Figure 2.7). The phenyl proton signals, H3´, H5´, and H6´, were identical in 37  shape and location through peptide coupling reactions with the integrated-area ratio of 1:1:1. In contrast, substitution with amino acid and peptide chains caused the signals from xanthene moiety to shift. H4 and H5 in cRho110-PEG3-N3 (compound 1) at δ 6.70 shifted to δ 7.87 in compound 3, and further separated from each other, δ 7.93 and δ 7.85, after coupling with tripeptide to produce compound 5. The presence of the α-protons of alanines in the region δ 5.0 – 4.0 indicated successful conjugation. In comparison to the starting material (compound 1), the presence of the peak at δ 4.09 in compound 3 is another evidence of the complete coupling of alanine with the ratio of H3´:H5´:H6´:Hα = 1:1:1:2. The four quartets at δ 3.96, 4.32, 4.38, 4.45 with a ratio of 1:1:1:1 confirmed the existence of four α-protons for each of the four alanines in compound 5 (split by the four corresponding methyl groups).  Figure 2.7 Close-up 1H-NMR spectrum of cRho110-PEG3-N3, compound 3 and compound 5 in MeOD-d4.  38  In the proton-decoupled 13C NMR spectrum of compound 5, the signals from the aromatic carbons were observed in the region of δ 100-160 ppm.  Notably, the unique spiro-carbon (C9) was observed at δ 85 ppm. Carbons from the tetraethylene glycol linker were observed at ~δ 68-72). Additional support for the presence of the tetra-alanine groups in compound 5 was obtained from the presence of signals corresponding to the eight α-carbons (δ 49-52 ppm), eight β-methyl carbons (δ 16-18 ppm), and five carbonyl carbons (δ 169-175 ppm). Additionally, the Boc protecting groups in compounds 2 and 4 gave rise to signals at δ 28 ppm (methyl carbons), δ 80 ppm (quaternary carbons) and δ 157 ppm (carbonyl carbons).  2.2.2 Synthesis of Monovalent Fluorogenic Substrate (Compound 13) The synthesis scheme of the mono-tetraalanine flurogenic HNE substrate is illustrated in Scheme 2.8. Selective Boc- protection of Rho110 (compound 6) was achieved to afford compound 7 by the procedure previously described48, prior to the coupling reaction with the linker, N3-PEG3-CH2COOH , in the presence of DIC and DMAP to obtain compound 8. A solution of TFA in DCM was utilized to remove the Boc- protecting groups, yielding compound 9 prior to the N-protected peptide coupling/deprotection cycles analogous to the synthesis of bis-tetraalanine substrate, compound 5. Some specific aspects of each of these steps are detailed in the following sections. 2.2.2.1 Selective Boc Protection of Rho110 The mono-protection of Rho110 was achieved by carefully controlling the amount of di-tert-butyl dicarbonate to slightly more than one equivalent. Initially, Rho110 was mixed with NaH in DMF under argon in order to generate the basic conditions in which Rho110 would be in the lactone form. The required amount of di-tert-butyl dicarbonate was added, and the reaction was stirred overnight until a TLC test indicated the formation of the fluorescent product, compound 7. 39  The reaction was then quenched with acetic acid. The crude thus obtained was purified via flash silica gel chromatography with a yield of 25%, comparable to the yield of 33%.48 2.2.2.2 Coupling of the linker with compound 7 Amide bond formation of the linker N3-PEG3-CH2COOH with compound 7 was achieved in the presence of DIC and DMAP. The reaction was initiated at 0 °C and then warmed room temperature. After stirring at room temperature for 2 h, the original reddish reaction mixture turned pale yellow. Eventually after 24 h of stirring at room temperature, the reaction mixture turned colorless. TLC tests indicated that the reaction was complete. The reaction mixture in DCM was then washed with water for 3 times. The solvent was removed and the crude thus obtained was concentrated and loaded onto the top of a silica gel column. Compound 8 was obtained, after elution with hexane and ethyl acetate (4:6), with a yield of 73%. 2.2.2.3 Boc- Deprotection The deprotection of compound 8, compound 10, and compound 12 was achieved in the similar manner. Each Boc- protected compound was dissolved in a mixture of DCM and TFA (1:1) and stirred for 30 mins to 1 hr until TLC indicated complete deprotection. Residual TFA was removed by repetitive addition of chloroform and evaporation under reduced pressure for 3 times. The crude compound was then purified by flash chromatography using hexane/ethyl acetate (4:6) as an eluent. Pure compound 9 and compound 11 were obtained with a yield of 58% and 71% respectively, as evidenced by 1H and 13C NMR as well as low and high resolution mass spectroscopy. Compound 13 was obtained by a slightly different procedure due to difficulty in monitoring the process of column purification. The pure compound was obtained by trituration with ethyl 40  acetate and hexane with a yield of 49%. The compound 13 was characterized by 1H and 13C NMR as well as low and high resolution mass spectroscopy.  2.2.2.4 Structural validation of Compound 13 and synthetic intermediates by NMR The target mono-peptide fluorogen 13 is comprised of 31 distinct protons and 40 distinct carbons (Figure 2.8). The 600 MHz 1H NMR and 150 MHz 13C NMR spectra are shown in Figure 2.9 and 2.10, respectively. Additional two-dimension NMR techniques were utilized to identify proton-proton and proton-carbon correlations due to the complex structure of the molecule and numerous overlapping signals.  2D Correlation spectroscopy (COSY, Figure 2.11) was used to detect the magnetization transfer between adjacent protons (3J coupling).  1H-13C Heteronuclear Single Quantum Coherence (HSQC, Figure 2.12) NMR spectroscopy was used to correlate proton signals with carbon signals (1JC-H coupling). Notably, phase information can be obtained by HSQC, thereby eliminating the need to run a separate 13C Distortionless Enhancement by Polarization Transfer (DEPT) experiment.  Finally, Heteronuclear Multiple-Bond Coherence (HMBC, Figure 2.13) reveals short- and long-range coupling between a proton and multiple carbon atoms.    Figure 2.8 Structure and position numbering of compound 13 and its intermediates  41    Figure 2.9 1H-NMR spectrum of 13 in MeOD-d4.  Figure 2.10 13C-NMR spectrum of 13 in MeOD-d4. 42   Figure 2.11 2D-COSY spectrum of 13 in MeOD-d4.  Figure 2.12 2D-HSQC spectrum of 13 in MeOD-d4. 43   Figure 2.13 2D-HMBC spectrum of 13 in MeOD-d4.  The corresponding 1D and 2D NMR spectra for each of the compounds listed in Scheme 2.8 are presented in the Appendix, Figures A34 – A84.  In the subsequent paragraphs, the assignment of final compound 13 will be described in detail. In the upfield region, three signals of β methyl protons, at δ 1.52, 1.45, and 1.39, with the integration ratio of 1:1:2 confirm the tetrapeptide coupling with two overlapping signals. Furthermore, in the downfield region, the characteristic aromatic protons of Rho110 are observed between δ 6.0 - 9.0 ppm as further evidence of the conjugation. In the region of δ 3.5 – 3.8 ppm, the 10 protons of the tetraethylene glycol linker, N3-PEG3-CH2COOH , appear as multiplets, while two protons at alpha carbon of linker are observed as a singlet at δ 4.16 ppm. The other two protons of the carbon adjacent to the azide group is a triplet at δ 3.36. For intermediates with a Boc- 44  protecting group, such as compound 10 and compound 12, all the three identical methyl groups on the Boc- protecting groups are observed as singlet at δ 1.40 ppm.  In the downfield region (δ 6.3-8.5 ppm) of the 1H NMR spectra of Rho110, compound 9, compound 11 and compound 13 shown in Figure 2.14, the aromatic signals of the xanthene moiety and the phenyl rings have been assigned. The selective attachment of the linker to one side of Rho110 provides asymmetry to compound 9. The free electron pair of the amine group is able to localize toward the attached ring causing shielding effect and, as a result, an upfield shift of H5, H7 and H8. Coupling of compound 9 with Boc-Ala-OH has an opposite effect and hence, only results in a very subtle change to the corresponding aromatic protons.  In the mono-peptide substrate, compound 13, the observation of α-protons of alanine in the region of δ 4.0-5.0 ppm provides evidence of conjugation. Due to difficulties in assigning the α-proton of alanine of compound 11, 2D COSY and 2D HSQC were performed to show that the α-proton overlaps with those from the ethylene glycol linker (Figure A59). Four quartets corresponding to 4 distinct alpha protons, at δ 3.96, 4.32, 4.38 and 4.45 ppm, indicate successful coupling to producecompound 13, with the ratio of H3´: Hα = 1:4.   45   Figure 2.14 Close-up 1H-NMR spectrum of Rho110 in MeOD-d4, compound 9 in DMSO-d6, compound 11 in acetone-d6 and compound 13 in MeOD-d4  In the proton-decoupled 13C NMR spectrum of compound 13, the signals from the aromatic carbons were observed in the region of δ 100-160 ppm.  Notably, the unique spiro-carbon (C9) was observed at δ 84.13 ppm. Carbons from the tetraethylene glycol linker were observed at ~δ 70-73. Importantly, the α carbons of tetra-alanine are observed at δ 51.80, 50.69, 50.50, and 50.14 ppm. Four β methyl carbons are observed at δ 17.92, 17.85, 17.53, and 17.43, while five carbonyl carbons, from four alanines and a linker, are detected at δ 174.62, 174.27, 173.42, and 171.49 ppm.   46  2.3 Summary In this chapter, the proposed HNE fluorogenic substrates were successfully synthesized. To obtain the bis-tetraalanine compound 5 from the commercially available starting material, cRho110-PEG3-N3, retrosynthetic analysis was utilized to design a 4-step pathway, which consisted of two cycles of amide coupling by HATU followed by Boc deprotection in TFA. On the other hand, because of its asymmetrical structure, mono-tetraalanine compound 13 required a 7-step process, due to an additional Boc protection/deprotection cycle and linker coupling reaction. In order to confirm the success of the syntheses, 2D NMR techniques, specifically COSY, HSQC, and HMBC, were used to characterize all intermediates and final products, allowing assignment of signals in 1H and 13C NMR spectra. Furthermore, all compounds have been confirmed by positive-mode high-resolution ESI mass spectroscopy.  With the two compounds in hand, their potential as HNE substrates in solution and on paper surfaces was examined, as described in the next chapter.        47  2.4 Experimental Section 2.4.1 Materials  cRho110-PEG3-N3 (catalog number: BP-22478) was purchased from Broadpharm (San Diego, CA, USA). Synthetic peptides including Boc-L-alanine (catalog number: A-1060.0100BA) and tri-L-alanine-OH (catalog number: H-1225.0001BA) were purchased from Bachem Americas, Inc. (Torrance, CA, USA). HPLC-grade HATU (99%) (catalog number: A518) was from AK Scientific, Inc. (Union City, CA, USA). Rho110 (chloride) (catalog number: 19061) was from Cayman Chemical Company (Ann Arbor, MI, USA). DIPEA and other reagents were purchased from Thermo Fisher Scientific Chemicals, Inc. (Ward Hill, MA, USA). All HPLC grade organic solvents were from MilliporeSigma Canada Co. (Oakville, ON, Canada). Native human neutrophil elastase protein (Catalog number: ab91099) was purchased from Abcam Inc. (Cambridge, MA, USA). For moisture-sensitive reactions, glassware was dried in a ~100°C oven and purged with inert gas (N2 or argon) before usage. 2.4.2 Thin Layer Chromatography (TLC) TLC was performed on aluminum plates (0.25 mm) pre-coated with Merck silica gel 60 F254, and was visualized using UV light (254 nm), 10% H2SO4 in MeOH, ninhydrin strain, and/or molybdate stain, including heating with a heat gun as appropriate. Silicycle SiliaFlash F60 (40-63 µm, 230-400 mesh) silica gel was used for all flash chromatography. 2.4.3 1D NMR and 2D NMR All 1H and 13C one dimensional NMRs were performed using a Bruker Avance 400 MHz spectrometer at 25°C. Chemical shifts are reported in parts per million (ppm) and coupling constants (J) in Hertz (Hz).  All 1H experiments were referenced to residual protonated solvent signals: acetone (δ 2.05), chloroform (δ 7.26), dimethylsulfoxide (δ 2.50) and methanol (δ 3.31).53 48  All 13C experiments were referenced to residual protonated solvent signals:  acetone peak (δ 29.84), chloroform (δ 77.16), dimethylsulfoxide (δ 39.52) and MeOD (δ 49.00).53 2D NMR experiments were performed using a Bruker Avance 600 equipped with a z-gradient TCI cryoprobe with a 1H resonance at 600.15 MHz and a 13C resonance at 150.92 MHz. Chemical shifts are recorded in ppm relative to the residual solvent peak in each spectrum, as above. NMR data was analyzed using Topspin data processing software.  2.4.4 High-Resolution ESI MS Positive-mode high resolution mass spectra (HRMS) were recorded on a Waters/Micromass liquid chromatography tandem time of flight mass spectrometer (LC-TOF MS) equipped with an electrospray ionization source.  2.4.5 Synthesis of compound 5 Synthesis of 2 Boc-L-Alanine (65.9 mg, 348.2 mmol) and HATU (132.4 mg, 348.2 mmol) were dissolved in 4 mL of DCM at RT for 10 min before DIPEA (135.0 mg, 1044 mmol) was added. The solution turned pale yellow after stirring for 20 min.  cRho110-PEG3-N3 (1) (25 mg, 43.5 mmol) was added into the reaction mixture and stirred for 24 h. The reaction mixture was washed with water (3×3mL). The organic phase was concentrated and purified by flash chromatography (mobile phase dichloromethane:acetone = 6:4, Rf = 0.4) followed by evaporation of the solvent under reduced pressure to yield compound 2 as a colorless solid (36.7 mg, 91.8 % yield).  1H-NMR (Fig A1, 400 MHz, acetone-d6): δ 9.55 (s, 1H, Boc-Ala-NH-), 9.53 (s, 1H, Boc-Ala-NH-), 8.23 (dd, J1=8.0, J2= 1.1, 1H, H5´), 8.07 (d, J=8.0, 1H, H6´), 7.99 (d, J=5.7, 1H, H4 or H5), 7.98 (d, J=1.9, 1H, H4 or H5), 7.91 (s, 1H), 7.75 (s, 1H, H3´), 7.24 (dd, J1=8.7, J2= 1.9, 1H, H2 or H7), 7.19 (d, J=8.3, 1H, H2 or H7), 6.82 (d, J=8.6, 1H, H1 or H8), 6.81 (d, J=8.6, 1H, H1 or H8), 6.32 49  (s, 2H, Boc-NH-CH), 4.28 (p, J=6.34, 2H, Hα), 3.59 (t, J=5.0, 2H, -OCH2CH2N3), 3.56-3.44 (m, 12H, -OCH2CH2O-), 3.32 (t, J=4.9, 2H, -OCH2CH2N3), 1.41 (s, 18H, (CH3)3-COC=O), 1.39 (d, J=1.8, 6H, Hβ). 13C-NMR(Fig A2, 100 MHz, acetone-d6) : δ 172.97, 168.82, 165.91, 156.49, 154.20, 152.34, 142.38, 142.20, 130.43, 129.44, 129.23, 125.70, 123.30, 116.26, 114.43, 107.76, 82.92, 79.55, 71.13, 70.81, 70.60, 69.97, 51.96, 51.31, 40.67, 28.54, 18.32. HRMS (ESI): Calculated for C45H57N8O13 [M+H+]: 917.4045; observed: 917.4040.  Synthesis of 3 To 4 mL of a DCM solution of the resulting solid (36.7 mg, 40.0 mmol), 4 mL of TFA was added dropwise and stirred at room temperature. After 1 hour, the DCM and TFA were removed by rotary evaporation. The resulting crude was purified by flash chromatography (mobile phase DCM:MeOH = 7:3, Rf = 0.2). The corresponding fractions were collected and evaporated to obtain compound 3 as a pale yellow solid (26.1 mg, 90.9% yield). 1H-NMR (Fig A9, 400 MHz, MeOD-d4) : δ 8.17 (dd, J1=8.1, J2= 1.3, 1H, H5´), 8.12 (dd, J1=8.1, J2= 0.5, 1H, H6´), 7.87 (q, J=0.9, 2H, H4 and H5), 7.64 (t, J=0.9, 1H, H3´), 7.23 (dt, J1=8.7, J2= 1.9, 2H, H2 and H7), 6.82 (d, J=8.7, 2H, H1 and H8), 4.09 (q, J=7.1, 2H, Hα), 3.60-3.48 (m, 14H, -OCH2CH2O-), 3.28 (t, J=5.0, 2H, -OCH2CH2N3), 1.61 (dd, J1=7.0, J2= 1.4, 6H, Hβ). 13C-NMR(Fig A10, 100 MHz, MeOD-d4) : δ 170.18, 169.70, 168.12, 154.62, 152.83, 142.65, 141.77, 130.71, 129.80, 129.70, 126.37, 123.90, 116.92, 115.44, 108.80, 83.85, 71.56, 71.48, 71.40, 71.11, 71.01, 70.25, 51.71, 50.99, 41.12, 17.49. HRMS (ESI): Calculated for C35H40N8O9Na [M+Na+]: 739.2816; observed: 739.2819.   50  Synthesis of Boc-tri-L-alanine-COOH  Scheme 2.9 Synthesis of Boc-tri-L-alanine-COOH   Tri-L-alanine (100mg, 0.43 mmol) and di-tert butyl dicarbonate (189 mg, 0.86 mmol) were dissolved in dioxane (1.5 mL) and a saturated solution of NaHCO3 (1.5 mL) was added as shown in Scheme 2.9. The reaction was stirred overnight at room temperature. After that, 3×3 mL of EtOAc was used to wash the aqueous phase. Followed by acidification with 6N HCl to pH 1, the product was extracted with 3×5 mL of ethyl acetate and the combined organic fraction was washed with brine (3×10 mL). Finally, the organic phase was dried over MgSO4 and evaporated to dryness, giving a white powder (89.4 mg, 62.4% yield).  1H-NMR (Fig A32, 400 MHz, MeOD-d4): δ 4.38 (quin, J=7.7, 2H), 4.08 (q, J=7.2, 1H), 1.44 (s, 9H), 1.40 (d, J=7.4, 3H), 1.37 (d, J=7.1, 3H), 1.31 (d, J=7.3, 3H). 13C-NMR (Fig A33, 100 MHz, MeOD-d4): δ 175.66, 175.50, 174.44, 157.70, 80.60, 51.46, 49.97, 28.68, 18.27, 18.19, 17.64. HRMS (ESI): Calculated for C35H40N8O9Na [M+Na+]: 739.2816; observed: 739.2819. Synthesis of 4  Boc-tri-Alanine-OH (129.3 mg, 390.5 mmol) and HATU (148.5 mg, 390.5 mmol) were dissolved in 5 mL of DCM at RT for 10 min before adding DIPEA (151.4 mg, 1171 mmol). The solution turned pale yellow after stirring for 20 min. Compound 3 (24.1 mg, 33.6 mmol) was added into the reaction mixture and stirred for 24 h. The reaction mixture was washed with water (3×5mL). The organic phase was concentrated and purified by column chromatography (mobile 51  phase DCM:Acetone:MeOH = 8:1:1, Rf = 0.4) followed by evaporation of the solvent under reduced pressure to yield compound 4 as a colorless solid (41.0 mg, 83.8 % yield).  1H-NMR (Fig A11, 400 MHz, acetone-d6) : δ 9.08 (d, J=8.0, 2H, Ala-C=O-NH-C3), 8.23 (dd, J1=8.0, J2= 1.4, 1H, H5´), 8.19 (d, J=1.8, 1H, H4 or H5), 8.17 (s, 2H, -NH-Ala-C=O), 8.10 (d, J=1.8, 1H, H4 or H5), 8.07 (dd, J1=8.1, J2= 0.5, 1H, H6´), 7.91 (d, J=5.8, 2H, -NH-Ala-C=O), 7.74 (q, J=0.7, 1H, H3´), 7.62 (d, J=7.7, 2H, -NH-Ala-C=O), 7.60 (dd, J1=9.0, J2= 1.8, 1H, H2 or H7), 7.52 (dd, J1=9.0, J2= 1.8, 1H, H2 or H7), 6.81 (dd, J1=8.7, J2= 6.8, 2H, H1 and H8), 6.69 (t, J=4.5, 2H, -NH-Ala-C=O), 4.42 (dquin, J1=7.3, J2= 2.5, 2H, Hα), 4.20-4.16 (m, 4H, Hα), 3.98 (dq, J1=7.0, J2= 3.8, 2H, Hα), 3.58 (t, J=4.9, 2H, -OCH2CH2N3), 3.53-3.45 (m, 12H, -OCH2CH2O-), 3.32 (t, J=4.9, 2H, -OCH2CH2N3), 1.50 (d, J=7.3, 6H, Hβ), 1.45 (s, 18H, (CH3)3-COC=O), 1.42 (dd, J1=7.4, J2= 2.2, 12H, Hβ), 1.37 (d, J=7.3, 6H, Hβ). 13C-NMR(Fig A12, 100 MHz, acetone-d6) : δ176.51, 175.67, 173.03, 172.53, 168.92, 165.92, 157.79, 154.52, 152.29, 142.40, 142.36, 142.32, 130.56, 129.27, 129.16, 125.66, 123.08, 116.60, 114.41, 107.77, 82.94, 80.55, 71.18, 71.13, 70.81, 70.59, 69.99, 53.22, 52.58, 51.70, 51.33, 50.86, 40.62, 32.63, 30.34, 28.56, 17.63, 17.30, 17.06, 17.04. HRMS (ESI): Calculated for C63H87N14O19 [M+H+]: 1343.6273; observed: 1343.6265. Synthesis of 5 In 4 mL of a DCM solution of 4 (41.0 mg, 40.0 mmol), 4 mL of TFA was added dropwise and stirred at RT. After 1 hour, DCM and TFA were removed by a rotary evaporation. 10 mL of chloroform was added and evaporated 3 times to dryness to remove residual TFA in the crude. The resulting solid was triturated by hexane (2×10 mL), EtOAc (2×10 mL) and acetone (1×10 mL) before drying under reduced pressure to obtain compound 5 as a pale orange solid (6.7 mg, 19.2% yield). 52  1H-NMR (Fig A19, 600.15 MHz, MeOD-d4) : δ 8.17(dq, J1=8.1, J2= 1.7, 1H, H5´), 8.12 (dtt, , J1=8.1, J2= 4.7, J3= 0.8, 1H, H6´), 7.93 (dd, J1=9.8, J2= 2.0, 1H, H4 or H5), 7.85 (m, 1H, H4 or H5), 7.66 (s, 1H, H3´), 7.31 (td, J1=13.3, J2= 2.0, 1H, H2 or H7), 7.22 (m, 1H, H2 or H7), 6.78 (m, 2H, H1 and H8), 4.45 (q, J=6.7, 2H, Hα), 4.38 (q, J=7.2, 2H, Hα), 4.32 (qd, J1=7.0, J2= 1.1, 2H, Hα), 3.96 (q, J=6.6, 2H, Hα), 3.61-3.47 (m, 14H, -OCH2CH2O-), 3.28 (t, J=4.9, 2H, -OCH2CH2N3), 1.52 (dd, J1=7.6, J2= 1.6, 6H, Hβ), 1.46 (d, J=7.3, 6H, Hβ), 1.40 (dd, J1=7.1, J2= 2.4, 12H, Hβ). 13C-NMR (Fig A20, 150.03 MHz, MeOD-d4): δ 173.78/173.65, 172.52, 170.15, 169.30, 167.17, 162.17/161.94, 153.77, 151.87, 141.68, 141.36/141.24, 129.76, 128.81/128.63, 125.36, 122.97, 116.32/116.01, 113.95, 107.90/107.70, 83.27, 70.62, 70.53, 70.45, 70.14, 70.06, 69.28,  HRMS (ESI): Calculated for C53H71N14O15 [M+H+]: 1143.5223; observed: 1143.5218. 2.4.6 Synthesis of compound 13 Synthesis of 7 Compound 7 was synthesized according to a previously reported method:48 Rho110 hydrochloride (200 mg, 0.55 mmol) was dissolved in 10 mL of anhydrous DMF, before NaH powder, 60 %wt moistened with oil (43 mg, 1.10 mmol), was slowly added. The reaction was stirred for 1 h under argon, at which time it became dark brown. Di-tert-butyl dicarbonate (99 mg, 0.46 mmol) was then added, and the solution turned red overnight. Acetic acid (1 mL) was used to quench the NaH, and DMF was partially removed by a rotary evaporation. Flash chromatography (hexane:DCM:EtOAc =5:3:2) was used to purify the product, yielding compound 7 as a red powder (66 mg, 25% yield). 1H-NMR (Fig A34, 400 MHz, acetone-d6): δ 8.67 (s, 1H), 7.97 (dt, J1=7.6, J2= 0.7, 1H), 7.79 (td, J1=7.5, J2= 1.0, 1H), 7.72 (s, 1H), 7.71 (td, J1=7.5, J2= 0.7, 1H), 7.26 (dt, J1=7.6, J2= 0.7, 1H), 53  7.13 (dd, J1=8.7, J2= 2.2, 1H), 6.68 (d, J=8.6, 1H), 6.57 (d, J=2.1, 1H), 6.48 (d, J=8.5, 1H), 6.43 (dd, J1=8.5, J2= 2.1, 1H), 5.17 (s, 2H), 1.49 (s, 9H). 13C-NMR (Fig A35, 100 MHz, acetone-d6): δ 169.61, 154.08, 153.55, 152.89, 151.95, 142.64, 135.86, 130.56, 129.59, 129.21, 128.04, 125.23, 124.92, 114.70, 114.27, 112.23, 107.86, 106.12, 100.87, 84.14, 80.56, 28.43. HRMS (ESI): Calculated for C25H23N2O5 [M+H+]: 431.1607; observed: 431.1608. Synthesis of 11-Azido-3,6,9-trioxaundecanoic Acid (N3-PEG3-CH2COOH)   Scheme 2.10 General synthesis of N3-PEG3-CH2COOH   The synthesis of the linker N3-PEG3-CH2COOH has been previously reported (Scheme 2.10).54 [2-[2-[2-chloroethoxy]ethoxy]ethanol (17.7 g, 0.1 mol) was mixed with 100 mL of water and sodium azide (13.0 g, 0.2 mol). The reaction mixture was heated to 75°C for 24 h with stirring. Water was then removed under reduced pressure until 30 ml remained. The aqueous solution was extracted by 10 ml of DCM for 3 times. The organic phase was dried with anhydrous sodium sulfate, filtered, and removed by rotary evaporation, yielding a colorless liquid (16.9 g, yield = 92%). The resulting azido-alcohol (7.6g, 0.04 mol) was dissolved in 200 mL of dry THF. NaH (60% powder, 2.2 g, 0.06 mol) was slowly added, and the solution was stirred for 1 h. Ethyl bromoacetate (10 mL, 0.06 mol) was added dropwise to the solution, which consequently turned yellow and then pale orange over 30 min. The reaction was stirred overnight, monitored by TLC, and quenched with ethanol (10 mL). THF was evaporated to dryness, and the crude was dissolved 54  in 120 mL of water. The product was extracted by 3×100 mL DCM before drying over Na2SO4 and rotary evaporation, giving a amber liquid (12.1 g). The resulting ethyl ester was then hydrolyzed with 30 mL of 3 M NaOH at room temperature for 18 hours. After washing with 30 mL of DCM 3 times, the aqueous phase was acidified with 10 %wt HCl to pH 1, before extraction with ethyl acetate (3 x 50 mL). The organic phase was collected and dried over Na2SO4. Upon rotary evaporation and further drying, the resulting product was a yellow liquid (5.1 g, 50.5 % yield). 1H-NMR (Fig A85, 400 MHz, chloroform-d1): δ 9.59 (s, 1H), 4.15 (s, 2H), 3.74-3.63 (m, 10H), 3.36 (t, J= 5.0 Hz, 2H). Synthesis of 8  N3-PEG3-CH2COOH (44.7 mg, 0.19 mmol), DIC (29.0 mg, 0.23 mmol) and DMAP (23.4 mg, 0.19 mmol) were mixed in 4mL of anhydrous DCM at 0°C. After 15 minutes, compound 7 (66 mg, 0.15 mmol) was added. The solution was stirred under argon and gradually turned yellowish, then colorless. The reaction mixture was washed by 4 ml of deionized water for 3 times before DCM was then removed on a rotary evaporator. The crude was purified by flash chromatography (hexane: EtOAc =4:6) resulting in compound 8 as a colorless solid (73 mg, 73% yield).  1H-NMR (Fig A36, 400 MHz, acetone-d6): δ 9.39 (s, 1H), 8.76 (s, 1H), 8.01 (dt, J1=7.6, J2=0.8, 1H, H3´), 7.97 (d, J=2.0, 1H, H4), 7.82 (td, J1=7.5, J2=0.9, 1H, H5´), 7.77 (s, 1H, H5), 7.75 (td, J1=7.5, J2=0.7, 1H, H4´), 7.32-7.28 (m, 2H, H2 and H6´), 7.21 (dd, J1=8.7, J2=2.1, 1H, H7), 6.81 (d, J=8.7, 1H, H1), 6.75 (d, J=8.7, 1H, H8), 4.16 (s, 2H), 3.80-3.70 (m, 8H), 3.67 (t, J=5.0, 2H), 3.38 (t, J=9.8, 2H), 1.50 (s, 9H). 55  13C-NMR (Fig A37, 100 MHz, acetone-d6): δ 169.85, 169.49, 153.96, 152.58, 152.56, 152.47, 142.97, 141.31, 136.26, 136.20, 130.94, 130.90, 129.48, 129.29, 129.26, 127.46, 125.49, 124.93, 124.89, 122.18, 116.47, 116.42, 115.44, 115.27, 115.21, 113.68, 108.25, 108.00, 106.17, 82.90, 80.69, 79.40, 71.83, 71.16, 71.03, 70.95, 70.83, 70.53, 51.30, 38.72, 28.43. HRMS (ESI): Calculated for C33H36N5O9 [M+H+]: 646.2513; observed: 646.2513. Synthesis of 9  1 mL of TFA was added dropwise to a 1 mL of DCM solution of 8 (73 mg, 0.11 mmol),  and stirred for 1 h. DCM and TFA were evaporated under vacuum and the crude was purified by flash chromatography (hexane: EtOAc = 4:6) to yield compound 9 as a red powder (90 mg, 58% yield). 1H-NMR (Fig A47, 400 MHz, DMSO-d6): δ 9.97 (s, 1H), 8.00 (d, J=7.7, 1H, H3´), 7.86 (s, 1H, H4), 7.79 (td, J1=7.5, J2=0.2, 1H, H5´), 7.71 (t, J=7.5, 1H, H4´), 7.27 (d, J=7.6, 1H, H6´), 7.24 (dd, J1=8.7, J2=1.1, 1H, H2), 6.70 (d, J=8.3, 1H, H1), 6.50 (s, 1H, H5), 6.41 (q, J=11.0, 2H, H7  and H8), 4.11 (s, 2H, -C=OCH2-OCH2CH2O-), 3.67-3.54 (m, 10H, -OCH2CH2O-), 3.35 (t, J= 4.9, 2H, -OCH2CH2N3). 13C-NMR (Fig A48, 100 MHz, DMSO-d6): δ 169.63, 169.28, 152.49, 151.94, 135.98, 130.68, 129.40, 128.97, 127.21, 125.60, 125.05, 116.14, 114.86, 112.66, 112.49, 107.34, 106.00, 99.77, 85.86, 70.97, 70.79, 70.40, 70.23, 69.83, 50.60. HRMS (ESI): Calculated for C28H27N5O7Na [M+Na+]: 568.1808; observed: 568.1806. Synthesis of 10  Boc-L-alanine (12.5 mg, 0.07 mmol), HATU (30.1 mg, 0.08 mmol) and DIPEA (21.3 mg, 0.16 mmol) were mixed in 1 mL of anhydrous DCM. After 30 minutes, compound 9 (30 mg, 0.05 mmol) was added and the solution was stirred under inert atmosphere, gradually turning yellow, 56  then colorless. The solvent was evaporated upon a rotary evaporator, and the crude was purified by flash chromatography (hexane: EtOAc =4:6) to yield compound 10 as a colorless solid (25 mg, 64% yield).  1H-NMR (400 MHz, chloroform-d1) : δ 9.54 (s, 1H), 9.33 (s, 1H), 8.02-3.97 (m, 3H), 7.81 (td, J1=7.5, J2=0.8, 1H), 7.75 (td, J1=7.5, J2=0.6, 1H), 7.34-7.31 (m, 2H), 7.23 (dd, J1=8.3, J2=1.8, 1H), 6.82 (d, J=8.6, 1H), 6.78 (d, J=8.6, 1H), 6.28 (d, J=6.0, 1H), 4.27 (quint, J=6.4, 1H), 4.12 (s, 2H), 3.79-3.68 (m, 8H), 3.66 (t, J=5.0, 2H), 3.34 (t, J=4.9, 2H), 1.43 (s, 9H), 1.41 (d, J=7.2, 3H). HRMS (ESI): Calculated for C36H40N6O10 [M+H+]: 717.2885; observed: 717.2883. Synthesis of 11  One mL of TFA was added dropwise to a 1 mL DCM solution of 10 (25 mg, 0.03 mmol), and the reaction was stirred for 3 h in an open flask to allow carbon dioxide to escape. DCM and TFA were evaporated under vacuum and the crude was purified by flash chromatography (hexane: EtOAc = 4:6) to give compound 11 as a colorless powder (15 mg, 71% yield). 1H-NMR (Fig A57, 400 MHz, acetone-d6): δ 9.35 (s, 1H), 8.03 (s, 1H, H4), 8.02 (d, J=5.1, 1H, H3´), 7.83 (t, J=7.3, 1H, H5´), 7.77 (t, J=7.4, 1H, H4´), 7.38 (d, J=7.5, 1H, H6´), 7.34-7.31 (m, 2H, H2 and H5), 7.04 (dt, J1=8.4, J2=2.2, 1H, H7), 6.92 (d, J=8.4, 1H, H8), 6.85 (d, J=8.6, 1H, H1), 4.12 (s, 2H, -C=OCH2-OCH2CH2O-), 3.79-3.64 (m, 11H, -OCH2CH2O- and Hα), 3.33 (t, J=4.8, 2H, -OCH2CH2N3), 1.34 (dd, J1=6.8, J2=2.4, 3H, Hβ). 13C-NMR (Fig A58, 100 MHz, acetone-d6): δ 176.39, 169.71, 169.39, 153.85, 152.27, 141.49, 140.01, 136.37, 131.09, 129.33, 129.29, 127.30, 125.65, 125.03, 124.75, 124.45, 117.40, 116.60, 115.08, 107.88, 82.43, 71.96, 71.27, 71.16, 71.03, 70.94, 70.61, 54.60, 51.33, 17.02. HRMS (ESI): Calculated for C31H33N6O8 [M+H+]: 617.2360; observed: 617.2354.  57  Synthesis of 12  Boc-tri-alanine (17.4 mg, 0.05 mmol), HATU (20.0 mg, 0.05 mmol) and DIPEA (20.4 mg, 0.15 mmol) were mixed in 2 mL of anhydrous DCM. After 30 min, compound 11 (10.8 mg, 0.02 mmol) was added. The solution was stirred under inert atmosphere and the progress of the reaction was examined using TLC. After 24 h, the reaction mixture was washed with 3 mL of deionized water 3 times before evaporating the organic phase on a rotary evaporator. The crude was purified by flash chromatography (DCM: Acetone = 4:6) to yield compound 12 as colorless solid (12.5 mg, 76% yield). Full characterization of compound 12 was not performed due to difficulties in NMR signal assignment resulting from overlap between the exchangeable protons of the 4 amide groups peaks in the aromatic region. Hence, full characterization of the final, deprotected product was carried out instead. HRMS (ESI): Calculated for C45H56N9O13 [M+H+]: 930.3998; observed: 930.3995. Synthesis of 13  0.5 mL of TFA was added dropwise to a 0.5 mL of DCM solution of 12 (12.5 mg, 0.013 mmol), and the reaction was stirred for 1 h in an open flask. DCM and TFA were evaporated under vacuum, followed by repeated addition of 10 mL of chloroform and rotary evaporation (3 times). The resulting solid was triturated by hexane (2×10 mL), EtOAc (2×10 mL) and acetone (1×10 mL) before drying under vacuum to obtain compound 13 as a pale orange solid (5.5 mg, 49% yield). 1H-NMR (Fig A69, 600 MHz, MeOD-d4): δ 8.04 (dtt, J1=3.2, J2=0.9, 1H, H3´), 7.92 (d, J=3.2, 1H, H4) 7.90-7.71 (m, 3H, H4´, H5´ and H5), 7.25 (t, J=8.3, 2H, H6´, H2 and H7), 6.76 (t, J=8.3, 2H, H1 and H8), 4.47 (q, J=7.1, 1H, Hα), 4.40 (q, J=7.2, 1H, Hα), 4.33 (q, J=7.1, 1H, Hα), 4.16 (s, 2H, -C=OCH2-OCH2CH2O-), 3.94 (qd, J1=7.1, J2=1.5, 1H, Hα), 3.80-3.61 (m, 10H, -OCH2CH2O-), 3.36 (s, 2H, -OCH2CH2N3), 1.53-1.37 (m, 12H, Hβ). 58  13C-NMR (Fig A70, 150 MHz, MeOD-d4): δ 174.77, 174.63, 174.28, 173.42, 171.49, 171.45, 171.27, 154.43, 152.98, 142.19, 142.11, 141.42, 136.86, 131.41, 129.50, 129.47, 127.68, 126.02, 125.18, 117.32, 116.87, 115.84, 115.61, 115.48, 109.12, 108.67, 84.14, 72.15, 71.66, 71.57, 71.43, 71.36, 71.11, 51.73, 51.18, 50.69, 50.66, 50.51, 50.14, 17.92, 17.85, 17.53, 17.43. HRMS (ESI): Calculated for C40H48N9O11 [M+H+]: 830.3473; observed: 830.3471.  59  Chapter 3: Application of Fluorogenic Substrates for Human Neutrophil Elastase 3.1 Background and Synopsis This chapter describes the application of the newly-designed and synthesized compounds 5 and 13 as fluorogenic substrates for Human Neutrophil Elastase. The compounds were first validated as HNE substrates in vitro, qualitatively by visual inspection and quantitatively by UV-vis spectroscopy. By the latter technique, Michaelis-Menten kinetic parameters were determined, thus revealing the relative specificity of HNE toward each substrate. Subsequently, the utility of these substrates in fabricating a point-of-care, bioactive paper-based sensor for wound monitoring was demonstrated. The resulting biosensor was tested by incubation with various amounts of enzyme to demonstrate concentration-dependent response. 3.1.1 Enzyme Kinetics Kinetic studies provide information on the substrate specificity of enzymes.55 During catalysis, an enzyme binds to different substrates with different affinities. Higher specificity for a substrate facilitates enzyme binding which ultimately increases the overall reaction velocity.  Once an enzyme has bound a substrate, one or more chemical steps occurs, leading to product formation and then release.  As the complexity of enzyme mechanisms increases, kinetic models can become highly sophisticated and challenging to derive. However, in many cases, and especially for hydrolytic enzymes like proteases, the standard Michaelis-Menten (MM) kinetic model is appropriate.  In the case of an irreversible, single-substrate enzymatic reaction,  product formation involves two steps (Equation 3.1): 1) the reversible binding between enzyme (E) and substrate (S) 60  to form the E-S complex, and 2) the irreversible chemical step and product release to regenerate free enzyme for the next round of catalysis.     (3.1) The Michaelis-Menten model is based on two assumptions: 1. The enzyme-substrate (E-S) complex is formed more quickly than the subsequent reaction, which converts the substrate molecules into the products. 56,57 2. The concentration of the E-S complex rapidly reaches a steady-state equilibrium concentration. 56,57   To study the kinetics of this reaction, it is necessary to consider the concentration of each species in each stage of reaction. Initially, the substrate is in large excess versus the enzyme, which rapidly form an E-S complex. This pre-steady state reaction generates a ‘burst’ in the complex which reaches saturation. This saturation brings about a steady rate of substrate depletion and product formation. Lastly, as substrate becomes depleted, the free enzyme concentration returns to the original level.  𝑣 =  𝑑[𝑃]𝑑𝑡=  𝑉𝑚𝑎𝑥[𝑆]𝐾𝑀+[𝑆]      (3.2) Under the steady-state approximation, the Michaelis-Menten equation (Equation 3.2), which relates initial reaction velocity to substrate concentration, can be derived. From the Michaelis-Menten equation three key parameters are used to consider enzyme efficiency: 1. The Michaelis constant, Km (M), defined as (k2+k-1)/k1, representing the likelihood of binding between the enzyme and the substrate leading to productive turnover.55 61  2. The maximum velocity, Vmax (M/s), is the velocity when all enzyme is in the E-S complex form. Vmax is dependent on the total enzyme concentration, [E]T (M), and the turnover rate of the enzyme, kcat, (s-1), i.e. Vmax=[E]T kcat.55 3. The ratio kcat/Km (unit in M-1s-1) reflects the specificity of an enzyme for a substrate and is valuable in comparing catalytic efficiency among substrates. To construct MM plot, the rate of product formation is essential to be evaluated at the various substrate concentrations. To evaluate Michaelis-Menten kinetic parameters is the initial-rate of the enzyme-catalyze reaction in determined in solution, typically by measuring product formation. Multiple initial-rate measurements, typically spanning the range 1/5 Km to 5 Km (Figure 3.1), are required to fully define the resulting Michaelis-Menten plot (Figure 3.2).      Figure 3.1 (Solid line) product(P) progress curves of enzymatic reactions with various substrate (S) concentration, (Dash line) Tangent line, representing the reaction velocities, of progress curves at initial substrate concentrations. Reprinted with the permission from John Wiley and Sons (58).  62   Figure 3.2 Michaelis Menten plot between initial velocity (V0) and substrate concentration ([S]). Reprinted with the permission from John Wiley and Sons (58).   3.1.2 Kinetic Model of Human Neutrophil Elastase The Michaelis-Menten model in section 3.1.1 explains the simplest case of an enzyme catalyzing a reaction of a single substrate to release a single product. The mechanism of the proteolysis catalyzed by HNE is formally a Ping-Ping Bi-Bi reaction, involving the peptide substrate, a covalent enzyme intermediate, and water (Figure 1.3, Figure 3.3A). Consisting of two major chemical steps, peptide-bond cleavage is initiated by the nucleophilic attack of Ser195 on the carbonyl carbon of the substrate, releasing the first N-terminal amine product, and forming an acyl-enzyme intermediate. Subsequent hydrolysis of the acyl-enzyme releases the second peptide product, regenerating the free enzyme.9 The large excess of water in the assay allows the simplification of the Ping-Pong Bi-Bi mechanism (Figure 3.3A) to the Uni-Bi mechanism (Figure 3.3B) by omitting the kinetic step corresponding to the binding of the water molecule.7,55 Furthermore, the release of both products from enzyme-product complex (EPNPC) is assumed to be relatively irreversible and rapid, due to their low concentrations under initial-rate conditions.  Typically, peptide bond cleavage of the enzyme-substrate complex (ES) is the rate determining step.7,55  63   Figure 3.3 (A) Ping-Pong Bi-Bi enzymatic mechanism of proteolytic reaction by HNE, (B) Uni Bi enzymatic mechanism. Reprinted with the permission from Springer Nature (7).    Thus, the Ping-Pong Bi-Bi mechanism (Figure 3.3A) results in an expansion of the original Michaelis-Menten kinetic model to include additional terms, 𝑣 =𝑉𝑀𝑎𝑥[𝑆][𝐻2𝑂]𝐾𝑚𝐻2𝑂[𝑆]+𝐾𝑚𝑆[𝐻2𝑂]+[𝑆][𝐻2𝑂].55 Because [H2O] is large and constant under practical assay conditions, this equation reduces to, 𝑣 =𝑉𝑀𝑎𝑥[𝑆]𝐾𝑚𝑆+[𝑆], for a Uni-Bi mechanism (Figure 3.3B), where 𝑉𝑀𝑎𝑥 = 𝑘𝑐𝑎𝑡[𝐸]𝑇, 𝑘𝑐𝑎𝑡 =  𝑘2𝑘3𝑘2+𝑘3 and 𝐾𝑚𝑆 =𝑘3(𝑘−1+𝑘2)𝑘1(𝑘2+𝑘3) .7 3.1.3 Spectrophotometry Compounds 5 and 13 were designed to enable the measurement of HNE activity, including the determination of Michaelis-Menten kinetic parameters, by either absorption or fluorescence spectroscopy.  Specifically, the rhodamine cores cRho110-PEG3-N3 (compound 1) and Rho110 (compound 6) have extended pi -conjugation system, including the two primary amine groups on the xanthene moiety. Reaction of these amine groups to form amide bonds, either to peptides or linkers, significantly alters the UV-visible and fluorescence spectroscopic properties, thereby providing a means to discriminate substrates and products in kinetic assays.  In considering the 64  assay design, the following parameters are important, which are based in the fundamental principles of absorption and fluorescence outlined in the Jablonski diagram (Figure 3.4): 1. The extinction coefficient (ε, expressed as M-1cm-1) is experimentally determined from the Beer-Lambert law and reflects the ability of a chromophore to absorb light in UV-vis spectroscopy. 2. Stokes’ law states that the emission wavelength of a fluorophore (λem) is always longer (lower in energy) than the wavelength of excitation (λex).  The Stokes shift is the difference λex and λem. (Figure 3.5)37  3. The quantum yield (Φ) is the ratio of the number of photons emitted to the number of photons absorbed. This parameter represents the fluorescence efficiency of a compound.  Figure 3.4 Jablonski diagram. Reprinted from Bioconjugate Techniques, 3rd, Greg T. Hermanson, Chapter 10: Fluorescent Probes, 396. Reprinted with the permission from Elsevier (37). 65   Figure 3.5 Stokes’ shift. Reprinted from Bioconjugate Techniques, 3rd, Greg T. Hermanson, Chapter 10: Fluorescent Probes, 397. Reprinted with the permission from Elsevier (37).  In this chapter, UV-visible spectroscopy (absorbance) was used to measure the hydrolysis of the new HNE substrates in solution to determine Michaelis-Menten parameters, and thus specificity, with reference to known HNE substrates.  On the other hand, fluorescence spectroscopy, using a fluorescence scanner, was used to quantify the HNE-catalyzed hydrolysis of the substrates after attachment to a paper substrate via click chemistry.    3.2 Materials and Methods 3.2.1 Solution-phase enzyme kinetics 3.2.1.1 Determination of extinction coefficients All absorbance measurements were performed by a UV-Vis spectrophotometer (Agilent Technologies Cary 60). Aqueous solutions of the compounds of interest (cRho110-PEG3-N3, compound 9, compound 5, and compound 13) at five or six different concentrations were prepared. The absorbance of the samples in the cuvettes was measured at the wavelength of 501 nm 35,42,48,59 in semi-micro cuvettes (1 cm × 0.4 cm). The Beer-Lambert law, A = εcl, (where ε is the extinction 66  coefficient, c is the concentration of solute, and 0.4 is the optical path length), was used to calculate the extinction coefficient of each compound in the linear region. 3.2.1.2 Preliminary verification of the activity of HNE on compound 5 25 µL HNE solution (8.33 µM in 0.1 M phosphate buffer, pH=7.6) was added to 0.19 ml of a 1 mg/ml solution of compound 5 in the same buffer at 37ºC and the sample was incubated for 10 min at 37 ºC.9 A control sample was prepared by addition of 60 µL 0.1 M phosphate buffer (pH=7.6) to 0.19 ml of 1 mg/ml solution of compound 5 in the same buffer. 3.2.1.3 Michaelis Menten kinetics of HNE on compound 5 and compound 13 Measurement of HNE kinetics on the rhodamine-based substrates was based on a procedure reported previously.12 All the reactions were performed in 1.5-ml semi-microcuvettes with the final reaction volume of 300 µL.  The HNE working solution was prepared by addition of 25 µL HNE stock solution (100 µg HNE in 100 µL of 50 mM sodium acetate and 150 mM NaCl, pH=5.5) into 975 µL of 0.1 M NaH2PO4, 0.5 M NaCl, pH=7.6 in a 1.5-ml Eppendorf tube. 8.75, 17.5, 35, 70, 140, 165, and 190 µL of 1 mg/ml stock solution of compound 5 in water were added to each cuvette to produce seven samples with different concentrations, 25.5, 51.0, 102.0, 204.1, 408.3, 481.2, and 554.1 µM respectively. 60 µL of the phosphate stock buffer (0.5 M NaH2PO4, 2.5 M NaCl, pH=7.6) was added to each cuvette. The cuvettes were incubated at 37 °C for 1 minute before addition of 50 µL HNE working solution. The measurement of HNE kinetics on compound 13 was executed in HEPEs buffer (25mM HEPES, 20 mM NaCl, pH7.4) at 37 °C. To 1.5-ml semi-microcuvettes with the final reaction volume of 300 µL, 2.2, 4.4, 8.8, 17.5, 35, 70, 95, and 123 µL of 2 mg/ml stock solution of compound 13 in water were added to each cuvette to produce seven samples with different 67  concentrations, 17.5, 35.2, 70.4, 140.7, 281.5, 562.9, 764.0 and 985.1 µM respectively. 30 µL of the HEPEs stock buffer (0.25M HEPES, 0.2 NaCl, pH7.4) was added to each cuvette. The cuvettes were placed in the UV-vis spectrometer and the UV-vis absorbance was recorded over time at the wavelength of 501 nm. The initial velocity (v0) of the reaction for each sample was calculated from the slope of the linear portion of the absorbance versus time plot, and the extinction coefficient. Origin Pro 10 60 was used to fit the Michaelis-Menten equation to plots of initial velocity divided by the concentration of the enzyme (V0/[E]T) versus the concentration of the substrate ([S]) using non-linear regression to derive  kcat (= Vmax/[E]T) and Km values. 3.2.2 Activation of cellulose paper surfaces and attachment of fluorophores 12.5 mg of (2,2,6,6-Tetramethylpiperidin-1-yl)oxyl (TEMPO) and 125 mg NaBr were dissolved in deionized water. To the reaction mixture, 1.5 ml of NaClO solution (13-15% chlorine) was added dropwise. 5M NaOH solution was employed to raise the pH of the reaction solution to 10. 1 g of 1-cm diameter Whatman No.1 filter papers - approximately 140 discs - were soaked in the reaction mixture for 5 min with gentle agitation, followed by washing with 40 ml deionized water (3 times) and 40 ml acetone (1 time) prior drying in reduced pressure.  0.5 g of TEMPO-oxidzed paper discs – approximately 70 discs - was suspended in 20-ml 1 M MES buffer (pH 4) in a 100-ml Erlenmeyer flask. EDC (0.24 g), NHS (0.14 g), and propargyl amine (0.5 g) were added, and the reaction mixture was agitated at 100 rpm for 24 h. The paper discs were washed with 40 ml deionized water 3 times, then once in 40 ml acetone and dried under reduced pressure. The resulting material was kept in a cool place and away from light. Per 14 mg of alkyne-functionalized paper – 2 discs, 0.5 ml of deionized water was used to soak the paper prior the addition of 25 μL of 0.9 mM (1 mg/ml) aqueous solution of compound 5 (final concentration 12.3 μM). A solution of THPTA/Cu(II)SO4 was prepared by mixing aqueous 68  solutions of THPTA (20 mg/ml) and Cu(II)SO4 (20 mg/ml) in the volume ratio of 5:1, respectively. To the reaction vial, 120 uL of THPTA/Cu(II)SO4 solution, and 1 mg of sodium ascorbate were added. Finally, an additional volume of deionized water was added to adjust the total reaction volume to 1.77 ml. The reaction vial was capped, wrapped with aluminium foil, and gently shaken (100 rpm). After 24 h, the discs were washed with 10 ml deionized water, 10 ml methanol, and 10 ml acetone before drying under reduced pressure.  The same procedure was used to modify paper surfaces with compound 13, using a stock solution of 1 mg/ml (1.2 mM), except that  two batches were made using different concentrations of compound 13, 12.3 and 49.2 μM in the CuAAC reaction by addition of 18.2 and 72.8 μL of the stock solution, respectively.  To enable quantitation of fluorophore release on paper surfaces, known amounts of the parent fluorophores were conjugated via CuAAC.  Stock solutions of 0.9-mM cRho110-PEG3-N3 in deionized water, 1.8-mM compound 9 in methanol, 20 mg/ml THPTA in deionized water, 20 mg/ml of Cu(II)SO4 in deionized water, and 2 mg/ml of sodium ascorbate in deionized water were prepared. A working solution of 5:1 v/v of THPTA solution and Cu(II)SO4 solution was prepared to minimize THPTA degradation and stored at -20°C.  3.2.3 CuAAC optimization using cRho110-PEG3-N3 and compound 9 Per 14 mg of alkyne-functionalized paper, 0.5 ml of water was mixed with 10, 25, 50, 100, 150 μL of the cRho110-PEG3-N3 or compound 9 stock solution in 20 ml sample vials. 120 μL of the 5:1 THPTA/Cu(II)SO4 solution and 0.5 ml of the sodium ascorbate solution were added respectively. Finally, deionized water was added to adjust the total reaction volume to 1.77 ml. After agitation at 100 rpm for 24 h, the paper discs were washed with 10 ml deionized water, 10 ml methanol, and 10 ml acetone, before drying under reduced pressure.  Prior to washing, the amount of unreacted cRho110-PEG3-N3 or compound 9, respectively, in each reaction was 69  determined using UV-vis spectroscopy (A501,  = 76,000 M-1cm-1 for cRho110-PEG3-N3,  = 23,164 M-1cm-1 for compound 9).  3.2.4 Calibration curve by absorption of cRho110-PEG3-N3 and compound 9 Whatman No.1 filter paper discs (10mm-diameter) were first hydrophobized to facilitate homogenous dispersion of flurophore solutions by soaking with polydimethylsiloxane (PDMS) and placement in a petri dish and covering with another larger piece of Whatman filter paper to absorb excess PDMS. The discs were then dried under reduced pressure overnight. Solutions of cRho110-PEG3-N3 and compound 9 of over a range of 1.7 μM and 366.7 μM were prepared. To individual PDMS-coated discs, 20 μL of solution was spread and dried repeated to obtain discs with defined masses of absorbed fluorophore. 3.2.5 Fluorescence imaging  A Bio-rad ChemiDoc XRS imaging system with Image Lab software was used to quantify fluorescence from paper surfaces, using the Alexa Fluor 488 setting (blue epi-illumination,530:28 filter). An unmodified Whatman no.1 filter paper was used as a reference. Images are presented with an inverted white background, with the maximum intensity of 10,000 and a gamma value of 0.60.  Calibration curves were constructed using the series of discs loaded with known amounts of the corresponding fluorophores by click chemistry or repeated absorbtion/evaporation. 3.2.6 Analysis of HNE activity on modified cellulose surfaces To monitor the ability of HNE to hydrolyze the fluorogenic compounds 5 and 13 covalently attached to cellulose paper surfaces, 10 μL of PBS buffer (0.1 M NaH2PO4 0.5 M NaCl pH 7.6) was used first to wet the paper discs on a glass petri dish. Serial dilutions of HNE were prepared (10, 5, 2.5 and 1.25 U/ml) and 10 μL of each solution was applied to individual discs to initiate reactions, followed by incubation at 37 °C. 20 μL of PBS was added every 30 min to keep the 70  paper wet, and fluorescence intensity was measured a discrete times over 6 h. An unmodified Whatman paper and substrate-functionalized discs without HNE added, wetted with PBS, were included as controls.  3.3 Results and Discussion 3.3.1 Validation of Compounds 5 and 13 as substrates for Human Neutrophil Elastase In an initial test, the bis-tetraalanine conjugate 5 was incubated with HNE and visualized under ambient ambient and UV light (254 nm).  As shown in Figure 3.6, compound 5 has no detectable fluorescence under UV illumination, whereas treatment with HNE results in strong fluorescence.  Similar results were observed with the mono-tetraalanine conjugate, compound 13 (data not shown).  These initial results provide compelling preliminary evidence that compounds 5 and 13 are viable HNE substrates, warranting further detailed kinetic evaluation.  Figure 3.6 Control solution (left cuvette) and the HNE treated compound 5 solution (right cuvette) under ambient (left panel) and UV light with the wavelength of 254 nm (right panel).  As a prerequisite to kinetic analysis, the extinction coefficients of compounds 5 and 13, and the corresponding HNE hydrolysis products, cRho110-PEG3-N3 and compound 9, respectively, were measured at 501 nm in aqueous solution.  Linear relationships between concentration and absorbance were observed over appropriate ranges for each compound (Figure 71  3.7).  The slopes of the fitted lines were used to obtain the following extinction coefficients:  cRho110-PEG3-N3, 72000 M-1cm-1 (reported by the supplier, 76000 M-1cm-1); Compound 5, 604 M-1cm-1; compound 9, 23,164 M-1cm-1; compound 13 is 387 M-1cm-1.  For both pairs of compounds, complete amidation strongly reduces absorbance at 501 nm.  Notably, mono-amidation of the rhodamine core with the ethyleneglycol linker in compound 9 also results in a reduction of absorbance versus the parent chromophore/fluorophore (Rhodamine 110 max 498,  81000 48,59).  Nonetheless, the large  relative to compound 13 represents a significant difference for kinetic analysis of HNE hydrolysis by spectrophotometry.  Figure 3.7 Extinction coefficients of fluorogenic substrates and their fluorescent precursors in dH2O   The Michaelis-Menten kinetics of HNE on compounds 5 and 13 were subsequently determined in initial-rate assays and compared against reported data for two standard, commercially available HNE substrates, MeO-Suc-AAPV-pNA and MeO-Suc-AAPV-AMC (Table 3.1). V0 vs. [S] plots for the HNE-catalyzed hydrolysis of compound 5 are shown in Figure 3.8.  Kinetic analysis of compound 5 was originally performed in PBS. However, limited solubility y = 604xR² = 0.9994y = 71935xR² = 0.9996y = 23164xR² = 0.9996y = 387xR² = 0.989801234560 0.0002 0.0004 0.0006 0.0008 0.001 0.0012 0.0014Absorbance per 1-cm pathlength (a.u.)Concentration (M)Compound 5cRho110-PEG3-N3Compound 9Compound 13Linear (Compound 5)Linear (cRho110-PEG3-N3)Linear (Compound 9)Linear (Compound 13)72  of compound 13 in PBS necessitated the use of HEPES-NaCl, so kinetics on compound 5 were also obtained in this buffer for direct comparison.  Notably, the values of the Michaelis-Menten constant (Km) for compounds 5 and 13 are very similar to those of the benchmark substrates (sub-millimolar), which suggests that there is no major impact of the chromophore on HNE binding affinity. Only MeO-Suc-AAPV-pNA, which contains the comparatively small chromogenic leaving group p-nitrophenylaniline, had a markedly higher kcat value among the substrates shown in Table 3.1. Although the kinetic data for compound 5 are complicated by the possibility of single and double cleavage to release cRho110-PEG3-N3, thus precluding more detailed dissection of the data, the overall results demonstrate that both compounds 5 and 13 are very competent substrates for HNE, on par with other widely used chromogenic substrates.   Figure 3.8 Michaelis-Menten curve of compound 5 (black) in PBS pH 7.6 at 37°C (grey) in HEPES pH 7.4 at 37°C and (red) compound 13 in HEPES pH 7.4 at 37°C toward HNE  73   Table 3.1 Michaelis-Menten kinetic parameters of HNE substrates Substrate kcat (s-1) Km (μM) kcat/Km (M-1·s-1) Condition Reference Compound 5 0.71±0.04 335±41 2120 25mM HEPES, 20 mM NaCl, pH7.4 at 37 °C  Procedure from (61)  1.51±0.09 232±28 6510 0.1M NaH2PO4, 0.5 M NaCl pH 7.6 at 37 °C Procedure from (12) Compound 13 3.56±0.37 400±93 8900 25mM HEPES, 20 mM NaCl, pH7.4 at 37 °C Procedure from (61) MeO-Suc-AAPV-AMC 3.3 290 11,000 10% DMSO in 0.05 M Tris, 0.5 M NaCl, 0.1 M CaCl2 pH 7.5 at 25 °C Reported in (62,63,64) MeO-Suc-AAPV-pNA 17 140 120,000 10% DMSO in 0.1 M HEPES 0.5 M NaCl pH 7.5 at 25 °C Reported in (62, 64)  It is, however, worth noting that both compounds 5 and 13 are marginally poorer substrates than the benchmark fluorogenic substrate, MeO-Suc-AAPV-AMC based on kcat/Km values.  This may be due, in part, to differences in peptide sequence between the substrates.  The active site of HNE is a hemispherical pocket (Figure 3.9), composed of Val190, Phe192, Ala231, Val216, Phe228, and a disulfide bridge, at the subsite P1.65,66 P1 has a high affinity toward small hydrophobic amino acids such as valine, cysteine, alanine, methionine, and isoleucine.65,66 Subsite P2 consists of Phe215, Leu99, as well as His57 (one of the residues of the catalytic triad), and demonstrates a preference for a medium-sized hydrophobic side chains, such as proline.65,66 Therefore, the replacement of proline with alanine in the P2 subsite in compounds 5 and 13 may contribute to the slightly lower substrate specificity of HNE for these substrates. 74    Figure 3.9 HNE’s catalytic triad, which is composed of His57, Asp102, and Ser195, covalently inhibited by N-[4-[(4-morpholinyl)carbonyl]benzoyl]peptidyl pentafluoroethyl ketone (blue stick). PDB ID: 1B0F. Cregge RJ, Durham SL, Farr RA, et al. J Med Chem. 1998;41(14):2461-2480.66  3.3.2 Production and analysis of paper-based biosensors for human neutrophil elastase. To produce HNE biosensors, the validated bis- and mono-tetraalanine HNE substrates (compounds 5 and 13, respectively) were conjugated onto alkyne- modified Whatman No.1 filter paper.  Whatman No.1 is made from cotton fiber, and therefore serves as a solid support analogous to cotton medical gauze.  Subsequently, HNE activity was observed and quantified by fluorescence imaging using calibration curves obtained for papers with defined amounts of the corresponding parent fluorophore.  75  3.3.2.1 Bis-tetraalanine-based HNE biosensor To fabricate the calibrated analytical biosensors, two sets of experiments were conducted (1) to quantify the number of molecules of fluorogenic substrates on paper surface, and (2) to correlate the number with the fluorescence intensity monitored by the fluorescence imager. In order to study the reactivity of CuAAC reaction on cellulosic surface, five initial concentrations of cRho110-PEG3-N3 (5-74 μM) were reacted with alkyne-functionalized Whatman No. 1 filter paper. Absorption at 501 nm before and after the reaction was used to quantify the amount of the fluorophore attached to the surface (Table 3.2).  Regardless of initial concentration CuAAC resulted in a conjugation yield of ca. 80%, with loadings in the range 3.3-48.4 nmol/disc.  The resulting paper discs were strongly colored (Figure 3.10A) and fluorescent under epi-illumination (Figure 3.10B).  Table 3.2 CuAAC reactivity of free cRho110-PEG3-N3 on propargyl amine coupled filter paper Reaction batch Volume of 0.9 mM compound 1 used (μL) Initial compound-1 quantity Final compound-1 quantity Reacted compound-1 quantity %Reacted compound 1 (nmol) (μM) (nmol) (μM) (nmol/disc)  1 10 8.7 4.9 2.2 1.2 3.3 74.6 2 25 21.8 12.3 4.8 2.7 8.5 77.9 3 50 43.5 24.6 9.6 5.4 17.0 77.9 4 100 87.0 49.2 15.7 8.9 35.7 81.9 5 150 130.5 73.4 33.8 19.0 48.4 74.1   76    Figure 3.10 cRho110-PEG3-N3 clicked onto propargyl amine coupled paper discs with various reacted cRho110-PEG3-N3 quantities (from left) virgin Whatman No.1 disc, 3.3, 8.5, 17.0, 65.7, 48.4 nmol/disc visualized under (A) white light, (B) a laboratory epi-illuminator with orange filter    To generate calibration curves for quantitation using a fluorescence scanner, direct absorption into the paper matrix had the advantage of being practically simpler, could potentially achieve higher loading, and was not dependent on chemical coupling yields. It was immediately observed that repeated application of a solution of cRho110-PEG3-N3, followed by repeated cycles of drying and addition to increase loading, led to uneven distribution of the fluorophore (Figure 3.11A).  Pre-treatment of the paper discs with PDMS mitigated this issue (Figure 3.11B).  This method allowed very high fluorophore loadings to be obtained (12.5-3750 ng/disc), and fluorescence was readily observed visually with epi-illumination and with a fluorescence scanner (Figure 3.11C and D).  Notably, loadings beyond the dynamic range of the scanner were obtained, with fluorescence quenching observed at the highest loading levels.    77   Figure 3.11 (A) cRho110-PEG3-N3 absorbed Whatman no.1 filter paper without PDMS coating, (B) with PDMS coating, (C & D) Different amount of  cRho110-PEG3-N3 absorbed paper discs (from left) 0, 12.5, 25, 37.5, 87.5, 125, 625, 1250, 2500, 3750 ng visualized under (C) a laboratory epi-illuminator with orange filter, (D) Bio-rad ChemiDoc XRS with Alexa Fluor 488 setting (excitation by epi-blue illumination 480-490 nm and 530/28 nm emission filter)   Figure 3.12B illustrates the calibration curve used to quantify HNE activity as a function of cRho110-PEG3-N3 fluorescence. The relationship between fluorescence and fluorophore concentration was linear only between the range of 0 – 0.65 nmol (Figure 3.12A), while higher concentration of the fluorophore caused quenching of the fluorescence signal. Based on these results we decided to use the linear relationship to convert measured fluorescence intensities measured from compound-5 modified paper discs to HNE activity level.  Figure 3.12 (A) Calibration curve of cRho110-PEG3-N3 absorption method, (B) Close-up calibration curve from cRho110-PEG3-N3 absorption method in the range of 0 to 0.65 nmol giving a straight line before saturation and quenching effect occur. 78  With a suitable set of standards for calibration in hand, the kinetics of HNE-catalyzed hydrolysis of the bis-tetraalanine substrate (compound 5) on paper discs was explored. A preliminary analysis using epi-illumination for visual detection clearly revealed increasing fluorescence intensity of discs that was directly proportional to HNE concentration over a fixed incubation time (Figure 3.13A and B).  Gratifyingly, quantitative analysis using a fluorescence scanner of a larger subsequent experiment clearly revealed a time- and dose-dependent response of the modified cellulose discs (Figure 3.13C and D).    Figure 3.13 (A) a laboratory epi-illuminator with orange filter utilized for preliminary observation of fluorescence phenomena after 6 hours of enzymatic reaction, (B) Close-up image of HNE incubated divalent fluorogenic substrate-clicked bioactive paper with different HNE loaded (from left) Whatman no.1 unmodified disc, 0, 21, 42, 83, 176 pmol of HNE, (C) Enzyme concentration- and time-dependent hydrolysis of compound 5 on paper surface observed by Bio-rad ChemiDoc XRS with Alexa Fluor 488 setting, (D) Time-dependent fluorescence intensity progress curves of different HNE loading discs up to 6 hours    79  To calculate the reaction progress, the initial concentration of compound 5 functionalized on one paper disc (Table 3.2) and the concentration of reacted substrate (Figure 3.13D) were measured. The reaction progress after 6 h from different HNE concentrations was found to be: 0.27 % for 21 pmol, 0.35% for 42 pmol, 0.66% for 83 pmol, and 0.94% for 176 pmol. Interestingly, and unlike previous studies using a comparable fluorescein-based esterase biosensor28, each applied amount of enzyme approached a different plateau value, implying that HNE was inactivated over time on the solid phase. Nonetheless, the data on the model substrate indicate that a functional biosensor had been produced using the design principals originally outlined.   3.3.2.2 Mono-tetraalanine-based HNE biosensor Analogous to the approach described above for the bis-tetraalanine biosensor based on cRho110-PEG3-N3, fluorescence calibration curves were constructed by direct absorption of compound 9 on paper surfaces for subsequent analysis of the mono-tetraalanine-based biosensor as well as the reactivity of compound 9 undergoing CuAAC. As before, the extent of covalent conjugation of 9 ( 23,164 M-1cm-1) was determined by absorbance in a subtractive assay, which showed a coupling yield of ca. 60%, regardless of initial fluorophore concentration (Table 3.3).  This value is ca. 10% lower than for the cRho110-PEG3-N3 fluorophore over a comparable concentration range (Table 3.2).      80  Table 3.3 CuAAC reactivity of compound 9 on propargyl amine coupled filter paper Reaction batch Volume of 1.8 mM compound 9 used (μL) Initial compound-9 quantity Final compound-9 quantity Reacted compound 9 % Reacted compound 9 (nmol) (μM) (nmol) (μM) (nmol/disc)  1 10 13.2 7.5 6.1 3.4 3.6 53.8 2 25 33.4 18.8 12.9 7.3 10.3 61.4 3 50 66.7 37.7 27.3 15.4 19.7 59.1 4 100 133.1 75.2 44.9 25.4 44.1 66.3 5 150 199.8 112.8 66.8 37.7 66.5 66.6   According to Figure 3.14A, at low loading of compound 9, a linear correlation between intensity and mass absorbed is observed in the range of 0 to 2.9 nmol. Unlike cRho110-PEG3-N3, the intensity begins to level off at 7.3 nmol without reduction due to quenching. The close-up linear region, shown in Figure 3.14B, can be utilized to correlate fluorescence intensity with free-compound 9 upon HNE cleavage of monovalent fluorogenic substrate, compound 13.  Figure 3.14 (A) Calibration curves of compound-9 absorption method, (B) Close-up calibration curve from compound-9 absorption method in the range of 0 to 2.9 nmol giving a straight line before saturation.  Subsequent kinetic studies of the HNE-catalyzed hydrolysis of compound 13 on paper surfaces, quantified by fluorescence scanning, revealed, notably, that this biosensor was 81  comparatively insensitive. For direct comparison with bis-tetraalanine fluorogenic substrate (compound 5), compound 13 was functionalized onto the paper discs with the same initial concentration of 12.3 μM (“Batch 1”). As shown in Figure 3.15A and B, the time- and enzyme concentration-dependent evolution of fluorescence was significantly lower than that shown in Figure 3.13.  Consequently, it was not possible to discriminate the activities of low concentrations of HNE (21-42 pmol). In this experiment, the lowest concentration of HNE practically detectable was 83 pmol, which was 4 times higher than the lowest concentration detected with the bis-tetraalanine-modified paper discs (Figure 3.13). Therefore, in an attempt to improve sensitivity, a 4-fold greater amount of compound 13 was conjugated onto the surface by increasing the initial concentration of the substrate in the CuAAC reaction (“Batch 2”). Gratifyingly, the enzymatic reaction progress curves increased significantly, evidencing a better sensitivity for HNE (Figure 3.15C and D) that was comparable or better than the bis-tetraalanine-modified paper discs (Figure 3.13). The number of functionalized compound-13 molecules was estimated as 44.1 nmol/ disc in Table 3.3. The extent of solid-phase reaction on paper discs were varied according to HNE loading (pmol) as following: 0.98% for 21 nmol, 1.30.% for 42 nmol, 1.67% for 83 nmol, and 1.86% for 176 nmol. Accordingly, the tuning of the degree of functionalization of the cellulose surface can be used to optimize sensitivity as needed, in balance with reagent consumption, in the generation of HNE biosensors. 82   Figure 3.15 Enzyme concentration- and time-dependent hydrolysis of compound-13 modified paper (A) batch 1 (with initial compound-13 concentration of 12.3 μM) and (C) batch 2 (with initial compound-13 concentration of 49.2 μM) observed by Bio-rad ChemiDoc XRS with Alexa Fluor 488 setting. Time-dependent progress curves of different HNE loading discs up to 6 hours from compound-13 modified papers (B) batch 1 and (D) batch 2              83  3.4 Summary In this chapter, the synthetic compounds 5 and 13 were validated as fluorogenic substrates for human neutrophil elastase. Incubation with the enzyme in solution produced fluorescent hydrolysis products, which could be observed visually and by UV-visible spectroscopy. Using the latter method, Michaelis-Menten parameters of these substrates were determined, which demonstrated that both compounds were competent substrates on par with widely used chromogenic and fluorogenic HNE substrates.  Ultimately, HNE biosenors were constructed using cotton cellulose paper surfaces as a substrate. To fabricate this biosensor, Whatman No. 1 filter paper was modified with TEMPO oxidation and propargyl amine coupling to produce an alkyne-activated surface, to which was coupled the azide-terminated fluorogenic substrates, compounds 5 and 13, via CuAAC. Subsequent quantitative analyses of time- and enzyme concentration-dependent evolution of fluorescence validated the modified paper discs as sensitive HNE biosensors. 84  Chapter 4: Conclusion and Future Works 4.1 Conclusion To conclude, this thesis ultimately illustrates a method for the successful fabrication of an HNE biosensor on a cellulose-based material, inspired by a previous project in the Brumer lab, in which a cellulosic medical materials were produced to detect  porcine liver esterase (PLE).28  The present work included the design of specific fluorogenic substrates that were characterized by 1H and 13C NMR to confirm successful synthesis. Using the Michaelis-Menten kinetic model, solution phase kinetic parameters were determined for HNE toward these fluorogenic substrate. Finally, a paper-based biosensor was constructed utilizing CuAAC as a key coupling reaction. In chapter 2, the design and synthetic procedures of fluorogenic substrates were investigated. Compound 5 was successfully designed and synthesized from cRho110-PEG3-N3, having a PEG-based linker terminated with an azide group for further CuAAC reaction with alkyne-functionalized paper. With a 4-step yield of 13%, two reactions were utilized: amino acid coupling by HATU, and Boc deprotection. Similarly, the rationale of the synthesis pathway and the synthesis procedure of compound 13 were illustrated with a 7-step yield of 2.7%. Characterization of the synthesized substrates were executed using one dimensional NMR, 1H and 13C NMR, with the assistance of 2D NMR techniques such as COSY, HSQC, and HMBC. HRESI-MS confirmed the molecular weight of compound 5 and compound 13. Chapter 3 explored the application of the synthesized substrates both in solution and solid phases. Incubation of the final synthetic products (described in Chapter 2) with HNE were validated as HNE fluorogenic substrates by the presence of the fluorescent by-products using UV-vis spectroscopy. Michaelis-Menten kinetic parameters, Km and kcat, were evaluated by initial-rate velocities. The novel fluorogenic compounds assayed had shown to have 60% lower turnover 85  numbers compared to commercially available substrates, indicating lower catalytic efficiencies. To develop higher-affinity clickable substrates, different tetrapeptide sequences could enhance overall interactions with HNE.   At the end of the project, we have developed a method to construct a bioactive paper modified by clickable HNE fluorogenic substrates. In brief, glucose-C6 hydroxyl groups from cellulose chains were activated via TEMPO oxidation. Alkyne groups were functionalized onto paper discs by operating propargyl amine coupling with the presence of EDC and NHS. As the final step of HNE paper-based biosensor fabrication, CuAAC was used to react the clickable substrates with alkyne-terminated paper. With several HNE-loading concentrations, the biosensor discs became fluorescent with the intensity depending on enzyme activity, which could be quantified by combining calibration curve data. It is worth noting that no spontaneous hydrolysis was observed, which implies that the amide linkages to the rhodamine core are generally stabile.  This research has provided a fundamental procedure to fabricate cellulose-based biosensors including substrate design towards a specific enzyme. The next section discusses further improvement that can be taken to enhance and optimize overall biosensor performance by 1) improving substrate affinity toward HNE, 2) optimizing the method for cellulose surface modification, and 3) fabricating an HNE dual-mode bio-responsive surface. 4.2 Future Work 4.2.1 Synthesis of HNE substrate library by SPOT synthesis As mentioned in section 1.2.2, HNE activity toward one substrate is regulated by the affinity of the enzyme for the tetrapeptide, which consequently affects the biosensor efficiency. However, in this case, selection of the tetrapeptide through a trial-and-error approach is impractical due to the, approximately, 16,000 combinatorial possibilities of natural tetrapeptide sequences.  86  SPOT synthesis can be successfully implemented for construction of a small scale tetrapeptide library, allowing for an array of reactions to operate on a cellulose surface in parallel as shown in Figure 4.1. Firstly, the paper is functionalized by either cRho110-PEG3-N3 or compound 9 into an array of circular spots, offering free amine groups on the surface. Secondly, the solid surface amine groups can be functionalized with amino acids by completing four cycles of coupling and deprotection with Fmoc, a technique often utilized in solid-phase peptide synthesis (SPPS). 67,68 A library of tetrapeptides can then be constructed to vary amino acids according to rows and columns, as shown in Figure 4.1B. As a result, a fluorescence intensity of each spot after the reaction with HNE to demonstrate the affinity of the enzyme towards a given tetrapeptide sequence. In vivo, chronic wounds not only consist of HNE but also matrix metalloproteases (MMPs) which play a role in extracellular matrix proteins degradation.69,70 Utilizing the tetrapeptide library array, specificity studies of MMPs can also be conducted to further optimize substrates.   Figure 4.1 (A) Molecular structure of free cRho110-PEG3-N3 functionalized as a 7-mm spot on a paper, (B) SPOT synthesis paper template that can vary 2 peptide variables at a time (distance between each spot’s center is ~8.5 mm). 87   4.2.2 Cellulosic surface chemistry exploration and development Carboxyl groups are introduced by TEMPO oxidation of hydroxyl groups on a cellulose surface, facilitating subsequent reactions to convert to other derivatives such as ester, as well as amide groups, illustrated in section 3.2.1. With the 3-step surface modification, HNE active paper-based biosensors can be successfully fabricated with room for additional research to optimize this process. The opportunity to explore other surface modification techniques can be widened by adjusting the substrate structure through protection of two tetrapeptide amine groups and hydrogenation of azide, as shown in Figure 4.2. Further development could be achieved from reactions with TEMPO-oxidized cellulose and amide bond coupling reagents, such as EDC and HATU, shown in Figure 4.3 in red. The direct amide formation between carboxyl group on TEMPO-oxidized cellulose surface and the amine terminal of the modified substrate (Figure 4.2B) consequently shortens the number of fabrication steps required to obtain the desired biosensing paper.    Figure 4.2 (A) Original divalent substrate and (B)modified divalent fluorogenic substrate with amine-terminated linker and succinylated tetra-alanine chains as HNE recognition modules  Alternatively, another potential route could involve activating surface hydroxyl groups on virgin cellulose with p-phenylene diisothiocyanate (PDITC), indicated in Figure 4.3 in blue. 88  PDITC would not only serve as an activator, but can also form a thiourea bond with the amine-terminated fluorogenic substrate at the other terminal. This method’s efficiency has been proven and employed by the previous research in Brumer lab.27 Utilization of divinylsulfone (DVS) in similar ways to PDITC has also been previously demonstrated in our lab.36 By exploring several surface modification techniques, the construction of the biosensor can be optimized to ultimately obtain a highly stable, low-cost production and low toxicity for realistic medical applications.  Figure 4.3 (red) Amide bond coupling of HNE substrate directly toward TEMPO oxidized paper; (blue) Thiourea formation of HNE substrate using PDITC as a surface activator, and (green) Secondary amine formation of HNE substrate using DVS as a surface activator.  4.2.3 Fabrication of an HNE dual-mode bio-responsive hydrogel Responsive materials, having dual functions in sensing and responding to an analyte, have been receiving much attention from material scientists. Considering the current HNE biosensing paper, tetrapeptide of compound 5 acts as an enzyme recognizing module which can be cleaved by HNE. The consequent parent fluorophore is fluorescent and is observable under UV light, detecting the presence of the enzyme. In order to upgrade this bioactive paper into a responsive cellulosic material, the concept of enzyme inhibition is introduced to covalently deactivate the target analyte. 89  To pursue this goal, a tetrapeptide-linked chloromethylketone (CMK) is known to irreversibly form covalent bonds with HNE to decelerate the degradation of extracellular matrix proteins.71 Correspondingly, the HNE clickable inhibitor (Figure 4.4 right) possesses tetrapeptide chain as a recognition module with N-terminal connected to azide linker for CuAAC reaction. Instead of fluorophore at the C-terminal (Figure 4.4 left), a substituting chloromethylketone module would act as a covalent inhibitor of HNE.  Interestingly, co-functionalization of both substrates and inhibitors onto alkyne-terminated cellulosic materials applies dual functions, detect-and-inhibit, onto a single material which can be fabricated in many formats, for example, cellulosic-based hydrogel, gauze, and cotton. Having coordinated multiple mechanisms for wound recovery, the innovative detect-and-inhibit hydrogel, potentially assisting growth of skin cells and initiating the inflammatory stage of skin healing, could be considered as a promising prospect for a responsive functional material.72  90   Figure 4.4 Schematic illustration of dual-mode bio-responsive cellulosic surface  In summary, two clickable peptide-conjugated fluorogenic substrates specific to HNE were successfully synthesized for the ultimate purpose of developing cellulose-based medical point-of-care diagnostic tools. The substrates were characterized with NMR and MS before testing the activity with the target enzyme. The molecules were designed to be compatible with the CuAAC click reaction, which is a powerful tool to efficiently construct functional materials. This research has proven the validity of molecular design, and cellulose-based biosensor fabrication procedure for further optimization and industrial application. 91   92  Bibliography  1.  Myers. BA. Wound Management: Principles and Practice (2nd Ed.). New Jersey, NJ: Pearson Education Inc.; 2008. 2.  OpenStax. Chapter 5: Integumentary system, Anatomy & Physiology. OpenStax CNX. cnx.org/contents/14fb4ad7-39a1-4eee-ab6e-3ef2482e3e22@8.24. Published 2016. Accessed September 7, 2019. 3.  Kabashima K, Honda T, Ginhoux F, Egawa G. The immunological anatomy of the skin. Nat Rev Immunol. 2019;19(1):19-30.  4.  Eming SA, Martin P, Tomic-Canic M. Wound repair and regeneration: Mechanisms, signaling, and translation. Sci Transl Med. 2014;6(265).  5.  Demidova-Rice TN, Hamblin MR, Herman I. Acute and impaired wound healing: pathophysiology and current methods for drug delivery, part 2: role of growth factors in normal and pathological wound healing: therapeutic potential and methods of delivery. Adv Ski wound care. 2012;25(8):304-314.  6.  Hiebert PR, Granville DJ. Granzyme B in injury, inflammation, and repair. Trends Mol Med. 2012;18(12):732-741. 7.  Brix K, Stöcker W. Proteases: Structure and Function. Vol 9783709108. Springer, Vienna; 2013.  8.  Chakraborti S, Dhalla NS. Pathophysiological Aspects of Proteases. (Chakraborti S, Dhalla NS, eds.). Springer Nature Singapore Pte Ltd.; 2017.  9.  Ohbayashi H. Current synthetic inhibitors of human neutrophil elastase. Expert Opin Ther Pat. 2002;12(1):65-84.  93  10.  Ferreira A V., Perelshtein I, Perkas N, Gedanken A, Cunha J, Cavaco-Paulo A. Detection of human neutrophil elastase (HNE) on wound dressings as marker of inflammation. Appl Microbiol Biotechnol. 2017;101(4):1443-1454.  11.  O’Donoghue AJ, Jin Y, Knudsen GM, et al. Global Substrate Profiling of Proteases in Human Neutrophil Extracellular Traps Reveals Consensus Motif Predominantly Contributed by Elastase. PLoS One. 2013;8(9):1-12.  12.  Kasperkiewicz P, Poreba M, Snipas SJ, et al. Design of ultrasensitive probes for human neutrophil elastase through hybrid combinatorial substrate library profiling. Proc Natl Acad Sci. 2014;111(7):2518-2523.  13.  Ali J, Najeeb J, Asim Ali M, Farhan Aslam M, Raza A. Biosensors: Their Fundamentals, Designs, Types and Most Recent Impactful Applications: A Review. J Biosens Bioelectron. 2017;08(01):1-9.  14.  Zhang Q, Lu Y, Li S, Wu J, Liu Q. Peptide-based biosensors. Pept Appl Biomed Biotechnol Bioeng. 2017;136:565-601.  15.  Jorfi M, Foster EJ. Recent advances in nanocellulose for biomedical applications. J Appl Polym Sci. 2015;132(14).  16.  Fu L, Zhang J, Yang G. Present status and applications of bacterial cellulose-based materials for skin tissue repair. Carbohydr Polym. 2013;92(2):1432-1442.  17.  Trache D. Nanocellulose as a promising sustainable material for biomedical applications. AIMS Mater Sci. 2018;5(2):208-214.  18.  Petersen N, Gatenholm P. Bacterial cellulose-based materials and medical devices: Current state and perspectives. Appl Microbiol Biotechnol. 2011;91(5):1277-1286.  19.  Bodin A, Ahrenstedt L, Fink H, Brumer H, Risberg B, Gatenholm P. Modification of 94  nanocellulose with a xyloglucan-RGD conjugate enhances adhesion and proliferation of endothelial cells: Implications for tissue engineering. Biomacromolecules. 2007;8(12):3697-3704.  20.  Fink H, Ahrenstedt L, Bodin A, et al. Bacterial cellulose modified with xyloglucan bearing the adhesion peptide RGD promotes endothelial cell adhesion and metabolism – a promising modification for vascular grafts. J Tissue Eng Regen Med. 2010;12(3):181-204.  21.  Pelton R. Bioactive paper provides a low-cost platform for diagnostics. TrAC - Trends Anal Chem. 2009;28(8):925-942.  22.  Martinez AW, Phillips ST, Whitesides GM, Carrilho E, Chem A. Diagnostics for the developing world: microfluidic paper-based analytical devices. Anal Chem. 2010;82(1):3-10.  23.  Ellerbee AK, Phillips ST, Siegel AC, et al. Quantifying colorimetric assays in paper-based microfluidic devices by measuring the transmission of light through paper. Anal Chem. 2009;81(20):8447-8452.  24.  Martinez AW, Phillips ST, Whitesides GM. Three-dimensional microfluidic devices fabricated in layered paper and tape. Proc Natl Acad Sci. 2008;105(50):19606-19611.  25.  Thuo MM, Martinez R V., Lan WJ, et al. Fabrication of low-cost paper-based microfluidic devices by embossing or cut-and-stack methods. Chem Mater. 2014;26(14):4230-4237.  26.  Martinez AW, Phillips ST, Nie Z, et al. Programmable diagnostic devices made from paper and tape. Lab Chip. 2010;10(19):2499-2504.  27.  Araújo AC, Song Y, Lundeberg J, Ståhl PL, Brumer H. Activated paper surfaces for the rapid hybridization of DNA through capillary transport. Anal Chem. 2012;84(7):3311-3317.  95  28.  Derikvand F, Yin DLT, Barrett R, Brumer H. Cellulose-Based Biosensors for Esterase Detection. Anal Chem. 2016;88(6):2989-2993.  29.  Song Y, Gyarmati P, Araújo AC, Lundeberg J, Brumer H, Staìšhl PL. Visual detection of DNA on paper chips. Anal Chem. 2014;86(3):1575-1582.  30.  Edwards JV, Gaston-Pierre S, Bopp AF, Goynes W. Detection of human neutrophil elastase with peptide-bound cross-linked ethoxylate acrylate resin analogs. J Pept Res. 2005;66(4):160-168.  31.  Edwards JV, Prevost N, French A, Concha M, DeLucca A, Wu Q. Nanocellulose-Based Biosensors: Design, Preparation, and Activity of Peptide-Linked Cotton Cellulose Nanocrystals Having Fluorimetric and Colorimetric Elastase Detection Sensitivity. Engineering. 2013;05(09):20-28.  32.  Edwards JV, Prevost N, Sethumadhavan K, Ullah A, Condon B. Peptide conjugated cellulose nanocrystals with sensitive human neutrophil elastase sensor activity. Cellulose. 2013;20(3):1223-1235.  33.  Edwards JV, Prevost NT, French AD, Concha M, Condon BD. Kinetic and structural analysis of fluorescent peptides on cotton cellulose nanocrystals as elastase sensors. Carbohydr Polym. 2015;116:278-285.  34.  Johnson AF, Struthers MD, Pierson KB, Mangel WF, Smith LM. Nonisotopic DNA Detection System Employing Elastase and a Fluorogenic Rhodamine Substrate. Anal Chem. 1993;65(17):2352-2359.  35.  De Cremer G, Roeffaers MBJ, Baruah M, et al. Dynamic disorder and stepwise deactivation in a chymotrypsin catalyzed hydrolysis reaction. J Am Chem Soc. 2007;129(50):15458-15459.  96  36.  Yu A, Shang J, Cheng F, et al. Biofunctional Paper via the Covalent Modification of Cellulose †. Langmuir. 2012;28(30):11265-11273.  37.  Hermanson GT. Bioconjugate Techniques. 3rd ed. Elsevier Inc; 2013.  38.  Saito T, Isogai A. TEMPO-mediated oxidation of native cellulose. The effect of oxidation conditions on chemical and crystal structures of the water-insoluble fractions. Biomacromolecules. 2004;5(5):1983-1989.  39.  Isogai A, Saito T, Fukuzumi H. TEMPO-oxidized cellulose nanofibers. Nanoscale. 2011;3(1):71-85.  40.  Kolb HC, Finn MG, Sharpless KB. Click Chemistry: Diverse Chemical Function from a Few Good Reactions. Angew Chemie - Int Ed. 2001;40(11):2004-2021.  41.  Castro V, Rodríguez H, Albericio F. CuAAC: An Efficient Click Chemistry Reaction on Solid Phase. ACS Comb Sci. 2016;18(1):1-14.  42.  Terentyeva TG, Van Rossom W, Van Der Auweraer M, Blank K, Hofkens J. Morpholinecarbonyl-rhodamine 110 based substrates for the determination of protease activity with accurate kinetic parameters. Bioconjug Chem. 2011;22(10):1932-1938.  43.  Valeur E, Bradley M. Amide bond formation: beyond the myth of coupling reagents. Chem Soc Rev. 2009;38(2):606-631.  44.  Carpino LA. 1-Hydroxy-7-azabenzotriazole. An Efficient Peptide Coupling Additive. J Am Chem Soc. 1993;115(10):4397-4398.  45.  Han SY, Kim YA. Recent development of peptide coupling reagents in organic synthesis. Tetrahedron. 2004;60(11):2447-2467.  46.  Sébastien Vidal, ed. Protecting Groups: Strategies and Applications in Carbohydrate Chemistry. 1st ed. Wiley-VCH Verlag GmbH & Co.; 2019.  97  47.  Albericio F, Isidro-llobet A, Mercedes A. Amino Acid-Protecting Groups. Chem Rev. 2009;109(6):2455-2504.  48.  Davis, Luke D, Chao, Tzu-Yuan, Raines RT. Fluorogenic label for biomolecular imaging. Bioconjug Chem. 2010;1(4):252-260.  49.  Wang J, Liang YL, Qu J. Boiling water-catalyzed neutral and selective N-Boc deprotection. Chem Commun. 2009;(34):5144-5146.  50.  Komatsu T, Hanaoka K, Adibekian A, et al. Diced electrophoresis gel assay for screening enzymes with specified activities. J Am Chem Soc. 2013;135(16):6002-6005.  51.  Lundt BF, Johansen NL, Vølund A, Markussen J. REMOVAL OF t‐BUTYL AND t‐BUTOXYCARBONYL PROTECTING GROUPS WITH TRIFLUOROACETIC ACID: Mechanisms, Biproduct Formation and Evaluation of Scavengers. Int J Pept Protein Res. 1978;12(5):258-268.  52.  López-Soria JM, Pérez SJ, Hernández JN, Ramírez MA, Martín VS, Padrón JI. A practical, catalytic and selective deprotection of a Boc group in N,N′-diprotected amines using iron(III)-catalysis. RSC Adv. 2015;5(9):6647-6651.  53.  Fulmer GR, Miller AJM, Sherden NH, et al. NMR chemical shifts of trace impurities: Common laboratory solvents, organics, and gases in deuterated solvents relevant to the organometallic chemist. Organometallics. 2010;29(9):2176-2179.  54.  Liu Y, Zhang Z, Zhang Q, Baker GL, Worden RM. Biomembrane disruption by silica-core nanoparticles: Effect of surface functional group measured using a tethered bilayer lipid membrane. Biochim Biophys Acta - Biomembr. 2014;1838(1 PARTB):429-437.  55.  Kenemans L, Ramsey N, Kenemans L, Ramsey N. Enzyme Kinetics: Principles and Methods. 3rd ed. Wiley-VCH Verlag GmbH & Co.; 2013.  98  56.  Stroberg W, Schnell S. On the estimation errors of KM and V from time-course experiments using the Michaelis–Menten equation. Biophys Chem. 2016;219:17-27.  57.  Duggleby RG, Clarke RB. Experimental designs for estimating the parameters of the Michaelis-Menten equation from progress curves of enzyme-catalyzed reactions. Biochim Biophys Acta (BBA)/Protein Struct Mol. 1991;1080(3):231-236.  58.  Nelson DL. Lehninger Principles of Biochemistry. 4th ed. (W.H. Freeman, ed.). New York; 2004. 59.  Leytus SP, Melhado LL, Mangel WF. Rhodamine-based compounds as fluorogenic substrates for serine proteinases. Biochem J. 1983;209(2):299-307.  60.  Origin Pro, Version 10. :OriginLabCorperation, Northampton, MA, USA. 61.  Fayad S, Nehmé R, Lafite P, Morin P. Assaying human neutrophil elastase activity by capillary zone electrophoresis combined with laser-induced fluorescence. J Chromatogr A. 2015;1419:116-124.  62.  Castillo MJ, Nakajima K, Zimmerman M, Powers JC. Sensitive substrates for human leukocyte and porcine pancreatic elastase: A study of the merits of various chromophoric and fluorogenic leaving groups in assays for serine proteases. Anal Biochem. 1979;99(1):53-64.  63.  Sun Q, Li J, Liu WN, Dong QJ, Yang WC, Yang GF. Non-peptide-based fluorogenic small-molecule probe for elastase. Anal Chem. 2013;85(23):11304-11311.  64.  Nakajima K, Powers JC, Ashe BM, Zimmermann M. Mapping the extended substrate binding site of cathepsin G and human leukocyte elastase. Studies with peptide substrates related to the (α1)-protease inhibitor reactive site. J Biol Chem. 1979;254(10):4027-4032. 65.  Korkmaz, Brice; Horwitz, Marshall.S.; Jenne, Dieter E.; and Gauthier F. Neutrophil 99  Elastase, Proteinase 3, and Cathepsin G as Therapeutic Targets in Human Diseases. Pharmacol Rev. 2010;62(04):726-759.  66.  Cregge RJ, Durham SL, Farr RA, et al. Inhibition of human neutrophil elastase. 4. Design, synthesis, X-ray crystallographic analysis, and structure-activity relationships for a series of P2-modified, orally active peptidyl pentafluoroethyl ketones. J Med Chem. 1998;41(14):2461-2480.  67.  Frank R. The SPOT-synthesis technique. J Immunol Methods. 2002;267(1):13-26.  68.  Hilpert K, Winkler DFH, Hancock REW. Peptide arrays on cellulose support: SPOT synthesis, a time and cost efficient method for synthesis of large numbers of peptides in a parallel and addressable fashion. Nat Protoc. 2007;2(6):1333-1349.  69.  Blakytny R, Jude E. The molecular biology of chronic wounds and delayed healing in diabetes. Diabet Med. 2006;23(6):594-608.  70.  Harding KG, Morris HL, Patel GK. Clinical review Healing chronic wounds. 2002;324(January).  71.  Navia MA, Mckeever BM, Springer JP, et al. Structure of Human Neutrophil Elastase in Complex with a Peptide Chloromethyl Ketone inhibitor at 1.84 angstrom Resolution. Proceeding of the National Academy of Sciences of the United States of America,1989, 86(1):7-11. 72.  Chen G, Yu Y, Wu X, Wang G, Ren J, Zhao Y. Bioinspired Multifunctional Hybrid Hydrogel Promotes Wound Healing. Adv Funct Mater. 2018;28(33):1-10.    100  101  Appendices  Appendix A  : Selected NMR Spectra A.1 Compound 5 Intermediates  Figure A1: 1H NMR spectrum of 2 in acetone-d6. 102   Figure A2: 13C NMR spectrum of 2 in acetone-d6.  Figure A3: 2D-COSY spectrum of 2 in acetone-d6. 103   Figure A4: 2D-HSQC spectrum of 2 in acetone-d6.   Figure A5: 2D-HMBC spectrum of 2 in acetone-d6. 104   Figure A6: 2D-HMBC spectrum of 2 in acetone-d6.   Figure A7: 2D-HMBC spectrum of 2 in acetone-d6.  105   Figure A8: 2D-HMBC spectrum of 2 in acetone-d6.   Figure A9: 1H NMR spectrum of 3 in MeOD-d4. 106   Figure A10: 13C NMR spectrum of 3 in MeOD-d4.  Figure A11: 1H NMR spectrum of 4 in acetone-d6. 107   Figure A12: 13C NMR spectrum of 4 in acetone-d6.  Figure A13: 2D-COSY spectrum of 4 in acetone-d6. 108   Figure A14: 2D-HSQC spectrum of 4 in acetone-d6.  Figure A15: 2D-HMBC spectrum of 4 in acetone-d6. 109   Figure A16: 2D-HMBC spectrum of 4 in acetone-d6.  Figure A17: 2D-HMBC spectrum of 4 in acetone-d6. 110   Figure A18: 2D-HMBC spectrum of 4 in acetone-d6. A.2 Compound 5  Figure A19: 1H-NMR spectrum of 5 in MeOD-d4. 111   Figure A20: 13C-NMR spectrum of 5 in MeOD-d4.  Figure A21: 2D-COSY spectrum of 5 in MeOD-d4. 112   Figure A22: 2D-COSY spectrum of 5 in MeOD-d4.  Figure A23: 2D-COSY spectrum of 5 in MeOD-d4. 113   Figure A24: 2D-HSQC spectrum of 5 in MeOD-d4.  Figure A25: 2D-HSQC spectrum of 5 in MeOD-d4. 114   Figure A24: 2D-HSQC spectrum of 5 in MeOD-d4.  Figure A25: 2D-HMBC spectrum of 5 in MeOD-d4. 115   Figure A26: 2D-HMBC spectrum of 5 in MeOD-d4.  Figure A27: 2D-HMBC spectrum of 5 in MeOD-d4. 116   Figure A28: 2D-HMBC spectrum of 5 in MeOD-d4.  Figure A29: 2D-HMBC spectrum of 5 in MeOD-d4. 117   Figure A30: 2D-HMBC spectrum of 5 in MeOD-d4.  Figure A31: 2D-HMBC spectrum of 5 in MeOD-d4. 118  A.3 Boc-tri-alanine-OH  Figure A32: 1H-NMR spectrum of Boc-AAA-OH in MeOD-d4.  Figure A33: 13C-NMR spectrum of Boc-AAA-OH in MeOD-d4. 119  A.4 Compound 7  Figure A34: 1H-NMR spectrum of 7 in acetone-d6.  Figure A35: 13C-NMR spectrum of 7 in acetone-d6.  120  A.5 Compound 8  Figure A36: 1H-NMR spectrum of 8 in acetone-d6.  Figure A37: 13C-NMR spectrum of 8 in acetone-d6. 121   Figure A38: 2D-COSY spectrum of 8 in acetone-d6.  Figure A39: 2D-COSY spectrum of 8 in acetone-d6. 122   Figure A40: 2D-HSQC spectrum of 8 in acetone-d6.  Figure A41: 2D-HSQC spectrum of 8 in acetone-d6. 123   Figure A42: 2D-HSQC spectrum of 8 in acetone-d6.  Figure A43: 2D-HMBC spectrum of 8 in acetone-d6. 124   Figure A44: 2D-HMBC spectrum of 8 in acetone-d6.  Figure A45: 2D-HMBC spectrum of 8 in acetone-d6. 125   Figure A46: 2D-HMBC spectrum of 8 in acetone-d6. A.6 Compound 9  Figure A47: 1H-NMR spectrum of 9 in DMSO-d6. 126   Figure A48: 13C-NMR spectrum of 9 in DMSO-d6  Figure A49: 2D-COSY spectrum of 9 in DMSO-d6. 127   Figure A50: 2D-COSY spectrum of 9 in DMSO-d6.  Figure A51: 2D-HSQC spectrum of 9 in DMSO-d6. 128   Figure A52: 2D-HSQC spectrum of 9 in DMSO-d6.  Figure A53: 2D-HSQC spectrum of 9 in DMSO-d6.  129   Figure A54: 2D-HMBC spectrum of 9 in DMSO-d6.  Figure A55: 2D-HMBC spectrum of 9 in DMSO-d6. 130   Figure A56: 2D-HMBC spectrum of 9 in DMSO-d6. A.7 Compound 11  Figure A57: 1H-NMR spectrum of 11 in acetone-d6. 131   Figure A58: 13C-NMR spectrum of 11 in acetone-d6.  Figure A59: 2D-COSY spectrum of 11 in acetone-d6. 132   Figure A60: 2D-COSY spectrum of 11 in acetone-d6.  Figure A61: 2D-COSY spectrum of 11 in acetone-d6. 133   Figure A62: 2D-HSQC spectrum of 11 in acetone-d6.  Figure A63: 2D-HSQC spectrum of 11 in acetone-d6. 134   Figure A64: 2D-HSQC spectrum of 11 in acetone-d6.  Figure A65: 2D-HMBC spectrum of 11 in acetone-d6. 135   Figure A66: 2D-HMBC spectrum of 11 in acetone-d6.  Figure A67: 2D-HMBC spectrum of 11 in acetone-d6. 136   Figure A68: 2D-HMBC spectrum of 11 in acetone-d6. A.8 Compound 13  Figure A69: 1H-NMR spectrum of 13 in MeOD-d4. 137   Figure A70: 13C-NMR spectrum of 13 in MeOD-d4.  Figure A71: 13C-DEPT NMR spectrum of 13 in MeOD-d4. 138   Figure A72: 2D-COSY spectrum of 13 in MeOD-d4.  Figure A73: 2D-COSY spectrum of 13 in MeOD-d4. 139   Figure A74: 2D-COSY spectrum of 13 in MeOD-d4.  Figure A75: 2D-HSQC spectrum of 13 in MeOD-d4. 140   Figure A76: 2D-HSQC spectrum of 13 in MeOD-d4.  Figure A77: 2D-HSQC spectrum of 13 in MeOD-d4. 141   Figure A78: 2D-HSQC spectrum of 13 in MeOD-d4.  Figure A79: 2D-HMBC spectrum of 13 in MeOD-d4. 142   Figure A80: 2D-HMBC spectrum of 13 in MeOD-d4.  Figure A81: 2D-HMBC spectrum of 13 in MeOD-d4. 143   Figure A82: 2D-HMBC spectrum of 13 in MeOD-d4.  Figure A83: 2D-HMBC spectrum of 13 in MeOD-d4. 144   Figure A84: 2D-HMBC spectrum of 13 in MeOD-d4. A.9 Linker (N3-PEG3-CH2COOH)  Figure A85: 1H-NMR spectrum of Linker (N3-PEG3-CH2COOH) in chloroform-d1. 

Cite

Citation Scheme:

        

Citations by CSL (citeproc-js)

Usage Statistics

Share

Embed

Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                        
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            src="{[{embed.src}]}"
                            data-item="{[{embed.item}]}"
                            data-collection="{[{embed.collection}]}"
                            data-metadata="{[{embed.showMetadata}]}"
                            data-width="{[{embed.width}]}"
                            data-media="{[{embed.selectedMedia}]}"
                            async >
                            </script>
                            </div>
                        
                    
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:
https://iiif.library.ubc.ca/presentation/dsp.24.1-0389685/manifest

Comment

Related Items