Applications of Oligopeptide-Functionalized

APPLICATIONS OF OLIGOPEPTIDE-FUNCTIONALIZED

SURFACES AND MICROPARTICLES IN LIQUID CRYSTALS-

BASED ASSAYS

ONG LIAN HAO

(B.ENG. (HONS.), NANYANG TECHNOLOGICAL UNIVERSITY)

A THESIS SUBMITTED

FOR THE DEGREE OF DOCTOR OF PHILOSOPHY DEPARTMENT OF CHEMICAL AND BIOMOLECULAR ENGINEERING NATIONAL UNIVERSITY OF SINGAPORE

2016

DECLARATION

I hereby declare that this thesis is my original work and it has been written by me

in its entirety. I have duly acknowledged all the sources of information which

have been used in the thesis.

This thesis has also not been submitted for any degree in any university

previously.

Ong Lian Hao 10 September 2016

ACKNOWLEDGEMENTS

First of all, I am utmost grateful for the opportunity presented by my advisor, Dr.

Yang Kun-Lin, to complete this thesis under his guidance. His insightful perspectives and open-door policy provided a conducive environment for me to develop and hone my research skills and ideologies.

I am also thankful for the engaging, and more importantly enlightening discussions with our group members, and they include Dr. Liu Fengli, Dr. Chen Chih-Hsin, Dr.

Ding Xiaokang, Ms. Liu Yanyang, Ms. Nguyen Le Truc and Dr. Arumugam

Kamayan Rajagopalan G. I am also fortunate to receive guidance and advice from

Dr. Saif Khan and his group members during our regular intergroup sharing sessions. I would also like to thank the friendly laboratory staff for their assistance in the procurement and operation of equipment, and they include Mr. Boey Kok

Hong, Ms. Lee Chai Keng, Mr. Liu Zhicheng, Ms. Lim Kwee Mei, Dr. Yuan Ze

Liang and Dr. Yang Liming.

Finally, I am grateful to my parents and parents-in-law for their support. I would also like to express my heartfelt gratitude towards my wife, Rylia, for her relentless support and dedicated care for the family over the years. She has always stood by me, and with our beautiful son, Jophiel, they are my motivation to complete this fulfilling life chapter.

TABLE OF CONTENTS

DECLARATION ...... i

ACKNOWLEDGEMENTS ...... ii

TABLE OF CONTENTS ...... iii

SUMMARY ...... x

LIST OF TABLES ...... xiii

LIST OF FIGURES ...... xiv

LIST OF SCHEMES...... xviii

LIST OF SYMBOLS ...... xix

LIST OF ABBREVIATION ...... xx

CHAPTER 1 INTRODUCTION ...... 1

1.1 Background ...... 2

1.1.1 Importance of Proteases and Protease Assays ...... 2

1.1.2 Introduction of Liquid Crystals ...... 4

1.1.3 LC-Based Protease Assays ...... 7

1.1.4 Immobilization of Oligopeptide on LC-Compatible Surfaces ...... 8

1.1.5 Signal Enhancement in LC-Based Protease Assays ...... 10

1.1.6 Oligopeptide-Functionalized Microparticles in LC ...... 11

1.1.7 Real-Time Protease Activity Monitoring...... 12

1.2 Research Objective ...... 13

CHAPTER 2 LITERATURE REVIEW ...... 16

2.1 Introduction ...... 17

iii

2.2 Homogenous Protease Assays ...... 20

2.2.1 Organic FRET Donors and Acceptors ...... 21

2.2.2 Nanomaterials as FRET Acceptors or Donors ...... 22

2.2.3 QD as FRET Donors ...... 24

2.2.4 Gold Nanoclusters (AuNC) ...... 26

2.2.5 Nanoparticles in Colorimetric Assays ...... 27

2.3 Heterogeneous Protease Assays ...... 29

2.3.1 Electrochemical Assays ...... 30

2.3.2 Label-Free SERS and SPR Assays ...... 33

2.3.3 Enzyme-Linked Peptide Protease Assays ...... 33

2.3.4 LC-Based Assays ...... 35

2.4 Outlook ...... 40

CHAPTER 3 MECHANISTIC STUDY FOR IMMOBILIZATION OF CYSTEINE-LABELED OLIGOPEPTIDES ON UV-ACTIVATED SURFACES ...... 42

3.1 Introduction ...... 43

3.2 Experimental Section ...... 46

3.2.1 Materials ...... 46

3.2.2 Surface Modifications ...... 46

3.2.3 Immobilization of Oligopeptides ...... 47

3.2.4 X-ray Photoelectron Spectroscopy (XPS) ...... 47

3.2.5 Stability Test for Immobilized Oligopeptides ...... 48

3.2.6 Purpald Test ...... 48

3.2.7 Streptavidin Binding Assay ...... 49

3.2.8 Oxidation of Thiols ...... 50

3.2.9 Fluorescence Microscopy ...... 50

3.2.10 Matrix-Assisted-Laser-Desorption-Ionization Time-of-Flight Mass-

Spectrometry (MALDI-TOF-MS) ...... 51

3.2.11 DTNB Assay ...... 51

3.3 Results and Discussion ...... 51

3.3.1 Role of Cysteine in Oligopeptide Immobilization ...... 51

3.3.2 Effects of Reducing Agent and Buffers ...... 54

3.3.3 Effect of UV Irradiation ...... 55

3.3.4 Test for Surface Aldehyde Group ...... 57

3.3.5 Role of Cysteine in Immobilization Mechanism ...... 58

3.3.6 Stability and Orientation of Oligopeptide Immobilization ...... 62

3.4 Conclusion ...... 65

CHAPTER 4 LIQUID CRYSTALS-BASED PROTEASE ASSAYS WITH DUAL CYSTEINE-LABELED PEPTIDES FOR IMPROVING PEPTIDE STABILITY ...... 66

4.1 Introduction ...... 67

4.2 Experimental Section ...... 71

4.2.1 Materials ...... 71

4.2.2 Surface Modifications ...... 71

4.2.3 Immobilization of Peptides ...... 72

4.2.4 Fluorescence Protease Assay ...... 73

4.2.5 LC-Based Protease Assay ...... 73

4.3 Results and Discussion ...... 74

4.3.1 Effect of Dual Cysteine on Peptide Surface Density ...... 74

4.3.2 Fluorescence Protease Assays ...... 78

4.3.3 Detection of Immobilized Peptides Using LC ...... 81

4.3.4 LC-Based Protease Assays ...... 82

4.3.5 Limit of Detection (LOD) of LC-Based Protease Assays ...... 85

4.4 Conclusion ...... 88

CHAPTER 5 DISTORTION OF LIQUID CRYSTAL PHASE BY MICROPARTICLES FUNCTIONALIZED WITH OCTADECYL- TERMINATED OLIGOPEPTIDES ...... 90

5.1 Introduction ...... 91

5.2 Experimental Section ...... 94

5.2.1 Materials ...... 94

5.2.2 Surface Modifications ...... 95

5.2.3 Immobilization of Oligopeptides on PEI-coated Microparticles .... 95

5.2.4 Fabrication of LC Cells ...... 96

5.2.5 Protease Assay ...... 96

5.3 Results and Discussion ...... 97

5.3.1 Interactions of LC with Submerged Microparticles...... 97

5.3.2 Effect of Surface Density of Oligopeptide on LC Distortion ...... 101

5.3.3 Effect of Cell Thickness on LC Distortion Diameter ...... 102

5.3.4 Computational Model for LC Director Profile ...... 104

5.3.5 Effect of Oligopeptide Surface Density on Computed Distortion

Radius ...... 105

5.3.6 Effect of LC Cell Thickness on Computed Distortion Radius ..... 107

5.3.7 Application of Oligopeptide-Functionalized Microparticles

Submerged in LC as a Trypsin Assay ...... 110

5.4 Conclusion ...... 111

CHAPTER 6 SURFACTANT-DRIVEN ASSEMBLY OF POLY(ETHYLENIMINE)-COATED MICROPARTICLES AT THE LIQUID CRYSTAL/WATER INTERFACE ...... 112

6.1 Introduction ...... 113

6.2 Experimental Section ...... 117

6.2.1 Materials ...... 117

6.2.2 Surface Modifications ...... 117

6.2.3 Coating of Microparticles with PEI ...... 118

6.2.4 Labeling of PEI-Coated Microparticles with FTIC ...... 118

6.2.5 Zeta Potential Measurements of Microparticles ...... 118

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6.2.6 Microparticle Assembly at LC/Water Interfaces ...... 119

6.2.7 Visual Detection of Lipase Activity Using Microparticle Assembly .

...... 120

6.3 Results and Discussion ...... 121

6.3.1 Surface Coating of Microparticles ...... 121

6.3.2 Adsorption of Surfactants on PEI-Coated Microparticles ...... 122

6.3.3 Self-Assembly of PEI-Coated Microparticles at LC/Water Interfaces

...... 123

6.3.4 Effect of Tween 20 Concentration on Microparticle Assembly ... 126

6.3.5 Visual Detection of Lipase Activity in Water ...... 129

6.3.6 Effects of Cationic and Anionic Surfactants ...... 130

6.3.7 Effect of SDS Concentration on Microparticle Assembly ...... 133

6.4 Conclusion ...... 135

CHAPTER 7 CONCLUSIONS AND RECOMMENDATIONS ...... 137

7.1 Conclusions ...... 138

7.2 Recommendations ...... 141

BIBLIOGRAPHY ...... 146

APPENDICES ...... 172

Appendix A ...... 172

A.1 Effect of Coating Duration on Water Contact Angle of DMOAP-

Coated Surface ...... 172

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A.2 Effect of UV Irradiation on Immobilization of P2 and P3 ...... 174

A.3 Determination of Oxidation of Free Thiols ...... 175

Appendix B ...... 176

B.1 Surface Amine Density Calculations ...... 176

LIST OF PUBLICATIONS ...... 177

SUMMARY

Proteases play pivotal roles in a myriad of biological processes, ranging from food digestion to cancer metastasis. Such importance has spurred great interest in the development of protease assays. In particular, liquid crystals- (LC-) based assays are attractive due to its sensitive and convenient detection format. However, several key challenges remained. First, stable immobilization of oligopeptides on LC- compatible surfaces is lacking. Second, detection of minute changes in peptides upon cleavage is challenging. Moreover, the limited surface area has led to poor detection limits. Finally, most assay formats are incapable of quantitative and real time monitoring. In this thesis, we addressed these challenges in the development of LC-based protease assays.

Stable immobilization of oligopeptides on LC-compatible surfaces is imperative for giving consistent assay results. In Chapter 3, we studied the immobilization of cysteine-labeled oligopeptides on ultraviolet- (UV)-activated, LC-compatible surfaces. Our results show that cysteine residue, regardless of its position in the oligopeptide, is essential for the stable oligopeptide immobilization. A possible reaction mechanism is the nucleophilic addition of thiolates to surface aldehyde groups. By using this technique, we were able to incorporate specific anchoring points into oligopeptides through cysteine residues. Furthermore, immobilized oligopeptides are very stable under harsh washing conditions. Therefore, this stable oligopeptide immobilization technique is employed in this thesis.

In Chapter 4, we further improved the stability of immobilized peptides and developed strategies for detecting minute changes in LC-based protease assays. The main concept is to use dual cysteine and biotin-labeled peptides as protease substrates. The use of dual cysteine label increases surface peptide density due to the formation of inter-peptide disulfide bonds. Meanwhile, the biotin moiety offers a streptavidin binding site for the amplification of changes in surface properties after protease cleavage. This approach led to better detection limit compared to previously-published LC-based protease assays.

For LC-based assays on flat surfaces, interactions between oligopeptides and LC are limited. Therefore, functionalized microparticles with a high surface area were used to immobilize octadecyl- (C18)-terminated oligopeptides for detecting proteases as shown in Chapter 5. Upon cleavage of oligopeptides by proteases, drastic changes in LC orientation near the microparticles (from homeotropic to planar) result in observable optical signals. This is due to the removal of the C18 fragment, which is responsible for the homeotropic LC orientation. This phenomenon can be potentially exploited for the development of high-throughput and quantitative protease assays.

In Chapter 6, self-assembly of functional microparticles at the LC/water interfaces were investigated. Because microparticles are in constant interaction with both the

LC and water phases, this system can be potentially useful for real time monitoring.

When polyethylenimine- (PEI)-coated microparticles were used, they were able to

xi attract surfactants such as Tween 20 and sodium dodecyl sulfate (SDS) in the aqueous solutions and self-assembled into unique patterns such as short chains and circular rings, respectively. Interestingly, the addition of ≥100 ng/ml lipase led to the hydrolysis of Tween 20, and this disrupted the formation of short chains. This result shows a promising application of microparticles for detecting protease activity in real time.

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LIST OF TABLES

Table 2.1. Summary of Homogeneous Trypsin and Chymotrypsin Protease Assays Recently Developed ...... 28

Table 2.2. Summary of Heterogeneous Trypsin and Chymotrypsin Protease Assays Recently Developed ...... 40

Table 3.1. Sequences of Model Oligopeptides ...... 53

Table 4.1. Peptide Sequences ...... 76

Table 4. 2. Comparison of Trypsin and Chymotrypsin Assays Recently Developed ...... 88

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LIST OF FIGURES

Figure 3.1. (a) The structure of DMOAP. (b) Proposed surface structure presenting aldehyde groups after UV irradiation...... 45

Figure 3.2. Fluorescence image of oligopeptides immobilized on UV irradiated DMOAP-coated surface using different buffer solutions: (a) carbonate buffer (100 mM, pH 10), (b) carbonate buffer (100 mM, pH 10) with 10 mM sodium cyanoborohydride, (c) PBS buffer (100 mM, pH 7); spots from left to right correspond to P1 to P5. (d) Comparison of fluorescence data for oligopeptide immobilization using the three different buffers in (a) to (c)...... 53

Figure 3.3. Fluorescence intensity of immobilized oligopeptide for different UV exposure durations: (a) P1, (b) P4 and (c) P5...... 56

Figure 3.4. XPS results of DMOAP-coated glass surface illustrating the emergence of C (1s) shoulder peaks with increasing (254 nm) UV exposure duration: (a) 0 s, (b) 50 s, (c) 180 s and (d) 300 s...... 56

Figure 3.5. Purpald test results: (a) 50 s UV irradiated DMOAP-coated TLC plate, (b) no UV irradiation on DMOAP coated TLC plate...... 57

Figure 3.6. Fluorescence intensities for streptavidin binding assay used to examine the role of cysteine in oligopeptide immobilization: (a) effect of thiol oxidation, (b) effect of pH...... 59

Figure 3.7. Mass spectrometry results for the product of oxidation in DMSO at different durations: (a) 0 h, (b) 1 h, (c) 2 h, (d) 3 h and (e) 4 h...... 61

Figure 4.1. (a) Fluorescence image of a surface with immobilized P1 (top) and P2 (bottom) at different concentrations (in μM). (b) Effects of DTT on fluorescence intensities of P1 immobilized on the surface without DTT (solid diamonds) and with 100 mM of DTT (hollow diamond) or P2 immobilized on the surfaces without DTT (solid circles) and with 100 mM of DTT (hollow circle). Scale bar represents 1 mm...... 76

Figure 4.2. Fluorescence trypsin assay performed on (a-d) P1- and (e-h) P2- functionalized surfaces. Increasing trypsin concentration led to decreasing fluorescence intensity of the immobilized peptide. Surface density of peptide was controlled by using different concentrations of peptides. For P1 and P2, concentrations of peptides used were (a,e) 500, (b,f) 250, (c,g) 100, and (d,g) 10 μM. For clarity, error bars were omitted. Typical standard deviation is ± 7% and ± 10% of mean intensities for P1- and P2-functionalized surfaces, respectively. ... 80

xiv by using different concentrations of P2 (10, 100, and 500 μM from left to right) during peptide immobilization. Scale bar represents 1mm...... 82

Figure 4.4. LC-based protease assays for the detection of trypsin and chymotrypsin using P2- and P3-fucntionalized surfaces. Concentrations of P2 and P3 used in immobilization were 100 μM (first column) and 500 μM (second column). The P2- functionalized surfaces were used to detect (a) 250 ng/ml of trypsin and (b) 500 ng/ml of chymotrypsin, respectively. The P3-functionalized surfaces were used to detect (c) 250 ng/ml of chymotrypsin and (d) 500 ng/ml of trypsin, respectively. Scale bar represents 1mm...... 83

Figure 4.5. Limit of detections observed in the LC-based trypsin assays by using P2-functionalized surfaces. The first and second columns represent surfaces functionalized by using 100 and 500 M of P2, respectively. Concentrations of trypsin used were (a) 100, (b) 10, (c) 1, (d) 0.1, and (e) 0.01 ng/ml. Scale bar represents 1 mm...... 85

Figure 4.6. Limit of detections observed in the LC-based chymotrypsin assays by using P3-functionalized surfaces. The first and second columns represent surfaces functionalized by using 100 and 500 μM of P3, respectively. Concentrations of chymotrypsin used were (a) 100, (b) 10, (c) 1, and (d) 0.1 ng/ml. Scale bar represents 1mm...... 87

Figure 5.1. Interactions of LC with oligopeptide-coated microparticles and corresponding optical textures of LC. (a,b) Unfunctionalized microparticles, (c,d) P1-coated microparticles and (e, f) C18-terminated, P1-functionalized microparticles. (a,c,e) are bright-field images and (b,d,e) are dark-field images (under crossed polars). Scale bar represents 10 μm. The arrows indicate the orientations of polarizer and analyzer...... 100

Figure 5.2. Effect of oligopeptide concentration on the distortion radius in LC. C18- terminated, oligopeptide-functionalized microparticles submerged in the LC created a small halo near the microparticle as shown in the insert. Scale bar represents 10 μm...... 102

Figure 5.3. Effect of LC cell thickness on distortion radius near submerged microparticles...... 103

Figure 5.4. (a) Computational results for LC director profiles between the surface -4 2 of a microparticle (W2 = 4 × 10 J/m ) and an imaginary surface with preferential homeotropic and planar anchoring, respectively. The green-shaded region indicates the distortion region. (b) Influence of surface anchoring strength of microparticle on the computed distortion radius...... 107

xv the distortion region. (b) Influence of cell thickness on the computed distortion radius was introduced by varying the anchoring strength of the imaginary surface...... 109

Figure 5.6. Application of C18-terminated, oligopeptide-functionalized microparticles in LC as a trypsin assay. The C18-terminated, oligopeptide- functionalized microparticles were immersed in solutions containing (a) 0, (b) 0.5 or (c) 1 mg/ml of trypsin for 3 h to cleave the oligopeptides before immersion. Scale bar represents 10 μm...... 111

Figure 6.1. Chemical structures of (a) N,N-dimethyl-n-octadecyl-3- aminopropyltrimethoxysilyl chloride (DMOAP), (b) poly-(ethylenimine) (PEI), (c) sodium dodecyl sulfate (SDS), (d) Tween 20, and (e) cetyltrimethylammonium bromide (CTAB)...... 116

Figure 6.2. Effect of PEI-coating concentration on the surface density of reactive amine groups. Inset: Calibration plot of fluorescence intensity obtained using standard FITC solutions...... 122

Figure 6.3. Influence on the zeta potential of the adsorption of surfactants on PEI- coated microparticles. The surfactants studied were (a) cationic CTAB, (b) nonionic Tween 20, and (c) anionic SDS...... 123

Figure 6.4. Bright-field images of the assembly of PEI-coated microparticles at LC/water interfaces (a) decorated with 90 μM Tween 20 or (b) without Tween 20. PEI-coated microparticles assembled into short chains only in the presence of Tween 20. Control experiments in the presence of 90 μM Tween 20 were also conducted using (c) uncoated microparticles and (d) PEI-coated microparticles at 40 °C. 5CB became an isotropic liquid at this temperature. (e) Grid holes were completely filled with LC to remove the concave meniscus, and PEI-coated microparticles were introduced without Tween 20. The scale bar represents 100 μm...... 125

Figure 6.5. Assembly of PEI-coated microparticles that followed the LC director at the LC/water interface. (a) Cross-polarized image of the PEI-coated microparticles at an LC/water interface decorated with 90 μM Tween 20. (b) Observed LC director field. The scale bar represents 100 μm. The arrows indicate the orientations of polarizer and analyzer...... 126

Figure 6.6. Effect of the Tween 20 concentration on the assembly of PEI-coated microparticles at LC/water interfaces decorated with Tween 20. Concentrations of Tween 20 are (a) 45 μM, (b) 90 μM, (c) 900 μM, (d) 4.5 mM, and (e) 18 mM. Panels a−e show cross-polarized images, and panel f shows the bright-field counterpart of the image in panel e. (g) Magnified area of panel b illustrating the white halo surrounding microparticles in a short chain. The scale bars in panels a and g represent 100 and 10 μm, respectively...... 127

xvi

Figure 6.7. Visual detection of lipase activity using microparticles at an LC/water interface decorated with 90 μM Tween 20. Lipase concentrations are (a) 500, (b) 250, (c) 100, and (d) 10 ng/ml. Panels a−d show bright-field images, and the scale bar in panel d represents 50 μm...... 130

Figure 6.8. Assembly of PEI-coated microparticles at LC/water interfaces decorated with (a,b) 90 μM SDS or (c,d) 9 μM CTAB. Each pair consists of a bright- field image and a cross-polarized image for comparison. (e) Magnified cross- polarized image of the PEI-coated microparticles in panel b. Hedgehog defects are indicated by arrows. The scale bars in panels a and e represent 100 and 5 μm, respectively...... 131

Figure 6.9. Effect of SDS concentration on the assembly of PEI-coated microparticles at LC/water interfaces decorated with SDS. Concentrations of SDS are (a) 0.9, (b) 9, (c) 90, (d) 270, and (e) 450 μM. Panels a−e show cross-polarized images, and panel f shows the bright-field counterpart of the image in panel e. The scale bar represents 100 μm...... 135

Figure A.1. Effect of DMOAP coating duration on the water contact angles. .. 173

Figure A.2. Fluorescence intensity of immobilized P2 and P3 for different UV durations...... 174

Figure A.3. DTNB assay for concentration of free thiols: (a) Calibration of absorbance values using cysteine as standard. (b) Concentration of P6 at different oxidation durations. Absorbance values are tabulated at 412 nm and averaged from 3 readings...... 175

xvii

LIST OF SCHEMES

Scheme 2.1. Peptide Cleavage by Proteases ...... 18

Scheme 2.2. Principles of FRET-Based Protease Assays ...... 21

Scheme 2.3. Principles for Electrochemical-Based Protease Sensor ...... 32

Scheme 2.4. Principles of Enzyme-Linked Peptide Protease Assay ...... 35

Scheme 2.5. Principles of LC-Based Protease Assay...... 37

Scheme 3.1. Procedure for Streptavidin Binding Assay ...... 50

Scheme 3.2. Proposed Mechanism for Oligopeptide Immobilization ...... 61

Scheme 3.3. Oligopeptide Immobilization Density and Orientation ...... 64

Scheme 4.1. Surface Immobilization of Dual Cysteine-Labeled Peptides Through Formation of Inter-Peptide Disulfide Bonds ...... 77

Scheme 4.2. Detection Principles of the LC-based Protease Assays ...... 79

Scheme 5.1. Possible LC Director Profiles Surrounding Microparticles Submerged in LC ...... 100

Scheme 6.1. Microparticles Assembly at LC/Water Interface ...... 120

xviii

LIST OF SYMBOLS

 Actual anchoring angle

NA Avogadro’s number

Kd Dissociation constant

L Extrapolation length

K One constant approximation of Frank constant

K1 Frank constant for splay deformation

K2 Frank constant for twist deformation

K3 Frank constant for bend deformation d LC cell thickness

Mr Molecular weight

 Preferential anchoring angle

W Surface anchoring strength

 Surface density

xix

LIST OF ABBREVIATION

5CB 4-Cyano-4’-n-pentylbiphenyl

Ac Acetyl

ALP Alkaline phosphatase

AuNP Gold nanoparticles

BAEE Nα-Benzoyl-L-arginine ethyl ester

BoNT Botulinum neurotoxin

BRET Bioluminescent resonance energy transfer

BSA Bovine serum albumin

C- Carboxyl-terminus

C18 Octadecyl

CNT Carbon nanotube

CTAB Cetyltrimethylammonium bromide

Cy3 Indocarbocyanine

Cy5 Indodicarbocyanine

Cys C Cystatin C

Cyt C Cytochrome C

DABCYL 4-(4-dimethylaminophenylazo) benzoic acid

DHB 2, 5-dihydroxybenzoic acid

DI Deionized

DMOAP N,N-dimethyl-n-octadecyl-3-aminopropyltrimethoxysilyl

chloride

DMSO Dimethyl sulfoxide

DTNB 5, 5’-dithiobis-(2-nitrobenzoic acid)

EDANS 5-[(2-aminoethyl)amino] naphthalene-1-sulfonic acid

ELISA Enzyme-linked immunosorbent assays

FAM Carboxyfluorescein

Fc Ferrocene

FITC Fluorescein 5-isothiocyante

FRET Fluorescence resonance energy transfer

GO Graphene oxide

HAN Hybrid aligned nematic

HIV Human immunodeficiency virus

LC Liquid crystals

LcA Light chain protease

LCD Liquid crystals display

LOD Limit of detection

MALDI-TOF-MS Matrix-assisted-laser-desorption-ionization time-of-flight

mass-spectrometry

MB Methylene blue

MMP Matrix metalloproteinase

N- Amino-terminus

NHS N-hydroxysuccinimide

OBOC One-bead-one-compound pADA p-aminodiphenylamine

PBS Phosphate buffered saline

xxi

PEI Poly(ethylenimine)

PMT Photomultiplier tube

PSA Prostate-specific antigen

PVA Polyvinyl alcohol

QD Quantum dot

SDS Sodium dodecyl sulfate

Se Selenide

SERS Surface-enhanced Raman scattering

SPR Surface plasmon resonance

TAMRA Carboxytetramethylrhodamine

TEA Trimethoxysilane aldehyde

TEM Transmission electron microscopy

TLC Thin layer chromatography

UCP Upconverting phosphors

US FDA United States Food and Drug Administration uPA Urokinase-type plasminogen activator

UV Ultraviolet

XPS X-ray photoelectron spectroscopy

ZnS Zinc sulfide

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CHAPTER 1

CHAPTER 1 INTRODUCTION

CHAPTER 1

1.1 Background

1.1.1 Importance of Proteases and Protease Assays

Proteases are enzymes that catalyze the hydrolysis of peptide bonds in peptide chains. They are classified based on their catalytic sites, namely aspartic, glutamic, cysteine, serine, and threonine proteases. Matrix metalloproteinases (MMP), on the other hand, refer to proteases that require metal ions for their catalytic activities.1

In humans, 1.7% of human genes is coded for proteases, and proteases are responsible for basic functions such as protein digestion, wound healing, and activation of immune systems.2 Furthermore, protease activities are linked to diseases such as cancers,3 cardiovascular diseases,4 Alzheimer’s disease,5 human immunodeficiency virus (HIV),6 thrombosis,7 and diabetics.8 Inhibitions of disease-related proteases can potentially be used as drug molecules to treat these diseases. One canonical example is the case of HIV, where medications in the form of protease inhibitors have been proven to prolong the life expectancy of HIV- positive patients.9 Consequently, several drugs in the form of protease inhibitors are United States Food and Drug Administration- (US FDA)-approved for the treatment of several diseases, like multiple myeloma, blood coagulation disorders, and diabetes.10 Therefore, protease assays are valuable diagnostic tools, and importantly, they can serve as the screening platform for disease treatments. In comparison, DNA,11 RNA,12 protein,13 or lipid assays are also important for disease diagnostics.14 However, these assays are not readily applied in the screening for treatment drugs. Given such compelling motivations, extensive efforts have been devoted to developing sensitive, selective, and robust protease assays for

CHAPTER 1

quantitative analysis of proteases and their inhibitors. The primary function of proteases is to hydrolyze the peptide bond between two amino acids (endopeptidase) or terminal amino acids at the amino (N-) or carboxyl (C-) terminals

(exopeptidase).1 This results in the fragmentation of a single peptide chain into two shorter peptide chains. For example, serine proteases like trypsin hydrolyzes the peptide bonds of positively-charged lysine and arginine.15 On the other hand, chymotrypsin hydrolyzes the peptide bonds of hydrophobic tryptophan, tyrosine, and phenylalanine.15 Furthermore, studies showed that amino acids away from the hydrolysis sites also influence protease activities. For example, presence of aspartic acid at several positions away from the site of peptide hydrolysis increased thrombin activities.16

Mechanistically, hydrolysis of peptide chains by proteases involve specific interactions between catalytic sites of proteases and complementary amino acid sequences through covalent bond formations, electrostatic attractions, hydrogen bonding or van der Waal’s interactions at the active sties of proteases.15, 17 Such specificity of protease activities results in the hydrolysis of peptide chains that contain specific amino acid sequences. Given this understanding, custom-made peptide substrates are often used in protease assays to detect the presence of a target protease. This is complemented by the advent of solid-phase peptide synthesis that has empowered us to produce peptides with pre-determined amino acid sequences.18

CHAPTER 1

A range of detection methods are used in protease assays, including colorimetric,19-

20 mass spectrometry,21-23 fluorescence resonance energy transfer (FRET),24-33 electrochemical,34-47 surface-enhanced Raman scattering (SERS),48-51 and surface plasmon resonance (SPR).52-55 More recently, nanomaterials such as nanoparticles,56-67 quantum dots (QD),68-80 and graphene oxides (GO)81-85 are also used for the development of protease assays with impressive detection limits.

1.1.2 Introduction of Liquid Crystals

Liquid crystals (LC) usually consist of rod-like molecules that form phases with both fluidity of liquids and molecular ordering analogous to solid crystals in one or two dimensions. Several forms of LC phases can exist, and this depends on their molecular ordering patterns. For example, smectic phases of LC consist of two- dimensionally ordered stacks of molecules with defined inter-layer spacings. In comparison, nematic LC phase consists of molecules with one-dimensional order that are oriented in a preferential direction (or director ). It is noted that the signs of LC director ( or –) do not result in distinguishable physical and optical properties.86 In this thesis, nematic LC is used due to its sensitivity to physical and chemical environment. Furthermore, some commonly used nematic LC, for example 4-cyano-4’-n-pentyl-biphenyl (5CB), forms stable nematic LC phase at room temperature. This improves the robustness of nematic LC-based applications.

From henceforth, nematic LC shall be referred to as LC.

CHAPTER 1

Thermodynamically, free energy density of LC systems can be described by the

Frank free energy model:86-88

(1.1)

Where K1, K2 and K3 are Frank elastic constants for splay, twist, and bend deformations respectively, is the average LC director vector, Wa is the surface anchoring strength, and (θ-θ0) is the deviation of actual LC director (θ) from the preferential anchoring (θ0) at the boundary surface. The first three terms describe the energy penalty of the system due to elastic deformation of the director field, while the fourth term describes anchoring energies arising from the deviation of LC orientation from its preferred direction at the boundary surfaces. Essentially, overall frank free energy is the minimization between bulk elastic energy and surface anchoring energy. Importantly, for biosensors applications, changes to surface composition due to the presence of target analytes can potentially result in the alterations of preferential surface anchoring orientation and strengths.

Consequently, minimization of overall Frank free energy (1.1) may amplify surface composition changes to alter orientation of LC in the bulk phase. This important property is extensively used in LC-based assays, where changes to orientation of

LC due to surface composition changes can alter LC orientation in the bulk phase.

For example, peptide or protein functionalization of DMOAP-coated surfaces changes the preferentially anchor LC from homeotropic to planar, and such surfaces cause LC to be oriented planarly in the bulk phase.89-90 Interestingly, such changes in LC orientation can be visualized due to optical birefringence of LC. Light that

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propagates through them undergoes two distinct light paths of different refractive indices, namely the ordinary and extraordinary refractive indices. Importantly, polarization of these two light paths are orthogonal to each other. Therefore, when linearly-polarized light is incident on an LC sample, the interference patterns of emerging light result in elliptically-polarized light. With reference to a canonical setup with bulk LC phase confined within a 200 m-wide square grid, observations of such samples under cross-polarized light typically yield a characteristic “maltese cross” appearance. This is a result of the preferential anchoring of LC at the boundary surfaces, where the bottom glass surface and confining square grid surfaces orients LC homeotropic (or perpendicular) to the surfaces. In contrary, the top surface preferentially anchors LC planarly (or parallel) to the surface. The results in the radial orientation of LC director within the grid. Such radial orientation of LC results in the observation of the “maltese cross” appearance. For bright regions in the “maltese cross”, light had emerged elliptically polarized, and this occurs when the azimuthal angle of LC orientation is finitely different from the angle of incident, linearly polarized light. The azimuthal angle must also be finitely different from the angle of polarization at the analyser. Consequently, the dark regions in the “maltese cross” appearance demarcates regions of LC where azimuthal angles are either parallel to the incident, linearly polarized light source, or the angle of polarization at the analyser.91

Another interesting property of LC is their ability to nucleate topological defects and organize micrometer-size particles into one- or two-dimensional structures.

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This phenomenon stems from the elastic reorientation of LC director around each microparticle as a result from the minimization of overall Frank free energy. A canonical example is a microparticle with preferential homeotropic anchoring. For such a microparticle, LC director around the microparticle is preferentially anchored radially. However, when this microparticle is placed within a bulk LC phase with a uniform director, it is impossible to satisfy both the preferential surface anchoring and bulk LC director without the formation of a region of defect. Defects are regions where the orientational order of LC breaks down, and LC behaves as isotropic liquids in these microscopic regions. In this case of a microparticle with preferential homeotropic anchoring, a hyperbolic hedgehog defect is commonly observed. This results in a dipolar characteristic of LC director field around the microparticle. Consequently, the presence of multiple “dipolar” microparticles in close vicinity can lead to the assembly of chain-like assembly. Such phenomenon has been extensively studied, with the observations of various defect structures and assembly patterns.87, 92 However, there has been limited studies on how such microparticles assemblies can be applied in sensing applications.

1.1.3 LC-Based Protease Assays

Recently, LC-based assays are emerging as excellent label-free assays for many bioanalytical applications. These LC assays are simple to use, and yet highly sensitive. Such assays make use of the ability to influence orientation of LC by engineering chemical composition of surfaces supporting the LC.93-94 This phenomenon was rigorously exploited by Abbott and co-workers, who established

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methods to detect the adsorption of biomolecules on surfaces or LC/water interfaces.95-103 These events were visualized directly by using changes in optical textures of LC. Later on, Yang and co-workers demonstrated excellent sensitivity, selectivity, and multiplex capability of LC-based protease assays.89, 104 However, several key challenges remained. First, stable immobilization of oligopeptides on surfaces compatible with LC-based assays is lacking. Second, detection of minute changes in peptides upon cleavage is challenging, since the surface property does not change drastically. Moreover, the limited surface area has led to poor detection limits. Finally, most assay formats are not capable of providing quantitative and real time protease activity monitoring.

1.1.4 Immobilization of Oligopeptide on LC-Compatible Surfaces

Proteases are able to recognize and cleave complementary amino acid sequences through covalent bond formation, electrostatic attraction, hydrogen bonding, or van der Waal’s interactions at the active sites of proteases.15, 17 Thus, custom-made peptide substrates containing specific amino acids cleavable by target proteases are often used in protease assays. This is complemented by the advent of solid-phase peptide synthesis for the synthesis of peptides with pre-determined amino acid sequences.18 In order to use oligopeptides in heterogeneous protease assays, oligopeptides need to be immobilized on a solid surface with controlled orientation and a high density.105-107 Specifically, for LC-based detection, LC-compatible surfaces are required. These surfaces are typically modified glass surfaces that preferentially orient LC molecules planarly (parallel to the surface) or

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homeotropically (perpendicularly to the surface). For example, N, N-dimethyl-n- octadecyl-3-aminopropyltrimethoxysilyl chloride- (DMOAP)-coated surfaces are commonly used. DMOAP is an organosilane that presents an octadecyl (C18) group and a quaternary ammonium salt. The positive charge on the quaternary ammonium makes DMOAP soluble in water. In the past, glass slides coated with DMOAP were widely used in LCD to control the orientations of LC.93, 108 Recently, DMOAP- coated slides have also been used for immobilization of oligonucleotides and oligopeptides.89, 109 On DMOAP-coated surfaces, past studies showed that LC 5CB is oriented homeotropically. Such DMOAP-coated surfaces and 5CB were used by

Chen and Yang for the fabrication of LC sensors to detect protease activities.109

Because DMOAP does not have any reactive groups, the surfaces need to be activated by ultraviolet (UV) light before oligopeptides can be immobilized.109-110

Such UV activation produces aldehyde groups, which serve as reactive groups for oligopeptide immobilization. On the aldehyde-functionalized surfaces, it was proposed that immobilization of oligopeptides can be achieved through the presence of cysteine residues.109, 111 However, the exact mechanism of surface activation and oligopeptide immobilization on the DMOAP-coated surface are still unclear. Moreover, oligopeptides immobilized on the surface are unstable under rigorous washing conditions. Therefore, in-depth mechanistic study for the immobilization of cysteine-labeled oligopeptide on DMOAP-coated surfaces is required to overcome these limitations. Additionally, to improve the robustness of

LC-based protease assays, it is desirable to have stable peptide immobilizations that can withstand harsh washing conditions. These washing conditions are important

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to remove non-specifically adsorbed proteases from the surfaces. One solution was suggested by Chen and Yang, where they used multiple lysine residues for improved immobilization stability.104 However, lysine residues are cleavable by trypsin, and the specificity of the assay was questionable. Thus, the challenge to improve the stability of immobilized peptides remains.

1.1.5 Signal Enhancement in LC-Based Protease Assays

In a typical LC-based protease assay, immobilized oligopeptides are cleaved by a target protease, and short oligopeptide fragments are released from the surface. LC is then applied to the surface to detect the remaining oligopeptides. Ideally, cleavage of oligopeptides should change surface properties, which in turn influence the orientations of LC substantially. However, the changes in surface properties may be insufficient to disrupt orientations of LC and produce optical signals. For example, Bi et al. reported that oligopeptides shorter than 9 amino acid residues were unable to disrupt orientations of LC, and that resulted in a poor limit of detection (~3 g/ml) for chymotrypsin.89 More recently, Chen and Yang used peptides ≥36 amino acid residues with multiple lysine residues to ensure that cleavage of the peptides led to the significant disruption of LC orientations.104 They reported an improved limit of detection (LOD) of 1 ng/ml and 0.1 ng/ml for trypsin and chymotrypsin, respectively. However, due to the use of multiple lysine residues, trypsin and chymotrypsin activities could not be differentiated. Therefore, more work is required to develop an amplification mechanism to enhance LC

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signals, and this will aid in the development of sensitive LC-based protease assays.

Importantly, the protease assay must retain its selectivity.

1.1.6 Oligopeptide-Functionalized Microparticles in LC

For LC-based assay on flat surfaces, interactions between oligopeptides and LC are limited. Therefore, to increase interactions between LC and oligopeptides, oligopeptide-functionalized microparticles with a high surface area can be exploited in LC-based protease assays. Recent research has focused on the assembly of microparticles dispersed in LC or at heterogeneous LC interfaces,112-

120 and trapping of microparticles at specially fabricated surfaces.87, 121-123 However, the use of microparticles dispersed in the LC phase has not been studied for sensing applications. Compared to traditional LC-based assays on functionalized flat surfaces, the use of high surface-to-volume microparticles can potentially increase the interaction between functionalized surfaces of microparticles with LC.

Furthermore, distortion of LC director profiles by the microparticles can potentially produce distinct and observable LC optical signals for quantitative data interpretation. The use of microparticles can also potentially improve the throughput of assays, where microparticles can be functionalized with a library of oligopeptides for screening purposes. However, some fundamental studies on the interactions between oligopeptide-functionalized microparticles and LC are required for the development of microparticle-based protease assays.

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1.1.7 Real-Time Protease Activity Monitoring

In addition to the detection of proteases, real-time monitoring of protease reactions to obtain kinetic information such as protease activity is very important. In particular, understanding the inhibition kinetics of disease-related protease activities, like HIV-1 protease,9 are of great interest. However, most LC-based protease assays are not suitable for real-time monitoring of protease activity.89, 104,

109 For real-time monitoring of protease activity using LC-based assays, the use of

LC/water interfaces is a suitable platform. Work by Jang and co-workers showed the potential of using such heterogeneous interfaces for monitoring of enzymatic activities.124-130 For example, they showed that cleavage of poly-L-lysine membrane by trypsin at an LC/water interface caused the interface to be permeable to phospholipids in the water phase.129 This resulted in a planar-to-homeotropic transition of LC. However, some of these studies relied on the use of poly-L-lysine, and this limits the applicability of these studies for detection of other proteases.

Furthermore, immobilization of peptide substrates at such a fluid interfaces are more challenging and less-studied. To overcome these limitations, self-assembly of microparticles at the LC/water interfaces can be studied. Because microparticles are in constant contact with both LC and water phases, this system can also be potentially useful for monitoring protease activity in real-time. Additionally, surfaces of microparticles can be functionalized with oligopeptides for selective protease sensing. However, more work is required to provide a fundamental understanding on how microparticles assemblies are influenced by changes in surface compositions of the microparticles.

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1.2 Research Objective

The research objectives of this thesis are as follows:

(1) The first objective is to understand the role of cysteine during the immobilization of oligopeptides on DMOAP-coated surfaces. This is crucial for

LC-based protease assays, as stable immobilization of oligopeptides with controlled orientation on DMOAP-coated surfaces will provide optimal interactions between target proteases and immobilized-oligopeptides. First, we carry out a mechanistic study on the role of cysteine during the immobilization.

Second, we study the stability of these immobilized oligopeptides by subjecting oligopeptide-functionalized surfaces to stringent washing conditions. Third, we investigate whether the orientation of immobilized oligopeptides was optimized to interact freely with any analytes present in the aqueous phase. Finally, we study the use of dual cysteine-labeled peptides to further improve stability and density of immobilized peptides. Besides using cysteines as oligopeptide immobilization sites, thiols of cysteines can also form disulfide bonds. Therefore, we can investigate if this property of thiol groups in cysteines can be utilized to improve the stability and density of oligopeptide immobilization.

(2) The second objective is to develop an LC-based functional protease assay with high sensitivity and selectivity, using trypsin and chymotrypsin as model proteases.

The current challenge for LC-based protease assay involves the need for oligopeptide-functionalized surfaces to remain stable under stringent washing conditions, and these surfaces must preferentially orient LC planarly. To achieve

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this, we develop an amplification mechanism that can significantly enhance LC signals for oligopeptide-functionalized surfaces. One possible approach is the introduction of biotin labels to oligopeptides, where we can investigate whether binding of streptavidin to the surface can amplify LC optical signals. Based on this approach, we can potentially ensure that LC are oriented planarly in the presence of oligopeptides. Additionally, we can ensure a selective protease assay, since this approach does not require any specific amino acids to be present in the oligopeptide.

(3) The third objective is to understand interactions between oligopeptide- functionalized microparticles and LC, and to exploit this phenomenon for detecting protease activity. A fundamental issue for such protease assays is that preferential orientation of LC at oligopeptide-functionalized microparticles must change from homeotropic to planar orientation, or vice versa. To achieve this, we explore the interaction of LC with microparticles functionalized with C18-terminated oligopeptides. Since DMOAP also contains C18 groups, we investigate if microparticles functionalized with C18-terminated oligopeptides can preferentially orient LC homeotropically. Furthermore, we can assess if the cleavage of oligopeptide can change the preferential orientation of LC to planar orientation.

This way, observable and distinct LC optical signal can be produced and used for detection of target protease activities.

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(4) The last objective is to develop a sensing platform capable of real-time monitoring of protease activities. To achieve this, we explore the possibility of forming assemblies of microparticles at LC/water interfaces. In order to monitor protease activities in real-time, such microparticles assemblies must respond to the cleavage of protease substrates present in the water phase. To achieve this, a proof- of-concept study is carried out using polyethylenimine- (PEI)-coated microparticles.

Since amine groups of PEI-coated surfaces can interact with any analytes present in the water phase through hydrogen bonding or electrostatic interactions, surfactants such as sodium dodecyl sulfate (SDS), Tween 20, and cetyltrimethylammonium bromide (CTAB) can be introduced into the water phase as model protease substrates. Since some of these surfactants can selectively adsorb on these microparticles through electrostatic attractions or hydrogen bonding, we can study the response of microparticle assemblies when the surface composition of microparticles changes. Additionally, since Tween 20 is a synthetic substrate of lipases, preliminary studies can be carried out to assess the response of microparticles assemblies to lipase activities. This study can provide the basis for future development of real-time protease activity monitoring using microparticles assembly at LC/water interfaces.

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CHAPTER 2 LITERATURE REVIEW

In this chapter, we review on the recent developments of protease assays.

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2.1 Introduction

Proteases are enzymes that catalyze the cleavage of peptide bonds in peptide chains.

They are classified based on their catalytic sites, namely aspartic, glutamic, cysteine, serine, and threonine proteases. Matrix metalloproteinases (MMP), on the other hand, refer to proteases that require metal ions for their catalytic activities.1

In humans, 1.7% of human genes is coded for proteases, and proteases are responsible for basic functions like digestion of food, wound healing, and activation of immune systems.2 Furthermore, protease activities are linked to diseases like cancers,3 cardiovascular diseases,4 Alzheimer’s disease,5 human immunodeficiency virus (HIV),6 thrombosis,7 and diabetics.8 Inhibitions of disease-related proteases can potentially be used to treat these diseases. One canonical example is the case of HIV, where medication in the form of protease inhibitors has been proven to prolong the life expectancy of HIV-positive patients.9

Given such compelling motivations, extensive efforts are devoted to developing sensitive, selective and robust protease assays for identification and quantification of proteases and their inhibitors. The primary function of proteases is to cleave a peptide bond between two amino acids (endopeptidase) or terminal amino acids at the amino (N-) or carboxyl (C-) terminals (exopeptidase). This results in the fragmentation of a single peptide chain into two shorter peptide chains. Scheme 2.1 describes the cleavage at positions P1 and P1’ of a peptide chain. Such hydrolytic activities are highly specific, and proteases recognize specific amino acids at P1 and P1’. For example, serine proteases like trypsin cleave the peptide bonds of positively-charged lysine and arginine at the P1 position.15 On the other hand,

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chymotrypsin cleaves the peptide bonds of hydrophobic tryptophan, tyrosine and phenylalanine at P1 position.15 Furthermore, studies showed that amino acids at several positions away from the cleavage sites also influence protease activities.

For example, the presence of aspartic acid at the P10 position improved thrombin activities.16

Scheme 2.1. Peptide Cleavage by Proteases

Protease cleavage that recognizes specific amino acids P1 and P1’. For thrombin, aspartic acid at P10 was also important for optimum cleavage.

Mechanistically, cleavage of peptide chains by proteases involves specific interactions between catalytic sites of proteases and complementary amino acid sequences through covalent bond formation, electrostatic attraction, hydrogen bonding, or van der Waal’s interactions at the active sites of proteases.15, 17 Such specificity of protease activities result in cleavage of peptide chains that contain specific amino acid sequences. Given this understanding, custom-made peptide substrates are often used in protease assays to detect the presence of a target protease. This is complemented by the advent of solid-phase peptide synthesis that has empowered us to produce peptides with pre-determined amino acid sequences.18

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In terms of operation modes, protease assays can be classified into homogeneous or heterogeneous assays. For homogeneous assays, both substrates and samples are present in an aqueous phase. Conversely, heterogeneous assays consist of protease substrates that are immobilized on a solid platform, while samples are in the aqueous phase. On the basis of detection methods, homogeneous assays can be further classified into colorimetric assays,19-20 mass spectrometry-based assays,21-

23 and fluorescence resonance energy transfer (FRET) assays.24-33 More recently, nanomaterials such as nanoparticles,56-67 quantum dots (QD),68-80 and graphene oxides (GO)81-85 are also used in the homogenous protease assays with impressive detection limits. However, homogenous assays are generally not capable of multiplexed sensing. This arises due to a limited number of probes that can produce distinct signals. For example, fluorogenic probes are constrained by their emission profiles, and only a limited number of probes can be used to ensure well-resolved emission signals. This limitation can be circumvented by using heterogeneous assays, where a large number of substrates can be immobilized at discrete locations on a solid surface. This allows simultaneous and multiplexed detection of proteases present in a single sample. Detection results will depend on signal levels at discrete locations on the solid surface. Compared to homogeneous assays, heterogeneous assays are capable of label-free sensing and do not rely on probes that produce signals. Heterogeneous assays can be further classified into electrochemical assays,34-47 surface-enhanced Raman scattering (SERS) assays,48-51 and surface plasmon resonance (SPR) assays.52-55

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Recently, liquid crystals- (LC)-based assays are emerging as excellent label-free assays for many applications. These LC assays are simple to use, and yet highly sensitive. They also generate optical signals that are visible to the naked eye. LC are typically rod-like molecules that are anisotropic. They orient in a preferential direction when subjected to a magnetic or electric field. Such preferential orientation results in birefringent of LC, and they display unique optical textures when viewed under cross polarizers.86 This phenomenon is ubiquitously applied in

LCD. Another interesting property is the ability to influence orientation of LC by engineering the chemical or morphology of surfaces confining the LC phase.93-94

This phenomenon was rigorously studied by Abbott and co-workers, who established methods to detect adsorption of biomolecules on surfaces or LC/water interfaces. These events were visualized directly by using changes in optical textures of LC.95-103 Later on, Yang and co-workers demonstrated excellent sensitivity, selectivity, and multiplex capability of LC-based protease assay.89, 104

In this review, we discuss recent developments of protease assays according to their assay formats (homogenous or heterogeneous) and detection principles. We also assess the performance of LC-based assays compared to other protease assays.

2.2 Homogenous Protease Assays

In a homogenous protease assay, a common element is that protease activity leads to increase or decrease of fluorescence caused by FRET phenomenon. This type of sensor often consists of a short peptide link to an FRET donor and an FRET quencher in close proximity (1-10 nm), where emission of the fluorescence donor

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is quenched by an acceptor.131 Thus, cleavage of the FRET peptide substrate will cause the FRET pair to separate and fluorescence signal to increase (Scheme 2.2a).

For such homogeneous assays, both protease and its substrate are in the aqueous phase. The main difference among these assays lies in the donors and acceptors, which range from organic molecules to nanomaterials.

Scheme 2.2. Principles of FRET-Based Protease Assays

(a) FRET assays based on organic FRET donor and acceptors. (b) FRET assays based on nanomaterials as FRET acceptors and FRET donor- labeled protease substrates. (c) FRET assays based on nanomaterials as FRET donors and FRET acceptor- labeled protease substrates.

2.2.1 Organic FRET Donors and Acceptors

Traditionally, FRET-based protease assays use organic FRET acceptor and donor covalently linked to a peptide chain (Scheme 2.2a). For example, Matayoshi et al. synthesized HIV protease substrates with organic fluorescent donor, 5-[(2-

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aminoethyl)amino] naphthalene-1-sulfonic acid (EDANS), and fluorescent quencher, 4-(4-dimethylaminophenylazo) benzoic acid (DABCYL), at N- and C- terminal of the peptide substrates, respectively.132 To improve detection limits,

Zauner et al. incorporated an enzyme cascade to improve the sensitivity of such

FRET-based protease assays.133 They introduced trypsinogen, which was activated in the presence of enteropeptidase to trypsin. This simple enzymatic cascade system greatly amplified enteropeptidase activities, and they detected 0.2 pM of enteropeptidase using optimized peptide sequence labeled with fluorescein (FRET donor) and carboxytetramethylrhodamine (TAMRA, FRET acceptor). Elsewhere,

Fuchs et al. incorporated peptide substrates labeled with FRET donors and receptors at the surface of polystyrene nanoparticles.59 It was concluded that cleavage of peptide substrate led to an increase in fluorescence signal. However, reliance on organic FRET acceptors and donors are susceptible to long-term stability and photo-bleaching issues. Thus, the focus has been shifted to nanomaterials which can overcome these stability issues.

2.2.2 Nanomaterials as FRET Acceptors or Donors

Nanomaterials like carbon nanotubes (CNT), graphene oxide (GO) and gold nanoparticles (AuNP) have been successfully employed as effective FRET acceptors in biosensing applications.134 For CNT- and GO-based materials, strong overlapping of -electrons stacking in these materials are ideal for application as

FRET acceptors.135 Furthermore, peptides can be immobilized on CNT or GO through bioconjugation to form an FRET-based protease assay (Scheme 2.2b).136

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For example, Huang et al. covalently immobilized three peptide substrates, each labeled with a different organic dye (FRET donor), on CNT (FRET acceptor).137

They detected three different proteases [limit of detection (LOD): 0.02 pM of matrix metalloproteinase- (MMP)-7, 0.07 pM of MMP-2 and 0.016 nM of urokinase-type plasminogen activator (uPA)] by monitoring fluorescence emissions of each organic dye. A similar approach was adopted by Zhu et al., where they used CNT-based material (FRET acceptor) to electrostatically attract fluorescence-labeled peptide substrates (FRET donor).138 Cleavage of peptide substrate resulted in observable emissions of fluorescence label, and they reported a LOD of 0.1 nM for thrombin.

Similar detection strategies involving GO (FRET acceptor) and fluorescence- labeled peptide substrates (FRET donor) were also demonstrated by other groups.83-

84, 139-141 For example, Feng et al. used GO (FRET acceptor) and fluorescence- labeled peptide substrate (FRET donor) to detect prostate-specific antigen (PSA).84

Fluorescence signals increased in the presence of target PSA, and they reported a

LOD of 0.3 nM. Gu et al. also demonstrated the detection of trypsin using a similar concept of GO (FRET acceptor) and fluorescence-labeled peptide substrate (FRET donor).139 Interestingly, GO has also been reported as FRET donors by Kwak et al., where fluorescent property of GO was used in a FRET system for the detection of chymotrypsin and MMP-2.85

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AuNP is also actively pursued as effective FRET acceptors.142-144 Furthermore, extensive studies and protocols on surface functionalization of AuNP surfaces are available.145 For peptide immobilization, thiols in cysteine residues are commonly used to covalently immobilize peptides. For example, Wang et al. used peptides with terminal cysteine residues to immobilize fluorescence-labeled peptides (FRET donor) on AuNP (FRET acceptor).146 Thus, cleavage of fluorescence-labeled peptides increased fluorescence signals, and a LOD of 0.033 nM to 0.142 nM for different proteases were achieved. Park et al. also demonstrated the use of fluorescence-labeled peptides (FRET donor) immobilized on AuNP (FRET acceptor) for detection of MMP-7.147

These developments showed good sensitivity and selectivity towards protease activities. Nanomaterials serve dual-roles as FRET acceptors and robust platform for immobilization of protease substrates. However, reliance on organic dyes as

FRET donors are still susceptible to stability issues.

2.2.3 QD as FRET Donors

As organic fluorescence dyes are susceptible to photo-bleaching and narrow excitation-emission peak separations, inorganic QD are emerging as excellent fluorescence materials. They boast superior photo-stability, high molar absorption coefficient, and wide excitation-emission peak separation.148 In the past, QD has been used as FRET donors in protease assays, in which peptide substrates were immobilized on the surface of QD and labeled with a FRET quencher. Such designs

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allow emissions of QD to regain upon cleavage of the immobilized peptide substrates (Scheme 2.2c). For example, Sapsford et al. studied the use of QD to detect botulinum neurotoxin (BoNT) serotype A light chain protease (LcA) using cadmium selenide/zinc sulfide (CdSe/ZnS) core/shell QD.73 Peptide substrates were immobilized on the surface of QD and labeled with indocarbocyanine (Cy3) dyes. Wang and Xia also developed a similar protease sensor for thrombin with a

LOD of 0.03 M. In this case, they used CdSe/ZnS QD and indodicarbocyanine

(Cy5) as the donor and acceptor pair.71 Elsewhere, Serrano et al. successfully detected trypsin activity using QD functionalized with TAMRA-conjugated peptide substrates, with a LOD of 1 nM.72 However, the above-mentioned studies involving

QD still required organic dyes as FRET acceptors, which are susceptible to stability issues. To overcome this, Lowe et al. used AuNP as FRET acceptors in combination with QD for the detection of protease uPA. However, this required extensive surface modifications of QD with streptavidin and peptide substrates with terminal cysteine and biotin.74 To simplify the process, label-free method was proposed by

He and Ma.70 They showed that dithiol peptides inhibited cadmium-telluride

(CdTe) QD formation, while monothiol promoted QD formation. Thus, trypsin activities that cleaved dithiol peptide substrates to monothiol peptide fragments resulted in QD formation and observable QD fluorescence emission. In this system, they reported a LOD of 0.8 pM. Another label-free method was proposed by Li et al..80 They used thrombin-specific peptide substrates that induced aggregation of

CdTe QD and quenched fluorescence emission. The presence of thrombin activities caused the dispersion of QD and increased fluorescence signals, and they reported

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a LOD of 0.2 nM. Wu et al. also suggested the use of label-free substrates for serine proteases.69 They showed that presence of native cytochrome C (Cyt C) quenches the phosphorescence of manganese-doped ZnS QD, and cleavage of Cyt C by trypsin increases phosphorescence signals.

2.2.4 Gold Nanoclusters (AuNC)

AuNC are clusters of tens of Au atoms, and they are highly fluorescent.149 They are potential candidates to replace organic fluorescence dyes for biosensing. In the literature, protein-encapsulated AuNC can be synthesized in the presence of bovine serum albumin (BSA). This system can be exploited as a platform for protease detection.150 Wang et al. reported that fluorescence of AuNC decreased when BSA was cleaved by proteases.151 They hypothesized that cleavage of BSA exposed

AuNC to ambient oxygen and reduced AuNC fluorescence. Lin et al. also made use of this concept to detect cystatin C (Cys C).152 However, numerous proteases also cleave BSA, resulting in poor selectivity. To overcome this, Gu et al. used peptide templates for AuNC synthesis.153 This allowed selective detection of elastase, with an impressive LOD of 0.03 nM. Thus, AuNC can potentially function as a label- free, fluorescence-based protease sensor. However, there are no reports of a generic approach to select peptide templates for AuNC synthesis, and this limits wide application of this technique for different target proteases.

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2.2.5 Nanoparticles in Colorimetric Assays

Nanoparticles are commonly used in colorimetric assays due to their plasmon resonance phenomenon.154 Aggregation of nanoparticles can lead to larger nanoparticles which exhibit different colors.155 Making use of such principles, Xue et al. developed trypsin assays by using peptides with repeated arginine residues to induce aggregation of AuNP.66 Cleavage of peptide substrate resulted in the the dispersion of AuNP and observable color change from purple to red with an LOD of 0.07 nM. To improve selectivity, Ding et al. reported the use of customized peptides that induced aggregation of AuNP.57 Thus, cleavage of peptides by trypsin caused the dispersion of AuNP. Elsewhere, Chen et al. reported the use of peptide- immobilized AuNP for the detection of MMP-7.62 Substrate peptides prevented aggregation of AuNP through their secondary structures,156 and cleavage of the substrates by MMP-7 induced AuNP aggregation. Using a similar approach, Chen et al. immobilized negatively-charged peptides on AuNP that remained dispersed, and cleavage of the peptide substrate promoted aggregation of AuNP.157 Using this method, they reported a LOD of 5 nM for trypsin and MMP-2. These developments demonstrated the feasibility of using AuNP for detecting proteases.145

Nanoparticle-based detection is advantageous because color changes can be detected with the naked eye or spectrometry for quantitative analysis. Moreover, plasmon resonance is not susceptible to photo-bleaching, as compared to fluorescence-based methods mentioned above.

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Table 2.1. Summary of Homogeneous Trypsin and Chymotrypsin Protease Assays Recently Developed Reference Detection Protease LOD Principle Substrate a Zauner et al. 133 Fluorescence Flb-CARSG- Trypsin (2 pM) TAMRAc 139 d Gu et al. Fluorescence R6-FAM Trypsin (~4 nM)

He and Ma 70 Fluorescence GCKGCG Trypsin (4 pM)

Serrano et al. 72 Fluorescence CKRVK- Trypsin (0.043 M) TAMRAc 31 Fan et al. Fluorescence R5 Trypsin (0.011 nM) Chymotrypsin (6 pM) Wang et al. 146 Fluorescence Bovine serum Trypsin (0.7 nM) albumin Chymotrypsin (0.4 nM) 66 Xue et al. Colorimetric R6 Trypsin (0.07 nM)

Ding et al. 57 Colorimetric YHPQMNPY Trypsin (0.5 nM)

TKAG3-C 157 Chen et al. Colorimetric E4GLLGALG Trypsin (5 nM) KC a Peptide sequences follow the one-letter code for amino acids. b Fl: acetamidofluorescein; c TAMRA: carboxytetramethylrhodamine; d FAM: carboxyfluorescein

We tabulated Table 2.1 to show a summary of homogeneous protease assays for trypsin and chymotrypsin, keeping in mind that the inherent activities of proteases used in these studies may vary, and the concentrations presented serve as a common unit for comparison. Among all techniques, fluorescence-based protease sensor developed by Fan et al. has the lowest LOD of 6 pM for chymotrypsin.31 However, this technique requires peptide substrates flanked with repeated arginine residues,

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and this may limit its applications for proteases that do not cleave such substrates.

For trypsin, the assay developed by He and Ma has the lowest LOD of 4 pM.70 It must be noted that this technique requires substrates with 2 cysteine residues, which is required to influence the formation of QD. However, no in-depth analysis on the role of cysteine residues was provided. Thus, more studies would be required for better understanding to generalize such protease assays. Furthermore, homogenous assays suffer from limited multiplex sensing capabilities.

2.3 Heterogeneous Protease Assays

Heterogeneous protease assays require the immobilization of protease substrates on solid platforms. These platforms allow proteases in the aqueous phase to interact with the immobilized substrates, and a large number of substrates can be immobilized on a solid platform for multiplex sensing applications. This is possible as optical or fluorescent signals at discrete locations can be identified and associated with a specific protease. An important consideration for heterogeneous assay is the choice of solid platforms for signal generation. For example, electrochemical-based sensors require conductive platforms like metals, while colorimetric assays usually require optically transparent platforms like glass. Thus, careful design of peptide substrates is required to ensure that functional groups are present for covalent immobilization of peptides on these platforms.

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2.3.1 Electrochemical Assays

Electrochemical assays rely on oxidative or reduction reactions on electrodes which cause measurable electrochemical signals.158 For electrochemical protease assays, a common approach involves covalent immobilization of peptide substrates on electrodes for protease sensing. Additionally, peptides are often labeled with redox reporters. For example, Ji et al. developed a thrombin assay using p- aminodiphenylamine (pADA)-labeled peptide substrates, where pADA acted as a redox reporter.38 Cleavage of peptide substrates decreased electrochemical signals due to the departure of the redox reporter (Scheme 2.3a). A similar approach was adopted by Shin et al. to detect MMP-9 using a methylene blue (MB)-labeled peptide substrate.159 Cleavage of peptide substrate by MMP-9 decreased electrochemical signals. In another example, Ko et al. used ferrocene (Fc), a common redox reporter, to label fibrinogen for detecting thrombin.41 Thrombin activities increased the density of Fc-labeled fibrinogen through coagulation, resulting in higher electrochemical signals (Scheme 2.3b). In order to further amplify the presence of protease activities, signal amplification techniques were investigated. For example, Wu et al. incorporated streptavidin-alkaline phosphatase

(ALP) at electrodes through biotin-labeled peptide substrates, and streptavidin-

ALP catalyzed the formation of electrochemically active phenol and increased initial electrochemical signals.45 Park et al. also incorporated a two-step enzymatic reaction to amplify the detection signals. For BoNT serotype E protease, the LOD could reach 2 pg/ml. Another amplification technique was demonstrated by Liu et al., who used electrodes functionalized with peptides containing repeated arginine

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residues to attract QD.42 Fluorescence arising from QD was then converted to measurable photocurrents. Thus, cleavage of these substrates by trypsin decreased photoelectrochemical signals. However, these techniques usually required synthetic labeling or binding of QD to peptide substrates, and this may adversely affect protease activities. Understandably, label-free substrates were desired to minimize such effects. For example, Cao et al. used peptides containing positively- charged arginine residue to prevent the interaction of electrodes with cationic electrochemical species in the aqueous phase.34 Therefore, cleavage of arginine by trypsin allowed electrochemical species to access the electrodes and increased electrochemical signals. A similar approach was also reported by Deng et al., who used peptide substrates that facilitated interaction of electrochemical species with the electrodes to produced large electrochemical signals.36 Cleavage of peptide substrates by PSA limited access of electrochemical species with the electrodes and decreased the measured signals, and they reported a LOD of 0.02 nM. This LOD is one order better than FRET-based PSA assay developed by Feng et al..84 Another interesting technique was demonstrated by Xia et al.; they used oxidation reaction catalyzed by peptide-heme complex to increase electrochemical signals.46 Thus, cleavage of peptide substrates by beta-site amyloid precursor protein cleaving enzyme 1 prevented the complex formation and decreased electrochemical signals.

Elsewhere, GO was also employed by Chen et al. to increase binding of MB near electrodes.35 This was achieved by using peptide-immobilized electrodes that promoted covalent attachment of GO upon cleavage by caspase-3, and the presence of GO attracted MB to increase electrochemical signals. They reported a

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remarkable LOD of 0.06 pg/ml (approx. 3 fM), which was among the lowest LOD reported. Given these recent developments in electrochemical-based protease assays, coupled with the ease of miniaturization of these assays, they are potential platforms for protease assays.

Scheme 2.3. Principles for Electrochemical-Based Protease Sensor

(a) “Turn –off” detection principle, where cleavage of immobilized peptide labeled with redox reporter decreases electrochemical signal. (b) “Turn-on” detection principle, where cleavage of immobilized peptide allow the binding of redox reporter to increase electrochemical signal.

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2.3.2 Label-Free SERS and SPR Assays

Label-free methods are attractive, as native peptides without additional tags are used as protease substrates. Thus, protease activities can be determined more accurately. SPR is a common label-free technique which can be used to monitor cleavage of peptide immobilized on a surface. In SPR, a beam of light with varying incident angles is impinged on a gold surface, and the reflected light is measured.

At a specific incidence angle, the absorbance of light is maximized whereas the reflected light is minimized. This incident angle is very sensitive to changes in surface properties. Therefore, it can be applied to detect cleavage of immobilized peptides by proteases. Such technique was demonstrated by Chen et al., who immobilized a highly cationic peptide substrate for detecting caspase-3.52 After the cleavage of peptides by caspase-3, the SPR angle decreased. By using this technique, they reported a LOD of 0.03 pM for caspase-3. SERS is another label- free technique that relies on Raman scattering by molecules adsorbed on rough metal surfaces. For example, Wu et al. used poly-arginine peptide to induce aggregation of AuNP and increased SERS signal.49 Cleavage of the peptide substrate by thrombin reduced the SERS signal.

2.3.3 Enzyme-Linked Peptide Protease Assays

Enzyme-linked peptide protease assays typically provide measurable signals through enzymes which are linked to peptide substrates. These enzymes catalyze the formation of colored compounds such that they are able to report the presence of peptide substrates (Scheme 2.4). For example, Ding et al. immobilized biotin-

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labeled peptides on a glass surface, and then conjugated the peptides to an enzyme,

ALP.160 This enzyme is able to remove a phosphate group from ascorbic acid 2- phosphate, generating a strong reducing agent ascorbic acid. Consequently, silver ions were reduced by ascorbic acid to form silver particles which were visible to the naked eye. In the presence of trypsin activity, substrates were cleaved and no silver particles were formed. Based on this principle, they reported a LOD of 0.4 nM trypsin. Cheng et al. also used silver deposition to produce optical signals for the detection of MMP-7, with a detection limit of 4.8 pM.161 Although these methods provide colorimetric responses, they require enzymes to generate the colorimetric response. Thus, specific conditions are required to carry out these enzymatic reactions, and this complicates the signal generation process.

Furthermore, the stability of secondary enzymes may require special attention to prevent any denaturation.

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Scheme 2.4. Principles of Enzyme-Linked Peptide Protease Assay

Formation of colored products from enzyme-linked peptides; absence of colored products indicates the presence of target protease.

2.3.4 LC-Based Assays

In LC-based assays, LC is used to examine solid surfaces immobilized with functional probes, which can be DNA, proteins, or peptides. Earlier, most LC-based assays were immunoassays, where antigens captured by antibody-coated surfaces caused changes to surface compositions.96, 101, 162 LC responded to such changes by altering its orientation near the surface, from homeotropic (perpendicular to the surface) to planar (parallel to the surface). Interestingly, changes at surfaces also influence LC in the bulk, and this was visualized as changes in LC optical signals.

More recently, Zhu and Yang demonstrated a method to quantify immunobinding events based on the length of bright LC optical signals in a microfluidic channel.163

These studies exemplified the feasibility of using LC-based assays to quantify

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changes in surface compositions. A similar concept was applied in LC-based protease assays, where peptide-immobilized surfaces caused LC to orient planarly and produce bright LC optical signals. In the presence of target proteases, peptides were cleaved and removed from the surface. Such surface composition changes resulted in dark LC optical signals. Based on this principle, Bi et al. developed an optical bar chart for semi-quantitative detection of protease activities.89 Cleavage of immobilized peptide substrates by trypsin resulted in dark LC optical signals, and they reported a LOD of 2 nM (Scheme 2.5). Later on, Chen and Yang also used a similar approach to detect trypsin and chymotrypsin activities, and they reported a LOD of 0.04 nM and 4 pM, respectively.104 These studies demonstrated the potential of carrying out heterogeneous protease assays that are capable of multiplex sensing. Furthermore, LC-based assays are not susceptible to long-term stability and photo-bleaching issues. However, these LC-based protease assays relied solely on immobilized peptides to influence the LC signals.89, 109, 164 To improve LOD of these assays, incorporation of an amplification mechanism is required to enhance changes in LC optical signals after cleavage of immobilized peptides.

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Scheme 2.5. Principles of LC-Based Protease Assay

Different LC optical texture in the presence of target protease. Peptide substrates are immobilized on glass substrates, and cleavage of peptide substrate changes surface composition.

Elsewhere, studies using LC/water interfaces were also studied as a possible sensing platform for protease activities. Such systems provide real-time detection of protease activity, since the LC interface can be monitored in real time with proteases in the aqueous phase. For example, Zhang and Jang reported the use of poly-L-lysine decorated LC/water interface for the detection of trypsin activity.129

They showed that the cleavage of poly-L-lysine at the LC/water interface resulted in a bright-to-dark transition in LC texture. To improve the selectivity of LC sensors at LC/water interface, Chang and Chen reported the detection of chymotrypsin activity using customized peptide substrates.165 They showed changes in LC texture of LC droplets in the presence of chymotrypsin activities, and the LOD was 0.02

M. Compared to detection at glass surfaces, the advantage of such LC/water interface detection is the real-time detection of protease activity. However, more work may be required to establish the robust immobilization of peptide substrates at LC/water interfaces.

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By comparing LC-based assays against other assay techniques, LC assays provide straightforward optical readouts that are photo-stable and generated based on interactions of LC with functionalized surfaces. This is in contrast to fluorescence detection methods, which are photo-sensitive and instability of the fluorophores used can often cause false results. Therefore, the LC-based assay provides a sensitive detection platform that is not susceptible to photo-instabilities commonly associated with fluorescence-based techniques. In relation to these issues of fluorescence-based techniques, optical absorbance of functionalized, inorganic nanoparticles were also discussed in this review as viable protease detection methods. However, aggregation of these nanoparticles due to fluctuations in pH, temperature, or salt contents of the samples can cause false results and limit their wide applications in real samples. Furthermore, these assays are usually carried out homogenously, and this restricts their applications in multiplexed assays. In these aspects, LC-based assays are not sensitive to fluctuations in physical parameters like pH or temperature. Importantly, due to their heterogeneous assay formats, multiplex detections are readily achievable for LC-based assays. Other detection methods include SPR and SERS. Although these techniques are highly sensitive, quantitative, and capable of real-time monitoring of protease activities, they require highly intricate optical setups. Therefore, they are restricted to applications in a controlled environment of a laboratory. In this aspect, LC-based assays require minimal instrumentation for straightforward and robust optical readouts that can be deployed in resource-limited regions. In summary, LC-based assays provide

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straightforward optical readouts, and they are portable, sensitive, robust, and capable of multiplexed detections. This makes such assays highly attractive.

Table 2.2 summarizes some of the recently developed heterogeneous trypsin and chymotrypsin assays based on concentrations of proteases as a common unit for the comparison of LOD. LC-based assays by Chen and Yang provided the best LOD for chymotrypsin (4 pM) and trypsin (0.04 nM).104 In this assay, peptides with multiple lysine residues were required for stable peptide immobilization. Because these peptides can be cleaved by trypsin or other proteases, the specificity of the assay was questionable. Therefore, further work is required for the development of heterogeneous LC-based protease assay with better selectivity and sensitivity.

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Table 2.2. Summary of Heterogeneous Trypsin and Chymotrypsin Protease Assays Recently Developed Reference Detection Protease LOD Principle Substrate a

42 Liu et al. Electrochemical CAPGGAR5 Trypsin (3 nM)

Cao et al. 34 Electrochemical RGGLAC Trypsin (0.6 nM)

Ding and Colorimetric PPLRINRHILT Trypsin (0.4 nM) 160 Yang RG3-Biotin Bi et al. 89 LC CDRVYIHPFH Trypsin (2 nM) LK Chen and LC CKGSNRTRID Trypsin (0.04 nM) Yang 104 EGNQRATRM

LGGKETSAA

KLKRKYWW

CLSELDDRA Chymotrypsin (4 pM)

DALQAGASQ

FESSAAKLKR

KYWWKNLK a Peptide sequences follow the one-letter code for amino acids

2.4 Outlook

Currently, there are many types of protease assays under development, but none of them are ideal. For homogenous assays, even though very low detection limits

(~pM) can be achieved, they suffer from issues such as limited stability of fluorescent molecules. To overcome this, inorganic QD and AuNC are exploited to

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provide additional stability. Besides fluorescence-based techniques, colorimetric assays with AuNP are widely studied due to its simplicity. However, due to the inherent instability of AuNP, these assays are susceptible to high salt concentrations, extreme pH, and temperature. On the other hand, heterogeneous assays provide a multiplexed platform for the detection of proteases. However, since protease substrates are typically immobilized on surfaces, only proteases near these surfaces interact with these substrates. Therefore, the sensitivity of heterogeneous assays is low. To overcome this, immobilized peptides on microparticles with a high surface-to-volume ratio can be used to improve interactions between proteases and peptides for better sensitivity. In the future, LC- based assays may emerge as an alternative platform which provides fast, sensitive and multiplexed detection of proteases. However, challenges in the LC-assays including stability of immobilized peptide substrates, the sensitivity of the assay, and their limited quantitative detection capabilities need to be addressed.

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CHAPTER 3 MECHANISTIC STUDY FOR

IMMOBILIZATION OF CYSTEINE-LABELED

OLIGOPEPTIDES ON UV-ACTIVATED SURFACES

DMOAP-coated surfaces are readily applied in LC-based assays. However, the mechanism involved in the immobilization of oligopeptides is still unclear. To address this, in this chapter, we aim to understand the mechanism for oligopeptide immobilization on ultraviolet- (UV)-activated, N, N-dimethyl-n-octadecyl-3- aminopropyltrimethoxysilyl chloride- (DMOAP)-coated surfaces. As such, this study will provide the fundamental understanding of oligopeptide immobilization through cysteine residue. This way, future oligopeptides can be optimally designed for LC-based assays.

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3.1 Introduction

Short oligopeptides have been increasingly used in microarrays and biosensors as molecular receptors in recent years due to several reasons. First, similar to antibodies, oligopeptides can bind to specific targets including small molecules, ions,166-167 explosives,168-170 and pesticides,171 and form a stable complex. Secondly, most short oligopeptides do not have quaternary structures such that they are less susceptible to environmental factors like temperature and pH. Lastly, high- throughput screening, like phage display technique, allows one to identify unique oligopeptide sequence for a particular target compound. After the sequence is identified, the oligopeptide can be custom-made by using solid-phase synthesis with high purity. Desired functional groups or amino acids (such as His-tag) can also be incorporated during this process. However, to use oligopeptides for sensor applications, oligopeptides need to be immobilized on a solid surface with controlled orientation and a high density.105-107 The former is important because specific binding motifs need to be exposed for binding target molecules.172-174 A high density of oligopeptide is desirable because it can increase the binding of target molecules to the surface and lower the detection limit.175-176

In the literature, immobilization of oligopeptides on an aldehyde-functionalized surface can be accomplished through a reaction between N-terminal cysteine and aldehyde groups, as reported by Liu et al..177 Using the immobilized oligopeptide with an N-terminal cysteine group, Bi et al. developed a protease assay in which the immobilized oligopeptide is cleaved by trypsin.111 They also showed that a

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reaction between the lysine group on the oligopeptide and surface aldehyde groups can be kinetically excluded under the experimental conditions. Meanwhile, Su et al. and Xiao et al. made use of cysteine residues for immobilization of oligopeptides on solid surfaces functionalized with maleimide groups, where maleimide-thiol reaction took place between the thiol group of cysteine and surface maleimide group.178-179 Notably, the approach of using thiol-ene reaction for oligopeptide immobilization is becoming more popular because of its robust reaction conditions and fast reaction rates.180-183 Another advantage is that this reaction can be activated by ultraviolet (UV) light to spatially immobilize oligopeptides. However, these immobilization techniques require prior surface modifications to present reactive groups necessary for coupling reactions.

N, N-dimethyl-n-octadecyl-3-aminopropyltrimethoxysilyl chloride (DMOAP) is an organosilane that presents an 18-carbon chain and a quaternary ammonium salt (see

Figure 3.1a). The positive charge on the quaternary ammonium makes DMOAP soluble in water. In the past, glass slides coated with DMOAP are widely used in

LCD to control the orientations of liquid crystals.93, 108 Recently, DMOAP-coated slides have also been used for immobilization of oligonucleotides and oligopeptides.89, 109 For example, Chen and Yang used DMOAP-coated surfaces for the fabrication of LC assays to detect protease activities.109 Because DMOAP does not have any reactive groups, the surface needs to be activated by UV before oligopeptides can be immobilized.109-110 Nevertheless, the exact mechanism of surface activation and oligopeptide immobilization on the DMOAP-coated surface

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are still unclear. Moreover, oligopeptides immobilized on the surface are unstable under rigorous washing conditions.

Figure 3.1. (a) The structure of DMOAP. (b) Proposed surface structure presenting aldehyde groups after UV irradiation.

In this study, we aim to understand the role of cysteine during the immobilization of oligopeptides on a DMOAP-coated surface. We first synthesized a set of model oligopeptides with a cysteine residue at various positions to better understand the role of cysteine during the immobilization. An acetyl group was also introduced to protect terminal amine, preventing it from reacting with surface functional groups.

Secondly, we studied the stability of these immobilized oligopeptides by incubating oligopeptide-functionalized slides in buffer solutions under stringent conditions.

The stability of the immobilized oligopeptides is crucial in the sensing applications, especially when repetitive washing steps are required. Finally, we investigated whether oligopeptides immobilized under optimized conditions bind to their target molecules with higher specificity and capacity.

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3.2 Experimental Section

3.2.1 Materials

Glass slides were purchased from Fisher Scientific (U.S.A.). Sodium bicarbonate, sodium cyanoborohydride, DMOAP, dimethyl sulfoxide (DMSO), polyoxyethylene (20) sorbitan monolaurate (Tween 20), 4-amino-5-hydrazino-1, 2,

4-triazole-3-thiol (Purpald), human serum, 2, 5-dihydroxybenzoic acid (DHB) and

5, 5’-dithiobis-(2-nitrobenzoic acid) (DTNB) were purchased from Sigma-Aldrich

(Singapore). Phosphate buffer saline (PBS) and Tris buffer were purchased from

First BASE (Singapore). Cy3-labeled streptavidin was purchased from Life

Technologies (Singapore). Calcium chloride and silica thin layer chromatography

(TLC) plates were purchased from Merck (Singapore). Trimethoxysilane aldehyde

(TEA) was purchased from United Chemicals Technologies (U.S.A.).

Oligopeptides (purity>95%) were synthesized by GenicBio Ltd (China). These oligopeptides include Ac-CGGK-FITC (P1), GGGK-FITC (P2), Ac-GGGK-FITC

(P3), CGGK-FITC (P4), GGCK-FITC (P5), CGGGWDPYSHLLQHPQ (P6)

GGGWDPYSHLLQHPQC (P7) and GGGWDPYSHCLLQHPQ (P8).

3.2.2 Surface Modifications

To clean the surface, glass slides were immersed in 5% (v/v) Decon 90 solution overnight. After this, the glass slides were sonicated thrice for 15 min in deionized

(DI) water. Subsequently, the glass slides were immersed in an aqueous solution containing 0.1% (v/v) DMOAP for 5 min, and then rinsed with copious DI water.

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DMOAP-coated glass slides were dried under a stream of nitrogen, followed by heating in a 100 C vacuum oven for 15 min. Water contact angle was measured to ensure the successful coating of DMOAP on the glass surfaces (see Appendix A.1).

For UV activation, the slides were placed 3 cm from a UV pen lamp (254 nm,

Spectroline®, model 11SC-1), and exposed to UV for 50 s.109 In the surface reduction experiment, the surface was immersed in carbonate buffer (100 mM, pH

10) containing 50 mM of sodium borohydride after the UV irradiation.

3.2.3 Immobilization of Oligopeptides

Solutions containing oligopeptides (10 µM for P1 to P5, 100 µM for P6 to P8) and

0.001% (v/v) of Tween 20 were dispensed on a UV irradiated surface by using a micropipette. Tween 20 was introduced to reduce non-specific adsorption of oligopeptides on the surface. The slide was then incubated in a dark and humidified chamber at room temperature for 3 h. The immobilization duration of 3 h was used based on results presented in Appendix A.2. After immobilization, the slide was dipped into 0.05% (v/v) Tween 20 solution for 5 s and rinsed with DI water. For P6 to P8, the increased oligopeptide concentration was aimed at increasing the surface density of immobilized oligopeptide to achieve better detection signals.

3.2.4 X-ray Photoelectron Spectroscopy (XPS)

Surface chemical composition on the DMOAP-coated glass surface was studied by using XPS (Kratos AXIS UltraDLD). The X-ray source is Al Kα (1486.71 eV, 75

W), and the pressure was kept at 5×10-9 Torr. The sample was cut into a 5 mm × 5

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mm piece, adhered to a standard stub and positioned 90° to the detector. Peak positions were calibrated by using C1s peak at 285.0 eV.

3.2.5 Stability Test for Immobilized Oligopeptides

Oligopeptide-immobilized slides were immersed in Tris buffer (50 mM Tris-HCl, pH 8.0) containing 20 mM of calcium chloride and 0.001% (v/v) of Tween 20.

Calcium chloride was added as it is commonly used to improve the stability of proteases in protease assays. The solution was maintained at 37 C in a water bath for 2 h. After the incubation, the slides were rinsed with copious DI water and dried under a stream of nitrogen gas.

To assess the stability of oligopeptide-immobilized slides in complex biological mediums, we immersed a slide with immobilized P6 in dilute human serum for 2 h at 37 C. Tris buffer (50 mM Tris-HCl, pH 8.0) was used as the diluent. After the incubation, the slides were rinsed with a copious amount of water and dried under a stream of nitrogen gas.

3.2.6 Purpald Test

To detect surface aldehyde groups, a purpald test was carried out on a TLC plate due to its high surface area and white background that allows easy visual detection of any color changes. Purpald solution was 1.0 M of sodium hydroxide solution containing 2% (w/v) of purpald. 20 µl of DMOAP was dropped onto the TLC plate,

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and the spot was allowed to dry in air for 10 min. After this, 50 s of UV irradiation was carried out on the surface. The purpald solution was dropped onto the TLC plate and left for 5 min at room temperature. Visual color changes of purpald solution to purple would indicate the presence of aldehydes on the surface.

3.2.7 Streptavidin Binding Assay

P6 to P8 were immobilized as described above. 0.6 μl of a solution containing 20

g/ml of streptavidin-Cy3 was spotted on an oligopeptide microarray. The spot was aligned at the same position as the oligopeptide spot. The slide was incubated for

30 min in a humidified chamber to allow the binding of strepatavidin-Cy3 to the oligopeptide. To remove unbound streptavidin-Cy3, the slide was washed briefly

(~5 s) by using a 0.05% (v/v) Tween 20 solution and DI water, respectively. Finally, the slide was dried under a stream of nitrogen gas. Scheme 3.1 shows procedures of this assay.

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Scheme 3.1. Procedure for Streptavidin Binding Assay

(a) Oligopeptide immobilization, (b) introduction of streptavidin-Cy3 to the same position where oligopeptide was immobilized, and (c) fluorescence scanning for detecting streptavidin-Cy3.

3.2.8 Oxidation of Thiols

For the oxidation of P6, 50% (v/v) of oligopeptide (500 μM) in DMSO was incubated at 37 C before diluting with carbonate buffer (100 μM, pH 10) to 100

µM for immobilization on the glass slide.184

3.2.9 Fluorescence Microscopy

Fluorescence data presented in this study were obtained using a fluorescence scanner (GenePix, 4100A) manufactured by Molecular Devices (U.S.A.). During a typical scan, photomultiplier tube (PMT) gain = 500 and pixel size = 20 μm. Ten readings were used to obtain an average intensity for each immobilization condition. For each reading, a circular area with 1 mm in diameter was used to

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obtain the fluorescence intensity using software (GenePix Pro 6.0) provided for the equipment.

3.2.10 Matrix-Assisted-Laser-Desorption-Ionization Time-of-Flight Mass-

Spectrometry (MALDI-TOF-MS)

Samples were prepared using 2, 5-dihydroxybenzoic acid (DHB) matrix with 1:1 mixing ratio. For this characterization, 1 μl of products from the DMSO oxidation reaction was mixed with 1 μl of DHB matrix. The experiments were carried out by using AutoFlex II (Bruker, U.S.A.).

3.2.11 DTNB Assay

In this assay, 2 mM of P6 was mixed with equal volume of DMSO. The increased concentration of P6 was used to achieve greater contrast in the absorbance values before and after the oxidation process. After the oxidation reaction, 15 μl of the reaction solution was added to 285 μl of 0.1 mM DTNB and incubated for 5 min.

The absorbance readings at 412 nm were recorded using microplate reader Infinite

200 PRO (Tecan, Switzerland), and cysteine was used as a standard for calibration.

3.3 Results and Discussion

3.3.1 Role of Cysteine in Oligopeptide Immobilization

To understand the role of cysteine for the immobilization of oligopeptides, we dissolved 5 oligopeptides (P1-P5) in carbonate buffer (100 mM, pH 10) and

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immobilized them on a UV-activated surface. As shown in Table 3.1, P1 and P4 have a cysteine at the N-terminus whereas P5 has a cysteine in the middle. P2 and

P3 do not have any cysteine. Moreover, the N-terminal amine groups in P1 and P3 were protected with an acetyl (Ac) group to determine whether free amine groups also affect oligopeptide immobilization. Figure 3.2a shows that only P1, P4, and P5 can be immobilized on the surface whereas P2 and P3 cannot. These results clearly indicate the importance of cysteine for the immobilization of oligopeptide on the surface. Moreover, even though P2 has an N-terminal amine, it cannot be immobilized on the surface. For P1, the immobilization was not achieved through the formation of a thiazolidine ring between thiol and amine groups.111, 177 This result suggests that N-terminal amine does not react with surface functional groups.

By assessing potential reactive groups on P1, P4 and P5, it can be seen that the presence of thiol alone, regardless of its position, can result in the immobilization of oligopeptides. This is in contrast to previous work in which only N-terminal cysteines were immobilized on aldehyde-terminated surfaces without reducing agents.111 These immobilized oligopeptides are very stable. Even after incubation in 0.001% (v/v) of Tween 20 at 37 C for 2 h, the fluorescence intensities only varied by less than 1% compared to initial readings. Finally, we note that in the case of P1, the lack of free N-terminal amine does not result in lower fluorescence intensity than P4.

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Table 3.1. Sequences of Model Oligopeptides Functional Oligopeptides Sequences a groups Ac-CGGK- P1 -SH (FITC) GGGK- P2 -NH2 (FITC) Ac-GGGK- P3 - (FITC) CGGK- P4 -SH , -NH2 (FITC) GGCK- P5 -SH, -NH2 (FITC) CGGGWDPY P6 -SH, -NH2 SHLLQHPQ GGGWDPYS P7 -SH, -NH2 HLLQHPQC GGGWDPYS P8 -SH, -NH2 HCLLQHPQ a Acetyl (Ac) group is introduced as a blocking group for N-terminal amine in P1 and P3; fluorescein 5-isothiocyanate (FITC) is introduced as a fluorescent marker; oligopeptide sequences are named according to the one-letter code for amino acids.

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correspond to P1 to P5. (d) Comparison of fluorescence data for oligopeptide immobilization using the three different buffers in (a) to (c).

3.3.2 Effects of Reducing Agent and Buffers

To understand the effects of reducing agent and buffers on oligopeptide immobilization, two additional immobilization buffers were used. The first one is carbonate buffer (100 mM, pH 10) with additional 10 mM of sodium cyanoborohydride. This reducing agent can reduce imines to form stable secondary amines. The second one is PBS buffer (100 mM, pH 7). The choice of PBS is based on a previous report showing that N-terminal cysteine and a surface aldehyde can react in this buffer.111 Figure 3.2b and 3.2d show that the reducing agent, sodium cyanoborohydride, does not result in any difference in fluorescence intensity. Thus, we can conclude that the reducing agent has minimal effect on the oligopeptide immobilization. In PBS buffer, however, P2 and P3 have higher fluorescence

(4196±460 and 2285±255) despite the absence of cysteine in P2 and P3 (see Figure

3.2c,d). Thus, in PBS buffer, it is possible to immobilize oligopeptides which do not have any cysteine residues. On the basis of these observations, we conclude that immobilization of oligopeptides in carbonate buffer results in well-controlled immobilization of cysteine-containing oligopeptides. On the other hand, immobilization reaction carried out in PBS buffer results in nonspecific immobilization of oligopeptides, regardless of the presence of cysteine residues.

Therefore, further experiments were carried out using carbonate buffer only, as this immobilization buffer provides stable and specific oligopeptide immobilization through cysteine residues only.

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3.3.3 Effect of UV Irradiation

To study the effect of UV irradiation duration on the density of oligopeptide immobilization, we prepared DMOAP-coated surfaces exposed to 0 s, 20 s, 50 s,

120 s, 180 s, and 300 s of UV irradiation. Oligopeptides P1, P4, and P5 were immobilized on these surfaces and their fluorescence intensities were measured and presented in Figure 3.3. From the trends observed in Figure 3.3, there is an optimum

UV irradiation duration at 50 s that results in the highest density of immobilized oligopeptide. For oligopeptides P2 and P3, fluorescence intensities remained low compared to P1, P4 and P5 (see Appendix A.3). Characterization of the surfaces after UV irradiation by using XPS was carried out. Figure 3.4 shows that the C1s shoulder peaks evolve with the increasing UV irradiation time. As shown in past studies, these peaks can be assigned to alcohol (C-OH ~286.5 eV), aldehyde

(HC=O ~288 eV) and carboxylic acid (HO-C=O ~289.2 eV),110 respectively. It can be observed that with increasing UV irradiation, more oxidized products were formed. In Figure 3.4b, where the samples were irradiated with UV for the optimum duration of 50 s, two shoulder peaks assigned to alcohol (B.E. ~286.5 eV) and aldehyde (B.E. ~288 eV) appear, suggesting that either aldehyde or alcohol group is responsible for the immobilization of oligopeptide. With prolonged UV irradiation up to 5 min, more alcohol and aldehyde groups are further oxidized to carboxylic acids, and this is seen from the emergence of the peak corresponding to the carboxylic acid group (B.E. 289.2 eV). By comparing the results in Figure 3.3 and 3.4, we conclude that surface aldehyde and alcohol groups are generated after

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50 s of irradiation, while surface carboxylic acid groups were observed after 180 s of irradiation.

Figure 3.3. Fluorescence intensity of immobilized oligopeptide for different UV exposure durations: (a) P1, (b) P4 and (c) P5.

Figure 3.4. XPS results of DMOAP-coated glass surface illustrating the emergence of C (1s) shoulder peaks with increasing (254 nm) UV exposure duration: (a) 0 s, (b) 50 s, (c) 180 s and (d) 300 s.

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3.3.4 Test for Surface Aldehyde Group

To examine whether aldehyde groups were produced after optimum UV irradiation of 50 s, purpald test was carried out by using a TLC plate coated with DMOAP.

TLC plate was used because of two reasons. First, because of its large surface area, the color change that happens on TLC plate is more observable than that on a glass slide. Secondly, the background color of TLC is white, and that provides a good contrast to color changes from the purpald test. From Figure 3.5, we can see purple color after the surface was irradiated with UV. This result confirms the production of aldehydes on the DMOAP surface after UV irradiation. In contrast, the TLC plate remained white if the surface was not irradiated with UV.

Figure 3.5. Purpald test results: (a) 50 s UV irradiated DMOAP-coated TLC plate, (b) no UV irradiation on DMOAP coated TLC plate.

To further confirm whether the aldehyde or alcohol group is responsible for immobilization of oligopeptides, we reduced DMOAP-coated surface with NaBH4 after UV irradiation. Then, oligopeptide immobilization was carried out using carbonate buffer (100 mM, pH 10). If oligopeptide immobilization requires surface aldehyde groups, the surface reduction will remove all aldehyde groups and greatly decrease the immobilization efficiency. The fluorescence intensity dropped from

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3467±814 to 702±74 for P1, 3972±466 to 532±64 for P4 and 3920±329 to 621±76 for P5. This result strongly suggests that the presence of aldehyde groups is responsible for oligopeptide immobilization. Thus, we propose that the UV- irradiated surface is aldehyde-terminated as shown in Figure 3.1b.

3.3.5 Role of Cysteine in Immobilization Mechanism

After the role of surface aldehyde groups is confirmed, we carried out two additional experiments to investigate the role of cysteine in oligopeptide immobilization and the immobilization mechanism. In these experiments, we used a new oligopeptide P6 with an HPQ motif which can bind to streptavidin with high specificity. The choice of P6 and the new binding assay is because variations of pH and temperature during experiments may affect the fluorescence intensity of FITC- labeled oligopeptide. In our first experiment, we study whether the free thiol group is indeed involved in a reaction that leads to immobilization of oligopeptide. In this experiment, the thiol group on P6 was oxidized to disulfides in 50% (v/v) DMSO solution at 37 C to deactivate its reactivity.184 The extent of oxidation was controlled by varying the incubation time from 0 to 4h and quantified by using

DTNB assay (see Appendix A.3). After which, streptavidin binding assay was carried out to assess the density of oligopeptide immobilized on the surface. Figure

3.6a shows that the fluorescence intensity decreases as the extent of P6 oxidation increases until 20% of thiol oxidation. Beyond 20% of thiol oxidation, the intensity values are close to those measured for un-modified surfaces (see section 3.3.6).

Thus, the presence of free thiols in cysteines is required for the immobilization of

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oligopeptides. To characterize the oxidation products, MALDI-TOF-MS was carried out at different oxidation durations. From Figure 3.7, two peaks at 1794-

1795 and 3587-3588 agree with the molecular weight of P6 (1793.97) and the dimer of P6 oligopeptide linked together with a disulfide bond.

Figure 3.6. Fluorescence intensities for streptavidin binding assay used to examine the role of cysteine in oligopeptide immobilization: (a) effect of thiol oxidation, (b) effect of pH.

The second experiment was to vary the buffer pH used during oligopeptide immobilization. Because pKa of thiol is 8.38, we varied the pH of carbonate buffer

(100mM) from pH 7 to 11 during the immobilization of P6. After which, streptavidin binding assay was carried out to assess the density of immobilized oligopeptide. Figure 3.6b shows that the fluorescence intensity increases when the pH is increased from 7 to 10. However, it decreases when the pH value is further increased to 11. The decrease in fluorescence intensity at pH 11 can be attributed to the instability of P6 at higher pH. Interestingly, there is a noticeable increase in fluorescence intensity between pH 8.4 and 9, which corresponds to pKa of thiol.

The increase in fluorescence intensity implies that the immobilization of oligopeptide prefers negatively-charged thiolate ions over thiol group. These

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results, when combined, suggest that the oligopeptide immobilization is realized through nucleophilic addition of the thiolates to surface aldehyde groups. This is supported by the efficient immobilization using carbonate (100 mM, pH 10) buffer, where the basic pH can catalyze the nucleophilic addition reaction by promoting the formation of thiols to the more nucleophilic thiolates (see Scheme 3.2). Under this condition, only cysteine-containing oligopeptide can be immobilized on surfaces presenting aldehyde group. Although the acid-catalyzed formation of thioactals has been reported,185 our results suggest that the immobilization is base- catalyzed. Additionally, the immobilization reaction probably stops after the formation of thioacetals, as shown in Scheme 3.2. Addition of a second thiolate ion to form dithioacetals is unlikely because alcohol is a poor leaving group. This proposed immobilization mechanism also suggested that NH2 did not aid in the immobilization of oligopeptides through reductive amination.186 Additionally, the role of N-terminal cysteine in the formation of thiazolidine ring was also not observed for UV-irradiated DMOAP surfaces.111

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Figure 3.7. Mass spectrometry results for the product of oxidation in DMSO at different durations: (a) 0 h, (b) 1 h, (c) 2 h, (d) 3 h and (e) 4 h.

Scheme 3.2. Proposed Mechanism for Oligopeptide Immobilization

A proposed mechanism for nucleophilic addition of thiols to aldehydes: (a) a thiol group deprotonates at pH = 10, (b) nucleophilic addition of a thiolate ion to the aldehyde group, (c) overall reaction scheme for immobilization mechanism.

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3.3.6 Stability and Orientation of Oligopeptide Immobilization

To investigate whether the orientation of oligopeptide immobilization can be controlled by the position of a cysteine residue, we selected oligopeptides P6, P7, and P8. All three oligopeptides contain a streptavidin binding motif, HPQ, and a cysteine residue at different positions. For P6, the cysteine is located at the N- terminus, whereas for P7, the cysteine is located at the C-terminus for P7. For P8, cysteine is positioned in the middle of the oligopeptide.

After immobilization of oligopeptides, streptavidin binding assay was carried out to assess the surface density and orientation of immobilized oligopeptides. For P6, the fluorescence intensity was 4287±233, suggesting that streptavidin-Cy3 can bind to the immobilized P6 effectively through HPQ. In contrast, the fluorescence intensities for P7 and P8 are 647±33 and 728±29, respectively. These values are much lower than that for P6. In fact, these values are even lower than a control experiment (fluorescence = 1129±60) in which no oligopeptide was immobilized on the surface. These data indicate that the position of cysteine affects the binding of streptavidin. One possible reason is that the oligopeptide immobilized through the N-terminal cysteine can expose the streptavidin binding motif (HPQ) located at the C-terminus. In contrast, the cysteine residue of P7 is located at the C-terminus.

Therefore, immobilization of P7 through the cysteine residue will hinder the interaction between HPQ and streptavidin-Cy3. For P8, the cysteine residue in the middle of the oligopeptide and that can result in a surface presenting both C- terminus (containing HPQ) and N-terminus. This surface reduces the density of

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HPQ motif interacting with streptavidin-Cy3, and the presence of the N-terminus may also interfere with the binding of streptavidin. Also, the density of P8 may be lower than in P6 and P7 considering the geometry of immobilized oligopeptide. As shown in Scheme 3.3, anchoring points in oligopeptides P6 and P7 are located either at the N-terminus or C-terminus. Therefore, immobilized oligopeptides can be viewed as closely packed straight chains on the surface. For P8, the immobilization through the cysteine in the middle hinders the packing of the oligopeptides, as each oligopeptide may take up a larger surface area. The control experiment of incubating streptavidin-Cy3 on the un-modified surface also produced a relatively low fluorescence, and this suggests low levels of non-specific binding by streptavidin-Cy3 on the substrate. From this experiment, we show that the orientation of the oligopeptide can be controlled by the location of the cysteine residue. We can maximize the binding of target protein to the surface by including a cysteine at the terminus opposite to the binding motif.

To study the stability of oligopeptide-immobilized surfaces in complex biological medium, we incubated P6-immobilized slides in human serums of 20×, 50× and

100× dilution factors. After the incubation, we introduced streptavidin-Cy3 to compare the binding capability of these surfaces with the results presented above.

We observed that the fluorescence intensities were 43%, 56% and 74% for dilution factors of 20×, 50× and 100× respectively. The decrease in fluorescence intensities can be attributed to the non-specific bindings arising from the complex medium.

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This may cause some immobilized P6 to be non-specifically bound to other proteins in the human serum, thus preventing them from binding with streptavidin-Cy3.

Scheme 3.3. Oligopeptide Immobilization Density and Orientation

Oligopeptides immobilization through cysteine residues in P6 to P8, respectively. (a) Optimum immobilization density and orientation of P6 for binding with streptavidin-Cy3. (b) Immobilization orientation of P7 that restricts the HPQ binding motif’s interaction with streptavidin-Cy3. (c) Oligopeptide immobilization of P8 that presents two different oligopeptide segments on the surface, and this reduces oligopeptide density and interaction of HPQ binding motif with streptavidin-Cy3.

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3.4 Conclusion

A cysteine residue, regardless of its position, is needed for oligopeptide immobilization on a UV-irradiated DMOAP surface. The UV irradiation produces surface aldehyde group, which reacts with cysteine and leads to oligopeptide immobilization on the surface. A possible reaction mechanism is the nucleophilic addition of thiols to surface aldehyde groups. This study provides a new method to immobilize oligopeptide on a DMOAP surface with good stability. Immobilized oligopeptides with specific binding motif can also bind to its target proteins, such as in the case of HPQ motif binding with streptavidin.

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CHAPTER 4 LIQUID CRYSTALS-BASED PROTEASE

ASSAYS WITH DUAL CYSTEINE-LABELED PEPTIDES FOR

IMPROVING PEPTIDE STABILITY

Liquid crystal- (LC)-based, heterogeneous assays that produce optical signals in response to protease activities are attractive. However, in these assays, issues such poor stability of immobilized peptides and difficulties in detecting cleavage of immobilized peptides remain. Based on the immobilization technique developed in

Chapter 3, we further improved the stability and applicability in LC-based assays by introducing dual cysteine and biotin-labeled peptides as protease substrates.These peptides, upon immobilization, exhibit higher surface densities and better stability than single cysteine-labeled peptides due to the formation of inter-peptide disulfide bonds. Meanwhile, the biotin moiety can bind to streptavidin and create substantial changes in surface properties, which can be detected with a thin layer of LC supported on the surface. This work aims to develop a proof-of- concept for a reliable and generic approach for LC-based protease assay.

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4.1 Introduction

Proteases are enzymes that catalyze the cleavage of specific peptide bonds in proteins or peptides. For example, trypsin cleaves peptide bonds at the carboxyl (C-

) terminal of positively-charged lysine and arginine.15 On the other hand, chymotrypsin cleaves peptide bond at the C-terminal of hydrophobic tryptophan, tyrosine, and phenylalanine.15 Proteases serve a whole range of biological functions, from blood clotting7 to food digestion.2 They also play important roles in cancers,3 cardiovascular diseases,4 Alzheimer’s disease,5 human immunodeficiency virus (HIV),6 thrombosis,7 and diabetics.8 Therefore, there is a strong motivation for developing protease assays which are able to detect proteases and their inhibitors. In general, protease assays can be classified as non-functional or functional assays. In non-functional protease assays, detection principles are based on the recognition of proteases by antibodies. For example, enzyme-linked immunosorbent assays (ELISA) were used to detect proteases through antibody- antigen recognition with the aid of secondary enzyme-linked antibodies for color generation.187-189 However, such assays cannot be used to distinguish active and inactive proteases. In contrast, functional protease assays detect enzymatic activities of proteases. Such assays are often designed bases on techniques such as bioluminescent resonance energy transfer (BRET),190-194 upconverting phosphors

(UCP),33, 195-198 and fluorescence energy resonance transfer (FRET).73, 76-77, 82, 139-

140, 146, 151 However, these assays are homogeneous assays, where both substrates and samples are present in an aqueous phase. Such homogenous assays are

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generally not capable of multiplex sensing. This issue arises due to overlapping of emission spectra of fluorescent probes.

In contrast, heterogeneous assays are more suitable for multiplex detection of proteases. In heterogeneous assays, a large number of peptides are first immobilized on a solid surface at discrete locations. Subsequently, the solid surface is immersed into an aqueous solution containing proteases which cleave peptides and remove peptide fragments from the surface. Next, mass spectrometry or surface plasmon resonance (SPR) can be used to detect cleaved peptides on the surface or peptide fragments in the solution.21, 199 Such detections can be performed at discrete locations on the surface, allowing multiplex detection in a single assay. For example, Hu et al. reported a multiplex protease assay by using matrix-assisted laser desorption/ionization-time-of-flight (MALDI-TOF) mass spectrometry to detect peptide fragments cleaved by target proteases.21 Due to the high sensitivity of MALDI-TOF, trypsin and chymotrypsin were detected with limits of detection at 2.3 and 5.2 nM, respectively. However, mass spectrometry detections require extensive and costly instrumentation. Surface spectroscopy methods, like surface plasmon resonance (SPR) and surface enhanced Raman spectroscopy (SERS), were also employed in heterogeneous protease assays. In such assays, cleavage of immobilized peptide substrates causes shift in surface plasmon signatures.48-54

However, their sensitivities are affected by poor signal-to-noise ratios in complex samples.200 Therefore, heterogeneous protease assays which provide simple optical outputs (e.g. visible to the naked eye) are highly desirable. An example is enzyme-

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linked peptide protease assay reported by Ding and Yang.160 In this assay, an array of biotin-labeled peptide substrates were immobilized on solid surfaces. In the absence of target protease, alkaline phosphatase-linked streptavidin (streptavidin-

ALP) bound to the surface, and ALP caused reduction of silver ions to silver particles. On the other hand, target proteases removed the biotin moiety and stopped the enzymatic reactions. Cheng et al. also developed a scanometric assay to detect matrix metalloproteinase-7, which is an important protease in cancer metastasis.161

In this assay, oligopeptides were conjugated to gold nanoparticles and immobilized on glass surfaces. Subsequently, gold nanoparticles on the surfaces induced the deposition of silver particles. These techniques provide facile signal readouts, while the heterogeneous assay formats provide multiplex sensing capabilities.

Alternatively, liquid crystals (LC) can be used as a simple signal transduction mechanism without secondary enzymatic reactions. LC-based assays rely on unique optical properties and orientational behaviors of LC. Molecules which form an LC phase are anisotropic and birefringent, allowing them to produce unique optical textures under a cross-polarized light source.86 Moreover, when these molecules are in contact with a solid surface, their orientations are highly sensitive to chemical compositions of the surface.93-94 Typically, surfaces decorated with N,

N-Dimethyl-n-octadecyl-3-aminopropyltrimethoxysilyl chloride (DMOAP) cause

LC to orient homeotropically,93-94 while surfaces decorated with proteins orient LC planarly.90 These unique phenomena form working principles of most LC-based assays pioneered by Abbott and co-workers for detecting metal ions, small organic

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molecules and biomolecules.79-83, 85, 192-194 More recently, some LC-based protease assays were demonstrated by Bi et al., who used LC supported on oligopeptide- functionalized surfaces to detect trypsin.89 In that assay, oligopeptide- functionalized glass surfaces were incubated in protease solutions to cleave oligopeptides, and LC was used to quantify protease activities. However, two main challenges in the LC-based protease assays were noted. First, highly stable immobilized peptides that can withstand harsh washing conditions is still lacking.

Thus, washing conditions need to be controlled carefully and retain immobilized peptides. Secondly, cleavage of the immobilized peptides may result in limited changes to surface composition. Therefore, LC may not detect protease activities properly.

To address the issue on the stability of immobilized peptides, we first explore the use of dual cysteine peptides to improve the stability of immobilized peptides. In the literature, it has been shown that cysteine-labeled peptides can be specifically immobilized on surfaces used in LC-based assays,201 and thiol groups in cysteine residues are capable of forming disulfide bonds. In this work, we investigate whether this functionality of thiols can be employed to increase stability of immobilized peptides. Next, to induce significant changes in surface properties after peptide cleavage, peptides were labeled with biotin, which can be used to disrupted LC orientations on these surfaces.

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4.2 Experimental Section

4.2.1 Materials

Glass slides were purchased from Fisher Scientific (U.S.A.). N, N-Dimethyl-n- octadecyl-3-aminopropyltrimethoxysilyl chloride (DMOAP), polyoxyethylene

(20) sorbitan monolaurate (Tween 20), sodium carbonate (≥99%), 1,4-dithiothreitol

(DTT, ≥99%), streptavidin from Streptomyces avidinii (≥13 units/mg protein), trypsin (≥10,000 BAEE units/mg protein) and -chymotrypsin (≥40 units/mg protein) from bovine pancreas were purchased from Sigma-Aldrich (Singapore).

Streptavidin labeled with Cy3 (streptavidin-Cy3) in a solution (~34 μM) was purchased from Thermo Fisher (Singapore). Calcium chloride dihydrate (≥99%) was purchase from Alfa Aesar (Singapore). Phosphate buffered saline (PBS, pH

7.4) and Tris buffer (1.0 M, pH 8.0) were purchased from First BASE (Singapore).

4-cyano-4’-n-pentyl-biphenyl (5CB) was purchased from Merck (Singapore).

Peptides P1 (N-CGGGGSDAQAKSAGGRGANDEGHPEDEGGK-Biotin-C) and

P2 (N-CCGGGSDAQAKSAGGRGANDEGHPEDEGGK-Biotin-C) with ≥95% purity were synthesized by Genic Bio (China). Peptide P3 (N-

CCGGGSDAQAWSAGGFGANDEGHPEDEGGK-Biotin-C) with ≥95% purity was synthesized by GL Biochem (China).

4.2.2 Surface Modifications

Glass slides were incubated in a 5% (v/v) Decon 90 solution for 16 h to clean the surfaces. After this, the glass slides were sonicated thrice for 15 min in deionized

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(DI) water. Subsequently, the glass slides were immersed in a solution containing

0.1% (v/v) of DMOAP for 5min and then rinsed with a copious amount of DI water.

Next, the DMOAP-coated glass slides were dried with compressed nitrogen gas, followed by heating in an oven at 100 C for 2 h. Finally, DMOAP-coated glass slides were cooled at room temperature for 1 h and used immediately.

4.2.3 Immobilization of Peptides

Peptides were immobilized following a protocol published earlier.201 Briefly, a

DMOAP-coated glass slide was first exposed to UV (Spectroline® 11SC-2 UV pencil lamp, 4.5 mW/cm2 of 254 nm) for 50 s. Afterward, droplets (0.5 µl each) of carbonate buffer (100 mM, pH 10.0) containing 10-500 µM of peptides and 0.001%

(v/v) of Tween 20 were spotted on the glass slide. After incubation in a humidified chamber for 3 h, the slide was cleaned by dipping into a Tween 20 solution [0.05%

(v/v)]. Finally, the slide was rinsed with DI water and dried with compressed nitrogen gas. To reduce disulfide bonds of the immobilized peptides, the surfaces with immobilized peptides were incubated in Tris buffer (50 mM, pH 8.0) containing 100 mM of DTT for 3 h at room temperature. Subsequently, the surfaces were rinsed with DI water and dried with compressed nitrogen gas. To produce fluorescence signal for detection, solutions containing 0.34 µM of streptavidin-Cy3 were spotted onto the surface. After 1 h of incubation in a humidified chamber, the surfaces were dipped into a Tween 20 solution [0.05% (v/v)] for cleaning. Finally, the surfaces were rinsed with DI water and blown dry with compressed nitrogen gas. Fluorescence signal was detected by using a microarray scanner (Roche

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NimbleGen MS200, Switzerland) with 2% gain, 100% laser intensity and 40

μm/pixel resolution. Fluorescence intensities were quantified by using software

GenePix Pro 6.0. Each data point was an average from fluorescence intensities of

5 spots.

4.2.4 Fluorescence Protease Assay

Peptide-functionalized surfaces were incubated in Tris buffer (50 mM, pH 8.0) containing 0.001% (v/v) of Tween 20 and 250 ng/ml of trypsin at 37 C. In all experiments, 20 mM of calcium chloride was also added to improve the stability of trypsin. After 3 h of incubation, the surfaces were rinsed with DI water and dried with compressed nitrogen gas. To produce fluorescence signal, solutions containing

0.34 μM of streptavidin-Cy3 were dispensed onto each spot and incubated for 30 min. Finally, the fluorescence intensity of each spot was determined according to the same procedure mentioned above.

4.2.5 LC-Based Protease Assay

Peptides were immobilized following the procedure described above with some modifications. First, the incubation time for peptide immobilization was increased to 16 h. Next, a peptide-functionalized surface was incubated in a Tris buffer (50 mM, pH 8.0) containing 0.001% (v/v) of Tween 20 and 250 ng/ml of trypsin for 6 h. After this, the surface was rinsed with DI water and dried with compressed nitrogen gas. Subsequently, PBS buffer containing 0.001% (v/v) of Tween 20 and

0.1 pM of streptavidin was pipetted onto the surface to cover each spot of the

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immobilized peptide fully. After 16 h of incubation in a humidified chamber, the surface was blown with compressed nitrogen gas to remove the buffer. Next, an LC cell consisting of a piece of peptide-functionalized glass and a piece of DMOAP- coated glass was fabricated. A piece of 50 μm spacer (3M, U.S.A.) was used to create an optical cell from the glass slides mentioned above. Finally, 30 μl of 5CB was added to the LC cell, and the cell was observed under a cross-polarized microscope (Nikon ECLIPSE LV100POL, Japan).

4.3 Results and Discussion

4.3.1 Effect of Dual Cysteine on Peptide Surface Density

An important requirement for a protease assay is that immobilized peptides must be highly stable even under rigorous rinsing conditions. The surface density of immobilized peptide also needs to be high enough to allow sensitive detection of protease activities. To achieve stable immobilization of peptides on a surface, two cysteine-labeled peptides P1 and P2 were tested. P1 is a peptide containing a single cysteine. This peptide was chosen because thiol moiety on the cysteine residue provides an anchoring point for peptide immobilization on a UV-treated surface as demonstrated in our earlier work.201 For P2, it has the same peptide sequence as P1, except that one glycine residue was replaced with a cysteine residue near the N- terminus (Table 4.1). We propose that the second cysteine is able to increase the stability of the immobilized peptide through the formation of disulfide linkages with other peptides. Both peptides were immobilized on a UV-modified surface, and fluorescence images (Figure 4.1a) were obtained to quantify the surface density

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of immobilized peptides as shown in Figure 4.1b. For P1, fluorescence intensities increased from 332±30 to 5196±330 when the peptide concentration was increased from 10 μM to 500 μM. This result shows that P1 was successfully immobilized on the UV-modified surface, and the surface density of peptide was concentration- dependent. The surface was saturated with P1 when 500 μM of P1 was used. For

P2, fluorescence intensities increased from 3920±585 to 11763±1017 as the peptide concentration was increased from 10 μM to 500 μM. For 0 μM of P1 and P2, average fluorescence intensities were 159±32 and 159±58, respectively. This showed that there was minimal non-specific binding of streptavidin-Cy3 on unfunctionalized surfaces. Apparently, the fluorescence intensity of P2 was always higher than that of P1 at the same concentrations. For example, when both peptide concentrations were 100 μM, the fluorescence intensity of P2 was 4.5 times higher than that of P1. This result suggests that the presence of the dual cysteine label led to a higher density of immobilized peptides on the surface. Since the only difference between P1 and P2 is the additional cysteine residue in P2, the higher density was probably caused by the second cysteine residue which formed inter-peptide disulfide bonds.202-204 As a result, more peptides could be immobilized on the surface.

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Table 4.1. Peptide Sequences Peptide Peptide Sequence (N- to C-)

P1 CGGGGSDAQAK-|-SAGGR-|-GANDEGHPEDEGGK-Biotin

P2 CCGGGSDAQAK-|-SAGGR-|-GANDEGHPEDEGGK-Biotin

P3 CCGGGSDAQAW-||-SAGGF-||-GANDEGHPEDEGGK-Biotin

P1 and P2 are specific substrates for trypsin, while P3 is a specific substrate of chymotrypsin. | indicates cleavage site by trypsin || indicates cleavage site by chymotrypsin

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To test if disulfide bonds exist, DTT, a strong reducing agent, was added to the immobilization buffer to reduce the disulfide bonds.205 Figure 4.1b shows the fluorescence intensities of different peptide-functionalized surfaces with and without the addition of DTT. For P1, the fluorescence intensity did not show much difference after the addition of DTT. For P2, however, the addition of DTT led to a decrease of fluorescence intensities by 39%, 48%, 50% and 55% when the P2 concentrations were 500, 250, 100 and 10 μM, respectively. This result suggests that P2 cross-linked through disulfide bonds with other peptides and formed a 2D network on the surface (Scheme 4.1). The formation of disulfide linkages led to an increase in the surface density of peptide. When the disulfide bonds were reduced by DTT, some cross-linked peptides were released and fluorescence intensities decreased. The formation of a cross-linked peptide network is expected to improve the mechanical stability of the immobilized peptides.

Scheme 4.1. Surface Immobilization of Dual Cysteine-Labeled Peptides Through Formation of Inter-Peptide Disulfide Bonds

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4.3.2 Fluorescence Protease Assays

To develop protease assays on the P1- and P2-functionalized surfaces mentioned above, trypsin was chosen as a model protease and fluorescence intensity was used as a tool to assess the cleavage of P1 and P2. Since P1 and P2 contain lysine and arginine residues (Table 4.1), trypsin is expected to cleave the immobilized P1 and

P2 peptides and remove terminal peptide fragments from the surface (Scheme 4.2).

Because each terminal fragment contains a biotin moiety which binds to fluorescent streptavidin-Cy3, cleavage of the peptide will lead to a decrease in fluorescence intensities. Figure 4.2a shows fluorescence intensities of P1-functionalized surfaces after incubation in trypsin solutions. In the absence of trypsin, fluorescence intensities were 5114±345, 4435±94, 2388±245, and 559±35 on the P1-modified surface with decreasing P1 densities (500, 250, 100, and 10 μM of P1). For comparison, when 250 ng/ml of trypsin was used, fluorescence intensities of these spots decreased to 1723±84, 1359±189, 544±27, and 239±52, respectively. The decrease in fluorescence intensities strongly suggests that the immobilized P1 peptides were cleaved by trypsin.

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Scheme 4.2. Detection Principles of the LC-based Protease Assays

(a) Mechanism of streptavidin-Cy3 binding to peptide-functionalized surfaces, where the presence of streptavidin can be detected by using fluorescence or LCs. (b) Cleavage of immobilized peptides by proteases leading to the removal of biotin from the surface, where the absence of biotin prevents binding of streptavidin-Cy3 to the surface.

Since the use of dual cysteine labels in P2 was shown to increase surface density of immobilized peptides, protease assays were also developed on the P2- functionalized surfaces and their performance was compared. Figure 4.2b shows fluorescence intensities of the P2-functionalized surfaces after incubation in trypsin solutions. In the absence of trypsin, fluorescence intensities were 11262±211,

10500±143, 8749±595, and 4729±236 on the surfaces functionalized using 500,

250, 100, and 10 μM of P2. For comparison, when 250 ng/ml of trypsin was used, fluorescence intensities of these spots decreased to 4606±97, 3410±114, 2555±205, and 1173±183. Similarly, the decrease in fluorescence intensities in the presence of trypsin strongly suggests that P2-immobilized surfaces were cleaved by trypsin.

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For both P1- and P2-functionalized surfaces, a significant decrease in fluorescence intensities was observed when trypsin concentration increased from 0.5 to 10 ng/ml.

By comparing Figure 4.2a and 4.2b, a decrease in fluorescence intensities on the

P1-functionalized surfaces ranged from 297 to 3119, whereas on the P2- functionalized surfaces, it ranged from 3556 to 7090. The decrease in fluorescence intensity is more pronounced on the P2-functionalized surfaces because the surface density of P2 was higher and the signal-to-noise ratio was better. Thus, the P2- functionalized surface with dual cysteine labels in P2 is a better candidate for the development of LC-based protease assays.

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4.3.3 Detection of Immobilized Peptides Using LC

To develop LC-based protease assays, we first investigated the ability of LC to detect immobilized P2 on UV-modified DMOAP coated surfaces. Figure 4.3a shows that LC supported on the P2-functionalized surface was not disrupted by P2, as the optical appearance of LC was uniformly dark. This is probably due to the small size of P2 which was unable to disrupt the LC. To increase the size of P2, the

P2-functionalized surface was incubated in solutions containing streptavidin with a concentration range between 0.001 and 0.1 pM. This range was chosen based on

206 the dissociation constant of biotin-streptavidin complex (Kd ~ 0.01 pM). The biotin label on the P2 allowed binding of streptavidin to P2 and increased its size to disrupt LC. Figure 4.3b-d showed that 0.01 pM (1 Kd) of streptavidin together with 100 M of P2 was the minimum concentration required to disrupt LC and produced bright signals. These results clearly demonstrated the importance of introducing streptavidin to disrupt LC orientation. Therefore, protease assay can be carried based on the ability of LC to detect full-length P2 immobilized on the surface. Cleavage of P2 results in the loss of biotin label and the ability of P2 to disrupt LC.

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Figure 4.3. Detection of immobilized P2 (conjugated to streptavidin) on a solid surface by using LC images under crossed polars. Streptavidin concentrations of (a) 0, (b) 0.001, (c) 0.01 and (d) 0.1 pM were used. Surface density of P2 was controlled by using different concentrations of P2 (10, 100, and 500 μM from left to right) during peptide immobilization. Scale bar represents 1mm.

4.3.4 LC-Based Protease Assays

Based on the detection principle mentioned above, LC-based protease assays were developed. In the first experiment, P2-functionalized surfaces were incubated in solutions containing 250 ng/ml of trypsin for 6 h, allowing the cleavage of P2 peptide and removal of the biotin label. Subsequently, the surfaces were incubated in another solution containing 0.1 pM of streptavidin. Figure 4.4a shows that in the presence of 250 ng/ml of trypsin, dark LC spots were observed. This is because cleavage of the immobilized P2 by trypsin removed the biotin labels, preventing the binding of streptavidin to the surface. Therefore, LC remained dark as its orientation was not disrupted. In contrast, if the surface was incubated in 500 ng/ml

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of chymotrypsin (which did not cleave P2 based on its specificity), bright LC spots were observed (Figure 4.4b). This is because immobilized P2 was not cleaved by chymotrypsin. Therefore, binding of streptavidin to the surface disrupted orientations of LC and resulted in bright LC spots. This result strongly suggests that this protease assay is a functional protease assay based on the specific cleavage of peptide.

To study the generality of the protease assay, we introduced P3 as a specific substrate for chymotrypsin (Table 4.1). Compared to P2, P3 has an identical amino acid sequence, expect for the substitution of lysine and arginine residues with

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tryptophan and phenylalanine, respectively. Following similar procedures described above, the P3-functionalized surfaces were incubated in solutions containing 250 ng/ml of chymotrypsin. Figure 4.4c shows that in the presence of

250 ng/ml of chymotrypsin, dark LC spots were observed. In contrast, if the surface was incubated in 500 ng/ml of trypsin (which did not cleave P3 based on its specificity), bright LC spots were observed (Figure 4.4d). These results demonstrated that specificity of the LC-based protease assay can be easily achieved by using custom-designed peptides that are specifically cleaved by target proteases.

Therefore, such LC-based protease assays can be generally applied to different protease targets.

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4.3.5 Limit of Detection (LOD) of LC-Based Protease Assays

To determine the LOD of LC-based protease assays, we decreased the concentrations of proteases and tested the ability of LC to detect whether the immobilized peptides were cleaved. For the P2-functionalized surfaces, we observed that LC remained dark when the trypsin concentrations were higher than

0.1 ng/ml (Figure 4.5a-d). In contrast, when the trypsin concentration was 0.01 ng/ml, bright LC spots were observed (Figure 4.5e). This implies that the minimal

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trypsin concentration required to cleave P2 and generate observable signals was 0.1 ng/ml (~4 pM) trypsin. Next, for the P3-functionalized surfaces, it was observed that LC remained dark when the chymotrypsin concentrations were higher than 1 ng/ml (Figure 4.6a-c). When 0.1 ng/ml of chymotrypsin was used, bright LC spots were observed (Figure 4.6d). This implies that the minimal concentration of chymotrypsin that can be detected is 1 ng/ml (~40 pM). By comparison with other

LC-based protease assays (Table 4.2), this work boasted better LOD and required shorter protease incubation period (6 h) than earlier work by Bi et al., where they reported a LOD of ~2 nM for trypsin with an incubation period of 8.4 h.89 Another work by Chen and Yang also reported a poorer LOD of ~200 pM for trypsin and chymotrypsin with a longer incubation period of 20 h.109 However, a more recent work by Chen and Yang showed comparable LOD of ~40 pM and ~4 pM for trypsin and chymotrypsin, respectively.104 However, this assay required the use of multiple lysine residues to enhance LC responses. Because lysine can be cleaved by trypsin or other proteases, the specificity of the assay was questionable. In this aspect, we are able to retain the specificity of the assays by using streptavidin to enhance LC response towards immobilized peptides.

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Table 4. 2. Comparison of Trypsin and Chymotrypsin Assays Recently Developed Reference Detection Principle Limit of Detection

Zauner et al. 133 Fluorescence Trypsin (2 pM)

He and Ma 70 Fluorescence Trypsin (4 pM)

Cao et al. 34 Electrochemical Trypsin (600 pM)

Fan et al. 31 Fluorescence Trypsin (11 pM)

Chymotrypsin (6 pM)

Bi et al. 89 LC-based Trypsin (2 nM)

Chen and Yang 109 LC-based Trypsin (200 pM)

Chymotrypsin (200 pM)

Chen and Yang 104 LC-based Trypsin (40 pM)

Chymotrypsin (4 pM)

This work LC-based Trypsin (4 pM)

Chymotrypsin (40 pM)

4.4 Conclusion

In this work, a heterogeneous protease assay based on dual cysteine-labeled peptides (as substrates) and LC (as optical readout) was successfully developed.

This protease assay relies on the ability of LC to distinguish cleaved and un-cleaved immobilized peptide on solid surfaces, and it can be used to detect trypsin and

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chymotrypsin with good specificity. LODs of this assay (0.1 ng/ml for trypsin and

1 ng/ml for chymotrypsin) are better or comparable with literature values (Table

4.2). Moreover, our LC-based assay requires simpler instrumentation than other fluorescence or electrochemical methods.31, 34, 70, 133 Peptide substrates can also be custom-made to tailor specific proteases. Therefore, the LC-based assay shows promises as a simple yet powerful analytical tool for detecting a wide range of proteases.

CHAPTER 5

CHAPTER 5 DISTORTION OF LIQUID CRYSTAL PHASE BY

MICROPARTICLES FUNCTIONALIZED WITH

OCTADECYL-TERMINATED OLIGOPEPTIDES

Based on work in Chapters 3 and 4, a generic approach was developed for LC- based protease assay. However, these assays do not provide quantitative assay results. To address this limitation, in this chapter, we explore the introduction of oligopeptide-functionalized microparticles within a bulk LC phase. Additionally, oligopeptides were labeled with octadecyl (C18) groups to provide interactions with

LC that are analogous to C18 groups of DMOAP. We studied how LC director profiles are distorted by these microparticles within a bulk LC phase. Such distortions can be measured, and surface density of oligopeptides can be potentially predicted using a facile computational model. This provides a quantitative analysis of our system based on a straightforward experimental setup.

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5.1 Introduction

Nematic liquid crystals (LCs) are rod-like molecules that possess both fluidity of conventional fluids and crystalline-like ordering of solid crystals. Such ordering of

LC molecules results in an average direction (director) in which the long axes of

LC molecules align. In order to control the LC director profile, external electromagnetic fields or chemical modifications to surfaces confining the LC phase can be used. For example, surfaces can be chemically modified with N,N- dimethyl-n-octadecyl-3-aminopropyltrimethoxysilyl chloride (DMOAP) to preferentially anchor LC homeotropic (perpendicular) to the surface.93 In contrary, uncoated polystyrene surfaces preferentially anchor LC planarly (parallel) to the surface.117 Interestingly, preferential anchoring of LC at surfaces is able to influence orientations of LC several micrometers away in the bulk due to long- range interactions of the LC phase. Several studies in the past showed that anchoring energy of LC on chemically functionalized surfaces can be expressed as:93-94, 207-209

∅ (5.1) where W is the anchoring strength,  is the preferential anchoring angle (easy axis), and  is the actual anchoring angle of LC at the surface.210 Due to the global minimization of free energy, the actual anchoring angle at surfaces may deviate substantially from the preferential anchoring angle, especially for surfaces with weak anchoring strengths. Therefore, to study the effects of anchoring strengths on the director profile, LC cells are commonly used. In an LC cell, two glass surfaces are used to confine an LC phase with a fixed thickness (~1 - 100 μm). Additionally,

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glass surfaces can be chemically modified to preferentially anchor LC homeotropically, and this produces LCs that are uniformly oriented homeotropically. In other cases, one of the glass surfaces can also be chemically modified to preferentially anchor LC planarly, and this results in a hybrid aligned nematic (HAN) cell. The director profile of an HAN cell can be solved by using the following modified equations proposed by Babero and Bartolino (and later by Price and Schwartz),210-211 where preferential anchoring at both surfaces are considered:

2 2∅ sin sin 2∅ sin2∅ 0 (5.2)

∅ sin sin 2∅ 0 (5.3) where 1 and 1 represents the actual and preferential LC anchoring angle at surface

1 whereas 2 and 2 represent the actual and preferential LC orientation at surface

2. Preferential anchoring angle of LC at surfaces are 0 and /2 for homeotropic and planar anchoring, respectively. d represents cell thickness. L1 and L2 are the extrapolation lengths of surfaces 1 and 2, respectively. L1 and L2 are defined as

L1=K/W1 and L2=K/W2. K is the one constant approximation of Frank elastic

-12 constant with a typical value of 9 × 10 N, while W1 and W2 are the anchoring strengths of surfaces 1 and 2, respectively. Typical values for anchoring strength of surfaces with preferential homeotropic or planar anchoring are 10-4 and 10-6 J/m2, respectively. For a given HAN cell with known d, 1, 2, W1, and W2, 2 can be solved by using Eq. 5.2, and 1 can be solved from Eq. 5.3. The resulting director profile can be assumed to vary linearly from one surface to another,211 and this causes the HAN cell to display birefringence. Consequently, such a HAN cell

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produces observable LC optical signals under crossed polars. Application of HAN cells for sensing purposes was rigorously studied by Abbott and co-workers, where they demonstrated the influence of biomolecules-decorated surfaces on the preferential anchoring angles of LC.95, 98, 103, 212 Such changes in preferential anchoring angles of LC were easily observed through LC optical signals, and these studies demonstrated the potential of using LC as an optical sensing platform for biomolecules. Later on, Yang and coworkers also made use of such principles to develop LC-based immunoassays.90, 213 For example, Xue and Yang used LC cells that oriented LC homeotropically, with one glass surface coated with DMOAP, while another surface was functionalized with antibodies.90 The binding of specific antigen caused preferential anchoring of LC to change from homeotropic to planar, and this produced bright LC optical signals.

Since LC anchor with preferential angles relative to contacting surfaces, the introduction of spherical microparticles into an LC phase provides an interesting platform to study how microparticles influence the LC director profile.214-216 For example, Poulin and Weitz used water-in-LC emulsions and water-in-LC-in-water double emulsions to study the possible LC director profiles around surfaces of emulsion droplets with preferential homeotropic or planar anchoring.216 Due to the geometry of spherical microparticles, non-uniform director profiles are observed.

For example, for homeotropic anchoring of LC at surfaces of microparticles, radial director profiles are formed around the microparticles. Due to such non-uniformity, submerged microparticles in LC phase can be easily observed under crossed polars.

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Recent research on the microparticles submerged in an LC phase has focused on the use of LC as a soft template for assembly of microparticles,112-120 and trapping of microparticles at specially fabricated surfaces.87, 121-123 However, there is no report of sensing applications using microparticles submerged in LC. In order to develop a sensitive and selective sensing platform, oligopeptides are ideal choices.

Oligopeptides can selectively bind to target analytes, and they are natural substrates for proteases.217 Therefore, more work is required to explore the interactions of oligopeptide-functionalized microparticles submerged in LC, and how protease sensing can be achieved.

In this work, we make use of an LC cell and introduced oligopeptide- functionalized, spherical microparticles in the LC phase. This setup provides a stable director profile for studying the influence of oligopeptide-functionalized microparticles on the director profile. We also explore the use of octadecyl- (C18)- terminated oligopeptides since these C18 groups resemble the long alkyl chains found in DMOAP. Therefore, we investigate how these oligopeptide-functionalized microparticles interact with LC, and whether such microparticles can be used to develop LC-based protease assays.

5.2 Experimental Section

5.2.1 Materials

Glass slides were purchased from Fisher Scientific (U.S.A.). 5% (w/v) silica microparticles (7.38±0.24 μm) were purchased from Microparticles GmbH

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(Germany). N, N-dimethyl-n-octadecyl-3-aminopropyltrimethoxysilyl chloride

(DMOAP), poly(ethylenimine) (PEI) (Mw ~ 750,000 Da, 50% (w/v) in water), polyoxyethylene (20) sobitan monolaurate (Tween 20), sodium carbonate

(≥99.5%), and trypsin from bovine pancreas (≥10,000 BAEE units) were purchased from Sigma-Aldrich (Singapore). Phosphate buffered saline (PBS) and sodium dodecyl sulfate (SDS) were purchased from First BASE (Singapore). Calcium chloride dihydrate (≥99%) and 4-cyano-4’-n-pentyl-biphenyl (5CB) was purchased from Merck (Singapore). Oligopeptides with purity ≥95% (P1: N-Cys-(Lys)4-C and

P1-C18: N-Cys-(Lys)4-C18-C) were synthesized by GL Biochem (China).

5.2.2 Surface Modifications

Glass slides were immersed in a 5% (v/v) Decon 90 solution overnight to clean the surfaces. After this, the glass slides were sonicated thrice for 15 min in deionized

(DI) water. Subsequently, the glass slides were immersed in a DMOAP solution

[0.1% (v/v)] for 5 min, and then rinsed with a copious amount of DI water. Finally, the DMOAP-coated glass slides were dried under a stream of nitrogen, followed by heating in an oven at 100 C for 1 h.

5.2.3 Immobilization of Oligopeptides on PEI-coated Microparticles

Microparticles [0.5% (w/v)] were immersed in an ethanol solution containing 1%

(w/v) of PEI for 3 h. After this, the solution was centrifuged briefly to collect the

PEI-coated microparticles. Subsequently, DI water was added to wash the PEI- coated microparticles three times. To introduce aldehyde groups, the PEI-coated

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microparticles were incubated in a glutaraldehyde solution [15% (w/v)] overnight.

Next, the solution was centrifuged briefly to collect the glutaraldehyde-coated microparticles, and then DI water was added to wash the microparticles three times.

To immobilize oligopeptide, the microparticles were incubated in carbonate buffer

(100 mM, pH 10.0) containing 200 μM of oligopeptide. After immobilization, the solution was centrifuged briefly to collect the oligopeptide-functionalized microparticles, and then DI water was added to wash the oligopeptide- functionalized microparticles three times. Finally, the microparticles were dried in a desiccator (under vacuum) for 5 h.

5.2.4 Fabrication of LC Cells

An LC cell was fabricated by using two DMOAP-coated glass slides and two strips of 100 μm spacer (3M, U.S.A.). After this, 30 μl of 5CB containing oligopeptide- functionalized microparticles were introduced into the cavity between the two glass slides. Optical appearance of the LC cell was observed using an optical microscope

(Nikon ECLIPSE LV100POL, Japan). Distortion radius of LC textures were measured, and the average and standard deviation of 40 readings were obtained using Image J software.

5.2.5 Protease Assay

Oligopeptide-functionalized microparticles were incubated in Tris buffer (50 mM, pH 8.0) containing 1 μg/ml of trypsin. 20 mM of calcium chloride was added as a protease stabilizer. The solution was incubated at 37 C for 3 h. After this, the

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solution was centrifuged to collect the microparticles, and then DI water was added to wash the microparticles three times to remove remaining trypsin. Finally, the microparticles were dried in a desiccator (under vacuum) for 5 h. After drying, LC was mixed with microparticles to a final concentration of 0.5% (w/v).

5.3 Results and Discussion

5.3.1 Interactions of LC with Submerged Microparticles

To understand how functionalized microparticles interact with LC, microparticles at different stages of functionalization were mixed with LC and the optical textures of LC were obtained. Figure 5.1a,b shows that when unfunctionalized microparticles were present, no observable distortions were observed in the bright field, while microparticles appeared bright with continuous optical texture under crossed polars. No pronounced LC distortions were observed in regions around the microparticles. Next, when microparticles functionalized with oligopeptide P1 were used, similar bright and crossed polar images were observed (Figure 5.1c,d).

These results show that unfunctionalized and P1-functionalized microparticles caused planar anchoring of LC at the surfaces. Based on previous reports for microparticles with planar anchoring, distortions to LC director profiles are typically limited to regions close to the surfaces of microparticles.216 Therefore, for unfunctionalized and P1-functionalized microparticles, only LC close to surfaces of microparticles were influenced, while LC away from these microparticles were not distorted. To produce pronounced LC distortions, microparticles were functionalized by using oligopeptide P1 with a terminal C18 group. As shown in

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Figure 5.1e,f, distortions of LC around the microparticles were observed under both bright and dark fields. From Figure 5.1e, the distortion around the microparticles was demarcated by a ring of distortion. This was also shown in images under crossed polars in Figure 5.1f. Additionally, a distinctive “cross” were observed, which corresponds to the regions where LC orientations are parallel to the polarizer and analyzer.91 This indicated that the director profile around the microparticle was radially orientated. This strongly suggested that the presence of C18 caused LC to be anchored homeotropically at the surfaces of microparticles. As a result, LC away from the microparticles was oriented radially under the influenced of these C18- terminated oligopeptides.

For microparticles with homeotropic anchoring, two types of defect structures are possible, namely, the hyperbolic hedgehog defects and Saturn rings.216 The observations in Figure 5.1e,f resembles that of Saturn rings formed around microparticles. Previous works by Gu and Abbott and Musevic et al. showed that with increased confinement of microparticles with homeotropic anchoring within an LC cell, Saturn rings were observed.218-219 In those previous experiments, LC cells were fabricated with boundary surfaces that anchored LC uniformly and planarly to the surfaces. As such, Saturn rings were formed perpendicular to the confining glass surfaces. Furthermore, for LC cells smaller than a critical thickness

(3.5 ± 0.1 μm), Saturn ring structures were observed in place of hyperbolic hedgehogs defects.219 These observations can be attributed to the effects of confinement, where influences by the boundary glass surfaces impose a significant

CHAPTER 5

Frank free energy penalty for deformation of LC away from the overall LC director.

This promoted the formation of Saturn rings. On the contrary, when confinement of microparticles was relaxed, LC director field oriented around the microparticles to form hyperbolic hedgehog defects. In our case, we observed Saturn rings that were parallel to the confining glass surfaces. One major difference is that surface anchoring of confining surfaces was uniformly planar in the earlier studies, while our study made use of confining surfaces with strong homeotropic anchoring.

Typically, homeotropic surface anchoring strengths are two orders of magnitude larger than planar surface anchoring strengths.210 Therefore, the strong homeotropic anchoring strength has likely contributed to the strong confinement effects of LC director field around the microparticles. This resulted in our observations of the

Saturn rings.

Based on these results, we proposed the LC director profile around the microparticles with preferential planar and homeotropic anchoring (Scheme 5.1).

Importantly, the presence of microparticles with preferential homeotropic anchoring causes pronounced LC distortions that are observable under bright field and crossed polars (Figure 5.1e,f).

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Scheme 5.1. Possible LC Director Profiles Surrounding Microparticles Submerged in LC

Microparticles are confined by two glass surfaces that preferentially anchor LC homeotropically. (a) and (b) show the LC director profile for microparticles with preferential planar and homeotropic anchoring, respectively.

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5.3.2 Effect of Surface Density of Oligopeptide on LC Distortion

Since the presence of C18-terminated oligopeptides preferentially anchors LC homeotropically, we further investigated the critical concentration of C18- terminated P1 required to induce homeotropic anchoring on the surfaces of microparticles. We also investigated the effect of surface density of C18-terminated

P1 on LC distortion. To quantify the distortion effects, we introduced the distortion radius (see insert in Figure 5.2). Figure 5.2 shows that at 20 μM of C18-terminated

P1, no distortion ring was formed. This implies that preferential anchoring of LC was planar, and the surface density of C18-terminated P1 was not sufficient to anchor LC homeotropically. Next, we observed that 100 μM was the critical C18- terminated P1 concentration required before distortion rings were observed around the microparticles, and distortion radius was 5.6 ± 0.5 μm. At this critical concentration, LC was preferentially anchored homeotropically, and this resulted in the radial orientation of LC near the microparticles and the formation of LC distortion ring. The increase of C18-terminated P1 concentration from 100 to 200

μM caused distortion radius to increase from 5.6 ± 0.5 to 8.4 ± 1.2 μm. Further increase of C18-terminated P1 concentration from 200 to 400 μM resulted in distortion radius of 8.1 ± 1.2 μm, suggesting that surface density of C18-terminated

P1 did not increase as concentration increased from 200 to 400 μM. These results also suggest that increase of C18-terminated P1 surface density led to increased homeotropic surface anchoring strength for surfaces of microparticles. Therefore,

LC was influenced, to a greater extent, by the increasing homeotropic surface anchoring strength with increasing surface density of C18-terminated P1. This led

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to the greater distortion of LC in the bulk phase by microparticles with a higher surface density of C18-terminated P1.

5.3.3 Effect of Cell Thickness on LC Distortion Diameter

So far, we showed the importance of anchoring strengths and preferential anchoring angle of LC at surfaces. Additionally, cell thickness can also affect the interaction of LC with the surfaces. Based on work by Price and Schwartz, they showed that LC director profile was more gradual for thicker cells.210 This was due to lower elastic energy caused by smaller gradients in the LC director, and this resulted in the weaker coupling of surfaces in the HAN cell. Therefore, we investigated the effect of cell thickness on the resultant LC distortion diameter around microparticles. In this experiment, we kept C18-terminated P1 concentration

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constant at 200 μM and varied cell thickness. From Figure 5.3, distortion radius decreased slightly from 10.7 ± 1.4 to 10.6 ± 1.6 μm as LC cell thickness decreased from 200 to 150 μm. Further decrease in cell thickness from 150 to 24 μm caused a gradual decrease in distortion radius to 7.7 ± 0.7 μm. The most significant decrease was observed when cell thickness decreased from 24 to 12 μm, where distortion radius decreased to 5.6 ± 0.7 μm. These results suggest that decrease in cell thickness reduced distortion of LC near the microparticles. From the perspective of energy minimization, this trend was likely due to the increase in elastic energy penalty for the formation of distortion rings. For a thinner cell, elastic energy penalty is larger for distortion in the LC director.210 Therefore, smaller distortion rings were observed in thinner cells.

Figure 5.3. Effect of LC cell thickness on distortion radius near submerged microparticles.

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5.3.4 Computational Model for LC Director Profile

In order to compute the orientation of LC molecules at surfaces of microparticles and the confining DMOAP surfaces, we made use of Eq. 5.2 and 5.3 previously developed.210-211 At discrete positions along the z-axis, we computed the director profile along the x-axis (in Scheme 5.1) based on an imaginary HAN cell formed between the surface of a microparticle (surface 2) and an imaginary surface (surface

1). This imaginary surface was created to incorporate the effect on LC director profile caused by the pair of DMOAP-coated glass surfaces used to fabricate the actual LC cell. In this imaginary HAN cell, LC preferentially anchored homeotropically with respect to the curvature of the surface of the microparticle, while LC preferentially anchored planarly at the imaginary surface. The planar anchoring of LC at the imaginary surface was to represent the director profile caused by the two DMOAP-coated glass surfaces, which was parallel to the z-axis.

The diameter of microparticles was 7.38 μm, while cell thickness, d, was set as 20

μm. This imaginary cell thickness value was chosen based on our experimental results, as maximum distortion radius measured was <12 μm (Figure 5.2 and 5.3).

After obtaining the orientation of LC at the two surfaces, we assumed a linear transition of LC orientation from one surface to the other.210 On the basis of assumptions mentioned above, we computed the LC director profile between the microparticle and the imaginary surface. From the computed director profile, we can identify the distortion ring started from a region where the director of LC exceeded a critical angle. This critical angle can be obtained from our experimental results. Additionally, In order to understand how parameters like the surface density

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of oligopeptides and actual LC cell thickness affect the radius of distortion rings observed, we can vary the surface anchoring strengths of the microparticle and the imaginary surface.

5.3.5 Effect of Oligopeptide Surface Density on Computed Distortion Radius

Since the presence of C18-terminated P1 causes the formation of distortion ring, we investigated the effects of surface density of the oligopeptide on the computed distortion radius. To achieve this, we varied surface anchoring strengths of the

-4 -4 2 microparticle, W2, from 4 × 10 to 1.8 × 10 J/m , which represents decreasing the

-4 2 surface density of oligopeptide functionalization. For W2 at 4 × 10 J/m (Figure

5.4a), which represents the case of saturated oligopeptide surface density, we observed that the director profile plot was similar to the radial profile proposed in

Scheme 5.1b. From this plot, we observed that LC director profile near the center of the microparticle deviated most from the z-axis. From the center of the microparticle, LC director profile rotated from homeotropic to planar orientation at the imaginary surface. On the contrary, at the top and bottom of Figure 5.4a, LC orientation was mostly planar. This was due to the curvature of the microparticle, where these regions were preferentially anchored parallel to the z-axis. Next, in order to determine the distortion radius from the computed data, we compared this computational results with our experimental observations. From Figure 5.2, the measured distortion radius was ~8 μm for surfaces saturated with C18-terminated

P1. Relating this to our computed director profile, at 8 μm away from the microparticle, the computed LC orientation that deviated most from the z-axis was

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~66 (with respect to the z-axis). With this benchmark, we propose that a distortion region started when the director of LC exceeded 66 (shaded region in Figure 5.4a).

Next, to study the effects of oligopeptide surface density on distortion radius, we reduced the surface anchoring strength from 4 × 10-5 to 1 × 10-6 J/m2. From Figure

5.4b, we observed that a minimum surface anchoring strength of ~1.8 × 10-6 J/m2 was required for the formation of distortion ring very close to the microparticle surface (~3.9 μm). Next, as surface anchoring strength of microparticle increased from 1.8 × 10-6 to 4 × 10-6 J/m2, distortion radius increased sharply from 3.9 to 7.8

μm. It further increased slightly to 8 m as the surface anchoring energy increased from 4 × 10-6 to 4 × 10-4 J/m2. This result shows a strong dependence on surface anchoring strength of a microparticle on the distortion radius. With increasing surface anchoring strength of a microparticle, LC far away from the microparticle was influenced and deviated from the z-axis, giving rise to the perceived increase in distortion radius. However, when anchoring strength was > 4 × 10-5 J/m2, the distortion radius did not increase significantly.

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5.3.6 Effect of LC Cell Thickness on Computed Distortion Radius

Next, to study the effect of actual LC cell thickness on distortion radius, we correlated the anchoring strength of the imaginary surface to the actual cell thickness. Since a smaller cell thickness increases coupling of the two DMOAP-

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coated surface and causes greater energy penalty for any distortion to the LC director profile,210 we can introduce this effect by adjusting surface anchoring strength of the imaginary surface. To represent LC cell of decreasing thickness, we can increase anchoring strength of the imaginary surface. For this study, surface anchoring strength of microparticle was kept constant at 4 × 10-4 J/m2, which represents saturated oligopeptide surface density, and this allows direct comparison with experimental results shown in Figure 5.3. To represent a thick LC cell, we

-6 2 computed the LC director profile for W1 at 1 × 10 J/m and observed a similar director profile as seen above (Figure 5.5a). Using the same criteria for distortion radius (≥66), we computed the distortion radius as surface anchoring strength of the imaginary surface increased from 1 × 10-6 to 4 × 10-4 J/m2 (Figure 5.5b). For anchoring strength of 1 × 10-6 J/m2, the computed distortion radius was 11.3 μm.

This corresponds well with the maximum distortion radius of 10.7 μm observed in

Figure 5.3. Next, as surface anchoring strength increased from 1 × 10-6 to 8 × 10-6

J/m2, distortion radius decreased from 11.3 to 8.2 μm. Subsequent increase of surface anchoring strength to 1 × 10-4 J/m2 only decreased distortion radius slightly to 7.9 μm, while a further increase of anchoring strength to 4 × 10-4 J/m2 did not affect the distortion radius. This computational result follows a similar trend observed in Figure 5.3, where decreasing cell thickness resulted in decreased distortion radius. However, an important trend from Figure 5.3 was the sharp decrease in distortion radius when cell thickness decreased from 24 to 12 μm. Such a sharp decrease in distortion radius was not present in Figure 5.5b. This was likely due to the thickness of the imaginary cell used for the computation, which was

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larger than the actual 12 μm cell thickness used in the experiment. To test this hypothesis, we reduced the cell thickness in our computation from 20 to 10 μm.

Indeed, we obtained a distortion diameter of 5.3 μm, and this corresponds well with the experimental value of 5.6 μm for a 12 μm-thick LC cell.

Figure 5.5. (a) Computational results for LC director profiles between the surface -6 2 of a microparticle and an imaginary surface (W1 = 1 × 10 J/m ) with preferential homeotropic and planar anchoring, respectively. The green-shaded region indicates the distortion region. (b) Influence of cell thickness on the computed distortion radius was introduced by varying the anchoring strength of the imaginary surface.

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Therefore, the above computational approach agrees well with the experimental trends observed. This model also provides a facile approach for the correlation of surface anchoring strengths to the observed distortion radius. Therefore, it can potentially be applied to quantify changes to oligopeptide surface densities by measurements of distortion radius.

5.3.7 Application of Oligopeptide-Functionalized Microparticles Submerged

in LC as a Trypsin Assay

To study the feasibility of using C18-terminated oligopeptides in protease assays, we incubated C18-terminated, P1-functionalized microparticles in solutions containing trypsin. From Figure 5.6a,b, preferential homeotropic anchoring of C18- terminated P1 was still observable. However, at 1 μg/ml of trypsin (Figure 5.6c), microparticles showed preferential planar anchoring (similar to the case of P1- functonalized microparticles in Figure 5.1d). This is attributed to the cleavage of

C18 groups of C18-terminated P1 in the presence of trypsin activity, and the absence of C18 caused LC to preferentially anchor planarly. Thus, this demonstrates the facile optical observations required to assess the presence of protease activities.

Importantly, the ability for C18-terminated oligopeptides to promote the formation of observable Saturn rings, and the correlation between the sizes of Saturn rings and surface anchoring strengths can potentially provide a quantitative measurement for cleavage of immobilized oligopeptides by protease activities. Such protease assays based on C18-terminated oligopeptides can potentially be combined with one-bead-one-compound (OBOC) technique to provide a high-throughput

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screening assay for protease activities.220 This potential technique is photo-stable, and does not rely on secondary enzymatic reactions for signal readout.

5.4 Conclusion

In this work, we demonstrated the use of octadecyl- (C18)-terminated oligopeptides to functionalize microparticles and influence the orientation of LC around it. This interaction can be easily controlled through manipulation of physical parameters like oligopeptide surface density and cell thickness. Interestingly, by applying a facile computational model, these parameters can be determined by measurements of distortion radius. The importance of C18 groups in influencing LC orientation is also employed in a proof-of-concept protease assay, where trypsin activity was shown to remove C18-terminated oligopeptides from microparticles. This resulted in observable changes in LC texture, and this demonstrates a facile system that can be readily combined with one-bead-one-compound (OBOC) methods for the high- throughput screening.

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CHAPTER 6 SURFACTANT-DRIVEN ASSEMBLY OF

POLY(ETHYLENIMINE)-COATED MICROPARTICLES AT

THE LIQUID CRYSTAL/WATER INTERFACE

Although Chapter 5 provided preliminary support for quantitative LC-based assays, this technique still lacks the ability for real-time protease activity monitoring. To overcome this, the introduction of C18-terminated, peptide-functionalized microparticles at LC/water interface can potentially provide a constant interaction of microparticles with analytes in the water phase and underlying LC for real-time monitoring. However, there is a lack of studies related to responsive assembly of microparticles at LC/water interfaces. Therefore, in order to provide a fundamental understanding of the assembly of microparticles at LC/water interface, we investigated a model system of poly(ethylenimine) (PEI)-coated microparticles that allow strong interactions with surfactants in the water phase. This study can provide us with a fundamental understanding of how adsorption of target analytes can be transduced as macroscopic changes in microparticles assembly patterns.

Fundamental understandings from this study will provide the foundation for future work related to real-time monitoring of protease activity based on microparticles assembly at LC/water interface.

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6.1 Introduction

The self-assembly of microparticles into uniform two- and three-dimensional structures has been exploited intensively in recent years because it has many applications in microphotonics,221 microelectromechanical systems,222 and tissue engineering.223 For these applications, there are strong motivations to influence and control the final assemblies. The surface morphologies, shapes, and chemical compositions of microparticles are all important factors in determining the final forms of assemblies. For example, Dendukuri et al. assembled amphiphilic microparticles at water and oil interfaces.224 They demonstrated the self-assembly of amphiphilic microparticles by minimizing the microparticles’ surface energy.

This resulted in the preferential location of the hydrophobic and hydrophilic portions of the microparticles in the oil and water phases, respectively. To form more distinct microstructures, Shields et al. synthesized anisotropic microparticles and formed linear and zigzagged chains under external electric and magnetic fields.225 Metal coatings resulted in highly polarizable microparticles, allowing external magnetic and electric fields to influence the final assemblies. These examples illustrate how the synthesis of microparticles with specially designed shapes and chemical compositions can influence their assembly at liquid interfaces.

Another crucial consideration for microparticle assembly is the supporting interfaces of these assemblies. Precise engineering of the supporting interfaces can provide templates for direct microparticle assembly, especially at fluid interfaces such as those of liquids or liquid crystals (LC). Because of the sizes of microparticles, the dominance of surface tension causes microparticles to be

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trapped at fluid interfaces.226 Consequently, the deformation of fluid interfaces by microparticles and the curvature of fluid interfaces can direct microparticle assembly.227-228 For example, Cavallaro et al. assembled rodlike microparticles at regions of high interfacial curvature created at water interfaces by microposts.229

They demonstrated how these microstructures and anisotropic microparticles drive microparticle assembly by long-range capillary forces. To control microparticle assemblies more actively, Chen et al. created unique microstructures with large surface areas (greater than square millimeters) at liquid interfaces using standing waves as templates.230 Interestingly, the global microstructure patterns formed were dependent on the drift energy gradients caused by the standing waves rather than on capillary forces. Such water interfaces provide soft templates with great flexibility for the creation of distinct microstructures. LC have also been utilized as soft templates for microparticle assembly. The anisotropic nature of LC allows for the creation of unique microparticle assemblies without intricate control of interfacial curvatures. Instead, microparticles self-assemble depending on the orientation of the LC molecules (or LC director field) in the LC phase. A variety of microparticle assemblies controlled by LC director fields have been reported by various research groups.92, 112-114, 231

To understand the effects of LC on microparticles, early works were carried out by

Poulin et al. and Muševič et al. using water droplets or solid microparticles

215, 219 dispersed in LC. The presence of microparticles in the bulk LC caused the formation of topological defects in the LC phase near the surfaces of the

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microparticles. Attraction among topological defects and interactions with the LC director field eventually led to different types of microparticle assemblies, as

88, 215-216, 219, 232-234 reported in the literature. Based on these understandings, studies involving microparticle assembly at LC/water and LC/air interfaces have been carried out to overcome the confinement of microparticles in the bulk LC phase.

Initial work by Koenig et al. demonstrated the assembly at the LC/water interface of silica microparticles coated with N,N-dimethyl-n-octadecyl-3- aminopropyltrimethoxysilyl chloride (DMOAP, Figure 6.1a).231 They reported the self-assembly of these microparticles as elongated chains or hexagonal lattices, depending on the presence of surfactant in the water phase. Later, Gharbi et al. studied the assembly of DMOAP-coated silica microparticles that imposed strong

114 homeotropic (perpendicular) boundary conditions at the LC/air interface. They concluded that the relative positions of defects formed near the microparticles led to distinct patterns such as elongated chains or hexagonal lattices. More recently, uncoated polystyrene microparticles that imposed planar (tangential) boundary

117-119 conditions were employed by Abbott and co-workers, who studied the ordering of microparticles at defect points of LC-in-water emulsion droplets.

Judging from these studies, there is strong interest in controlling microparticle assembly through interactions between microparticles and LCs at LC/water interfaces. In previous works, however, researchers focused on only uncoated surfaces or surfaces coated with a single layer of DMOAP. To further extend the possibility of forming assemblies of microparticles at LC/water interfaces, two coating layers on the surfaces of microparticles were employed in this work. The

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first layer was poly(ethylenimine) (PEI, Figure 6.1b) adsorbed on the surfaces of the microparticles, and the second layer was surfactants that reversibly bind to PEI through different interactions (electrostatic attraction or hydrogen bonding).

Because of this unique two-layer structure, it was possible to obtain different assembly patterns of microparticles. This approach also allowed for an investigation of how the binding strengths of surfactants affect the assembly patterns.

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6.2 Experimental Section

6.2.1 Materials

Glass slides were purchased from Fisher Scientific (U.S.A.). Silica microparticles

[5% (w/v), 4.40 ± 0.24 μm] were purchased from Microparticles GmbH (Germany).

N,N-Dimethyl-n-octadecyl-3-aminopropyltrimethoxysilyl chloride (DMOAP), poly(ethylenimine) [PEI; Mw ≈ 750000 Da, 50% (w/v) in water], polyoxyethylene

(20) sorbitan monolaurate (Tween 20), cetyltrimethylammonium bromide (CTAB), lipase from Candida rugose [type VII, activity > 700 U/(mg of protein)], and fluorescein 5-isothiocyante (FITC) were purchased from Sigma-Aldrich

(Singapore). Phosphate-buffered saline (PBS) and sodium dodecyl sulfate (SDS) were purchased from First BASE (Singapore). The LC 4-cyano-4′-n- pentylbiphenyl (5CB) was purchased from Merck (Singapore). Copper grids (100- mesh) for transmission electron microscopy (TEM) were purchased from Ted Pella

(U.S.A.).

6.2.2 Surface Modifications

Glass slides were soaked in a 5% (v/v) Decon 90 solution overnight to clean the surfaces. After this, the glass slides were sonicated three times for 15 min each in deionized (DI) water. Subsequently, the glass slides were immersed in a water solution containing 0.1% (v/v) DMOAP for 5 min and then rinsed with copious amounts of DI water. Finally, the DMOAP-coated glass slides were dried under a stream of nitrogen and then heated in a vacuum oven at 100 °C for 15 min.

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6.2.3 Coating of Microparticles with PEI

Microparticles (0.5%, w/v) were immersed in an ethanol solution containing 1%

(w/v) PEI for 3 h. After this, the solution was centrifuged briefly to collect the PEI- coated microparticles. Subsequently, DI water was added to wash the PEI-coated microparticles three times to remove remaining PEI solution. The PEI-coated microparticles were dispersed in DI water before use.

6.2.4 Labeling of PEI-Coated Microparticles with FTIC

PEI-coated microparticles were immersed in 1× PBS buffer containing FITC at a concentration of 1 μg/ml. After 1 h, the FITC-labeled microparticles were briefly centrifuged, and the FITC solution was removed. DI water was then added to wash the microparticles twice. After this, the microparticles were observed under a fluorescence microscope. Quantification of the fluorescence signal was done by using ImageJ software, and readings from 20 microparticles were used to obtain an average value for each sample. A calibration curve of the fluorescence intensity was obtained by measuring FITC solutions of known concentrations in a capillary tube (0.2 mm × 2.0 mm).

6.2.5 Zeta Potential Measurements of Microparticles

First, 0.5% (w/v) PEI-coated microparticles were diluted in 10 mM sodium chloride solution (pH 7.0) to a final concentration of 0.05% (w/v). Sodium chloride was used

235 to maintain the ionic strength of the solution. Surfactants were added to the

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aqueous phase, and the zeta potential of the microparticles was measured using a

Malvern Zetasizer Nano ZS instrument. For each data point, the average and standard deviation of 60 readings were used.

6.2.6 Microparticle Assembly at LC/Water Interfaces

First, copper grids (100-mesh) were supported on DMOAP-coated glass slides. The grids were filled with LC (5CB), and excess LC was removed using capillary tubes.

The LC-filled copper grids were immersed into water containing 90 μM Tween 20 for 30 min to allow the adsorption of Tween 20 at the LC/water interface. Next,

PEI-coated microparticles were added to the water phase to a final concentration of

0.1% (w/v), allowing them to sediment to the LC/water interface (Scheme 6.1a).

The optical appearance of microparticle assembly was observed using an optical microscope (Nikon ECLIPSE LV100POL, Japan), and images were recorded 10 min after the introduction of the microparticles. Intermicroparticle distances were measured based on the center-to-center distances between pairs of microparticles, and the average and standard deviation of 40 readings were obtained using ImageJ software.

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Scheme 6.1. Microparticles Assembly at LC/Water Interface

(a) Experimental setup for microparticle assembly at the LC/water interface. (b) Self-assembly of PEI-coated microparticles in the presence of Tween 20, which imposed weak homeotropic boundary conditions on the LC. (c) Self-assembly of PEI-coated microparticles in the presence of SDS, which imposed strong homeotropic boundary conditions on the LC and resulted in the formation of a hedgehog defect (represented by a black dot) near each microparticle.

6.2.7 Visual Detection of Lipase Activity Using Microparticle Assembly

To detect lipase activity through microparticle assembly, LC-filled copper grids were immersed into water containing 90 μM Tween 20 and lipase for 35 min. Next,

PEI-coated microparticles were added to the water phase to a final concentration of

0.1% (w/v), allowing them to settle on the LC/water interface. The optical appearance of microparticle assembly was observed using an optical microscope

(Nikon ECLIPSE LV100POL, Japan), and images were recorded 10 min after the microparticles were added.

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6.3 Results and Discussion

6.3.1 Surface Coating of Microparticles

To confirm that the microparticles were coated with PEI using fluorescence, microparticles coated with different concentrations of PEI were immersed into a 1

μg/ml FITC solution. After the microparticles had been rinsed to remove unreacted

FITC, fluorescence images of the microparticles were taken, and fluorescence intensities were analyzed using ImageJ software. Figure 6.2 shows that the fluorescence intensities increased with increasing PEI-coating concentration up to

1% (w/v). This result suggests that the surfaces of the microparticles were successfully coated with PEI and that the density of surface amine groups increased with increasing PEI-coating concentration. When the PEI-coating concentration was higher than 1% (w/v), the fluorescence intensity reached a plateau, suggesting that the surfaces of the microparticles were saturated with PEI already. Therefore,

1% (w/v) PEI solution was used for coating microparticles in all subsequent experiments. Under these coating conditions, we estimated that surface density of

2 reactive amine groups was approximately ∼0.03/nm (see Appendix B.1). The stability of the coated PEI layers was tested with repetitive rinsing. After two additional washing cycles, a decrease in fluorescence intensity of only ∼10% was observed. This result implies that the PEI layers were stable.

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Figure 6.2. Effect of PEI-coating concentration on the surface density of reactive amine groups. Inset: Calibration plot of fluorescence intensity obtained using standard FITC solutions.

6.3.2 Adsorption of Surfactants on PEI-Coated Microparticles

To study the adsorption of surfactants on PEI-coated microparticles, we measured the zeta potentials of PEI-coated microparticles in the presence of different surfactants at different concentrations. In the absence of surfactant, the zeta potential of the microparticles was 62.0 ± 1.2 mV (Figure 6.3). Because PEI is positively charged at neutral pH, a positive zeta potential agrees well with the model. In the presence of SDS, the zeta potential decreased quickly with increasing

SDS concentration, suggesting that the negatively charged surfactant adsorbed onto the PEI-coated microparticles because of electrostatic attraction. For Tween 20, the zeta potential also decreased with increasing Tween 20 concentration. Because PEI contains nitrogen atoms and Tween 20 contains oxygen atoms, the decrease in zeta potential was probably caused by the adsorption of Tween 20 onto the PEI-coated microparticles through hydrogen bonding. This result is consistent with earlier

236-238 work showing that nonionic surfactants adsorb on hydrophilic surfaces. For

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instance, Zhang and co-workers proposed that nonionic surfactants adsorb on hydrophilic surfaces through hydrogen bonding. They also observed a slight

239 decrease in zeta potential after the addition of non-ionic surfactants. In contrast, the zeta potential did not decrease in the presence of CTAB. This is reasonable because both PEI and CTAB are positively charged and repel each other. These results suggest that only SDS and Tween 20 can adsorb on the surfaces of PEI- coated microparticles.

Figure 6.3. Influence on the zeta potential of the adsorption of surfactants on PEI- coated microparticles. The surfactants studied were (a) cationic CTAB, (b) nonionic Tween 20, and (c) anionic SDS.

6.3.3 Self-Assembly of PEI-Coated Microparticles at LC/Water Interfaces

To initiate the self-assembly of PEI-coated microparticles, we immersed an LC- filled grid into water. The LC/water interface was decorated with 90 μM Tween 20.

Next, we added PEI-coated particles to the solution to a final concentration of 0.1%

(w/v). Within 5 min, the microparticles settled onto the LC/water interface and started to assemble into short chains (each chain contained up to 20 microparticles),

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as shown in Figure 6.4a. The intermicroparticle distance was 4.61 ± 0.21 μm, which is very close to the diameter of the microparticles used (4.40 ± 0.24 μm). This showed that there was no spacing between any pair of adjacent microparticles in the short chains. In contrast, when the LC/water interface was not decorated with

Tween 20 (Figure 6.4b) or when the microparticles were not coated with PEI

(Figure 6.4c), the microparticles did not assemble into short chains. As shown in

Figure 6.4b,c, the microparticles appeared to organize into two-dimensional hexagonal structures. The intermicroparticle distances were 4.74 ± 0.14 and 4.71 ±

0.28 μm in panels b and c, respectively, of Figure 6.4 (there was little or no space between any pair of adjacent microparticles). This resulted in the close packing of the microparticles at the center of the grid hole (because of a concave meniscus).

Moreover, if the LC was heated above its clearing temperature (35 °C for 5CB), the microparticles also did not assemble into short chains (Figure 6.4d). Instead, they moved either to the center of the grid (because of a concave meniscus) or to the side (Figure 6.4e). These controlled experiments led us to conclude that all three elements (PEI coating, Tween 20, and LC phase) are essential to the assembly of microparticles into short chains.

Figure 6.4a also shows that the directions of the short chains all pointed toward a single point in the LC. This behavior is similar to the LC director field when a point defect is present in an LC. To obtain the LC field, the LC sample was rotated under crossed polars (Figure 6.5a) to determine the direction of the LC director field.

Figure 6.5b shows that the LC director field also converged to a single point defect

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in the center of the square. By comparing panels a and b of Figure 6.5, one can see that the directions of the short chains followed the LC director field. This result is consistent with past studies showing that microparticles with homeotropic boundary conditions assemble into short chains that follow the LC director field.114,

216, 231

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6.3.4 Effect of Tween 20 Concentration on Microparticle Assembly

To understand the effect of the Tween 20 concentration on the self-assembly of

PEI-coated microparticles, we varied the concentration of Tween 20. Figure 6.6 shows that 90 μM was the minimum Tween 20 concentration required to trigger the self-assembly of PEI-coated microparticles into short chains. Interestingly, when the Tween 20 concentration was 45 μM, the texture of the LC was not disrupted by the microparticles (Figure 6.6a). The LC texture was very continuous even in the region with microparticles. Additionally, adjustment of the focal planes showed that the microparticles and the LC/water interface were not at the same focal plane.

This result implies that the microparticles did not penetrate the LC interface when the Tween 20 concentration was too low. In contrast, Figure 6.6b (and the magnified view in Figure 6.6g) shows that, when the Tween 20 concentration was increased to 90 μM, the microparticles started to form short chains and were surrounded by a white halo in the LC phase. The white halo is strong evidence that these microparticles were able to submerge into the LC and distort its orientation.

When the Tween 20 concentration was between 900 μM and 4.5 mM, short chains

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of microparticles aggregated together and formed bands of three to five chains

(Figure 6c,d). This can be explained by the increasing surface density of Tween 20 adsorbed on the PEI-coated microparticles, leading to the aggregation of PEI- coated microparticles to form thicker bands. Again, white halos surrounding the microparticles are clearly visible in these cases.

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However, when the Tween 20 concentration was 18 mM, the LC/water interface was fully covered by Tween 20. This was evident from the uniformly dark image

102, 240 of the LC under crossed polars (Figure 6.6e). Under these conditions, all PEI- coated microparticles assembled at the center of the grid hole (because of a concave

241-242 meniscus), as shown in Figure 6.6f. This implies that, when the LC/water interface was fully covered with Tween 20, the PEI-coated microparticles did not submerge into the LC. This conclusion is supported by the uniform texture of the

LC under crossed polars (Figure 6.6e) and under a bright field (Figure 6.6f). Both images strongly suggest that, when the LC interface was saturated with Tween 20, the microparticles were prevented from breaking the LC interface.

Previous reports have suggested that nonionic surfactants (such as Tween 20) have

−6 2 −2 a weak anchoring strength (∼10 J/m ) compared to ionic surfactants (∼10

2 214, 232, 243 J/m ). Therefore, we propose that the short-chain assembly is due to the weak homeotropic boundary conditions imposed by microparticles covered with

Tween 20. When such a microparticle is placed at the LC/water interface, it is able to penetrate into the LC phase and cause a distortion of the director, as shown in

Scheme 6.1b. However, the anchoring strength is not strong enough to cause a hedgehog defect, as in the case of strong homeotropic anchoring. Therefore, we hypothesize that the LC director around the microparticles resembles surface rings,

243-244 as suggested previously by Mondain-Monval et al. and Stark. For surface rings, the LC director field is minimally distorted by the microparticles, as compared to the case of hedgehog defects. Although Saturn ring defects are also

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probable, we did not observe these defects in our experiments. When several microparticles were present with surface rings, they aggregated together to form short chains, as shown in Scheme 6.1b.

6.3.5 Visual Detection of Lipase Activity in Water

Because the assembly of the PEI-coated microparticles is dictated by the surfactants adsorbed on their surfaces, this phenomenon can be applied to detect any lipases

245-247 that hydrolyze Tween 20. We anticipated that, after the hydrolysis of Tween

20, microparticles would no longer be able to assemble into short chains, as shown in Figure 6.6b. To test this hypothesis, we added lipase to water at concentrations between 500 and 10 ng/ml. As shown in Figure 6.7a−d, when the lipase concentration was 100 ng/ml or higher, the microparticles did not form short chains.

Instead, they assembled at the center of the grid hole, similarly to the case of Figure

6.6a (when the concentration Tween 20 was lower than the minimum required).

However, when the lipase concentration was 10 ng/ml, the microparticles still assembled into short chains. These results show that a lipase concentration of 100 ng/ml is sufficient to hydrolyze Tween 20 and make its concentration fall below its minimum concentration for microparticles to form short chains.

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6.3.6 Effects of Cationic and Anionic Surfactants

To further study whether PEI-coated microparticles can self-assemble at LC/water interfaces decorated with cationic or anionic surfactants, we deposited PEI-coated microparticles at LC/water interfaces decorated with either SDS (anionic surfactant) or CTAB (cationic surfactant, Figure 6.1e). The concentrations of SDS and CTAB were 90 and 9 μM, respectively, corresponding to approximately 1% of their critical micelle concentrations. As can be seen in Figure 6.8a,b, the PEI-coated microparticles assembled into circular rings at the LC/water interface decorated with SDS. Interestingly, this microparticle assembly pattern was distinctly different from those observed in the presence of Tween 20 (Figure 6.6b). On the other hand, when CTAB was used, the microparticles assembled at the center of the grid hole, independent of the LC director field (Figure 6.8c,d). This was probably because positively charged CTAB cannot adsorb onto the surfaces of PEI-coated microparticles, as shown by the zeta potential measurements.

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To understand how the microparticles assemble in the presence of SDS, we observed the microparticles under a higher magnification (Figure 6.8e) and found that the LC surrounding the microparticles appeared dark under crossed polars. This phenomenon implies that the microparticles imposed strong homeotropic boundary conditions on the LC. Because the PEI microparticles were positively charged, they were able to attract the negatively charged SDS to their surfaces, as demonstrated by the zeta potential measurements. (In the presence of 100 μM SDS, the zeta potential decreased from 62.0 ± 1.2 to −54.4 ± 1.8 mV.) Moreover, hedgehog

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defects near the microparticles can be seen in Figure 6.8e. In the literature, it has been reported that microparticles with strong homeotropic boundary conditions can

216, 219 form hedgehog defects near the microparticle surface. Additionally, the intermicroparticle distance was 5.9 ± 0.7 μm, which agrees well with previous studies showing an intermicroparticle distance of 1.3d (where d is the diameter of the microparticle) for microparticles that impose strong homeotropic boundary

215, 231 conditions. Another notable difference is the significantly larger intermicroparticle distance compared to the case of short chains formed in the presence of Tween 20. Based on these observations, we propose a model for the assembly of PEI-coated microparticles in the presence of SDS. As shown in

Scheme 6.1c, after the formation of a hedgehog defect near a microparticle, the microparticles behave like dipoles in the LC field and attract other microparticles.

Eventually, these microparticles chain up together and form a linear pattern, as shown in Scheme 6.1c.

Another interesting phenomenon shown in Figure 6.8b is the dark domains surrounded by microparticles. We attribute the dark domains to a saturated

102, 240 monolayer of SDS adsorbed at the LC/water interface. This phenomenon strongly implies that PEI-coated microparticles at the LC/water interface are able to “squeeze” SDS into a saturated domain. Because SDS is a surfactant, it can lower the surface tension and compact SDS into a saturated domain through a surface tension gradient. The formation of surfactant-rich domains at LC/water interfaces has also been observed by other research groups. For example, Abbott and co-

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workers discovered phase separation of phospholipids and ionic surfactants at

248-250 LC/water interfaces. In particular, Gupta and Abbott demonstrated the phase

250 separation of ionic surfactants in a high salt concentration (1 M NaBr). They proposed that a high salt concentration is needed to screen the electrostatic repulsion of charged surfactants, allowing them to aggregate and form separate domains. In our system, however, the phase separation was initiated by the introduction of PEI-coated microparticles. Furthermore, we note that long chains of microparticles circled the perimeters of the saturated SDS domains (Figure 6.8b).

A possible explanation is that the LCs oriented homeotropically in the saturated

SDS domain whereas the LCs oriented planarly outside these domains. Therefore, the orientation of the LCs experiences homeotropic-to-planar deformation (Figure

6.8b), which comes with a high energy penalty. Previous works suggested that microparticles can be attracted to regions with the greatest LC deformation to

244, 251 minimize the energy. Therefore, this can explain why all of the microparticles moved to the perimeters of the saturated SDS domains.

6.3.7 Effect of SDS Concentration on Microparticle Assembly

To understand the effect of the SDS concentration on the self-assembly of the PEI- coated microparticles, we varied the concentration of SDS. At a lower SDS concentration (0.9 μM), no distinct microparticle assembly was observed (Figure

6.9a), likely because of the limited coverage of SDS on the PEI-coated microparticles. When the SDS concentration was 9 μM, the PEI-coated microparticles assembled into short chains. However, no dark domains formed

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(Figure 6.9b). As shown in our model (Scheme 6.1c), the short chains were caused by hedgehog defects attracting each other (Figure 6.9b). At 90−270 μM SDS, the

PEI-coated microparticles elongated and formed dark domains, as seen in Figure

6.9c,d. These dark domains appeared to increase as the SDS concentration was increased. The increase in SDS concentration caused more SDS to adsorb on the

PEI-coated microparticles and led to a higher SDS coverage at the LC/water interface. Both factors are responsible for the expanding dark domains. At 450 μM

SDS, the PEI-coated microparticles assembled at the center of the grid hole independent of the LC director field (Figure 6.9f). In Figure 6.9e, the dark cross- polarized image at 450 μM SDS indicates that the LC/water interface was saturated with SDS. The saturated coverage of SDS at the LC/water interface shielded the

LC phase and prevented interactions between the microparticles and the LC phase.

Such an observation is similar to the case of microparticle assembly at an LC/water interface decorated with 18 mM Tween 20 (Figure 6.6f).

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6.4 Conclusion

In this study, we found that poly(ethylenimine)- (PEI-) coated microparticles are able to assemble into different patterns at the liquid crystal (LC)/water interface.

However, this phenomenon was observed only when the interface was decorated with surfactants Tween 20 or sodium dodecyl sulfate (SDS), and the assembly pattern was very sensitive to the type of surfactant used. The two surfactants are able to adsorb on the surfaces of PEI-coated microparticles and interact with the

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director field of the LC to give different assembly patterns. Previous studies on microparticle assembly at the LC/water or LC/air interface used the approach of

114, 231 controlling the LC director field to achieve different assembly patterns.

However, such an approach relies on the use of microparticles coated with N,N- dimethyl-n-octadecyl-3-aminopropyltrimethoxysilyl chloride (DMOAP) that impose strong homeotropic boundary conditions on the LC. This limitation was addressed in this study, where we made use of PEI-coated microparticles that interact with surfactants present in the water phase to achieve different assembly patterns. Significantly, this study also suggests the potential of utilizing microparticle assembly at the LC/water interface for sensing applications, as exemplified through the detection of lipase activity by examining the assembly patterns, with a lipase detection limit of 100 ng/ml.

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CHAPTER 7 CONCLUSIONS AND RECOMMENDATIONS

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7.1 Conclusions

Even though there are already many types of protease assays under development, there is still no single protease assay that can meet all requirements. An ideal protease assay should be fast, sensitive, multiplexed, and robust. In this thesis, a new type of LC-based, heterogeneous protease assay is developed. LC-based protease assays provide facile optical readout system with multiplex sensing capabilities. They are customizable for different target proteases through the design of specific oligopeptide substrates. LC-based assays are highly sensitive to protease activities, as LC responds and amplifies cleavage of immobilized oligopeptide substrates into optical signals. These signals arise from birefringence of LC, and this property causes the appearance of the bright optical signal under cross- polarized light source. Such signal generation principle is photo-stable, and does not require intricate spectroscopy measurements. Thus, LC-based assays can address the requirements of an ideal protease assay. In this thesis, we provided the foundation for a robust and sensitive LC-based assay that can be generically applied for different target proteases.

In Chapter 3, we carried out a detailed mechanistic study on immobilization of oligopeptides and developed a technique for specific immobilization of oligopeptides through cysteine residues. It was concluded that nucleophilic addition of thiolate ions from cysteine to aldehyde groups present on the surfaces was responsible for the immobilization of cysteine-bearing peptides. Furthermore, this technique provided immobilized oligopeptides which were highly stable; there was

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CHAPTER 7 no significant loss of immobilized oligopeptides under assaying conditions. This study addressed stability issues previously encountered, and provides the foundation for incorporation of oligopeptides in LC-based protease assays.

In Chapter 4, we developed a working principle for the LC-based protease assay.

The principle of LC-based protease assay stems from the influence of oligopeptide- immobilized surfaces on LC orientation. Upon cleavage by target protease, oligopeptide fragments remaining on the surface will no longer influence LC. This causes LC to orient from planar to homeotropic, and this is transduced as bright-to- dark optical signals. However, there are uncertainties to whether oligopeptide- immobilized surfaces influence LC to cause planar orientations. Thus, we introduced a two-pronged approach that overcomes this issue. First, 30-mer peptides were incorporated with two N-terminal cysteines to serve as immobilization sites and inter-peptide cross-linking through disulfide bonds. This increased peptide immobilization density. Second, C-terminals were biotin-labeled to allow specific binding with streptavidin. This presents the surface with large streptavidin proteins (~52.8 kDa), compared to the smaller peptides (~3 kDa). In the presence of target proteases, cleavage of biotin-labeled peptides presented the surface with smaller peptide fragments (<3 kDa), and the sharp difference in surface composition instilled greater reliability and sensitivity to the LC-based protease assays. This two-pronged approach was tested against trypsin and chymotrypsin, and we reported a LOD of 0.1 ng/ml and 1 ng/ml respectively, which are better than

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In Chapter 5, we demonstrated how interactions between microparticles and LC phase provided distinct optical textures that were capable of protease sensing. This is motivated by greater interactions between LC and microparticles, given the confinement of microparticles within the LC phase. Many researchers reported the formation of unique LC textures around microparticles presenting different boundary conditions, and this phenomenon was applied in this protease assay. This was achieved by immobilization of octadecyl- (C18)-terminated oligopeptides on microparticles. Strong homeotropic boundary condition imposed by C18-terminated oligopeptides caused LC near the microparticles to be distorted greatly, resulting in observable LC texture. Subsequently, cleavage of oligopeptides by target protease removed the C18 groups and resulted in drastic changes in LC texture. Therefore, this study suggests an alternative approach to LC-based assay, and departs from the canonical platform of assays based on flat surfaces. The greater interactions between microparticles and LC, compared to flat glass surfaces, also suggest an alternative amplification mechanism to improve the sensitivity of LC-based assays.

In order to better study and monitor protease activities, real-time protease assays are valuable tools. Thus, in Chapter 6, we developed a platform for real-time, LC- based protease assay. Such assays require LC interface to be in constant contact with an aqueous phase containing proteases and their substrates, thereby allowing

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LC to respond to any protease activities in real-time. However, such assays require adsorption of oligopeptide substrates at LC/water interface, which is more complex and less-studied. Furthermore, we would require an amplification mechanism to improve the sensitivity of the assay. Thus, a proof-of-concept study was done to investigate the use of microparticles assembly at LC/water interface in detecting the presence of target analytes or enzymatic activities. We achieved this by the introduction of PEI-coated microparticles that interact with surfactants present in the water phase. Based on the nature of adsorption (electrostatics or hydrogen bonding) of surfactants on PEI-coated surfaces, we observed vastly different microparticles assembly. This showed the potential of using microparticles assembly to detect target analytes in the water phase. We further exemplified sensing capabilities of such microparticles assembly to detect lipase activity. Lipase was used as a model target enzyme, since it catalyzes the hydrolysis of Tween 20, which was a non-ionic surfactant used in this study. Through this work, we gained a greater understanding of the interactions between microparticles and LC, and demonstrated their potential for real-time sensing applications.

7.2 Recommendations

Based on studies carried out in this thesis, we provide some recommendations for future work:

(1) Based on the working principles proposed in Chapter 3 and 4, more

clinically-relevant proteases can be tested to validate our assay’s sensitivity.

For example, MMPs appear to attract great attention due to their role in

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cancers. Probably, they can also be detected by using the LC-based protease

assay developed in Chapter 4. Furthermore, multiplexed sensing can be

carried out to study the selectivity of LC-based assays in the presence of

multiple proteases.

(2) As shown in Chapter 4, the introduction of streptavidin as part of an

amplification mechanism improved the detection limits. Thus, more

amplification mechanisms can be explored to improve the sensitivity and

detection limit of LC-based assays. For example, C18-terminated

oligopeptides can be studied for their ability to amplify LC signals (as

observed in Chapter 5). These C18-terminated oligopeptides inherently

impose homeotropic boundary condition on LC. Therefore, cleavage of

these oligopeptides will cause a dark-to-bright change in a typical LC-based

assay on flat surfaces. Additionally, this sensing principle circumvents

limitations of typical LC-based assays, where surfaces required DMOAP

coating to provide a dark background and contrast for oligopeptide-

immobilized spots. In this case, surfaces do not require DMOAP coating,

since cleavage of C18-terminated oligopeptides can provide dark-to-bright

changes to LC optical for detection. Therefore, more surface

functionalization methods can be explored to optimize oligopeptide

immobilization.

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(3) Based on microparticles assembly at LC/water interface in Chapter 6, we

demonstrated how microparticles assembly can detect lipase activity. This

platform can be developed into real-time protease assays, where the

introduction of oligopeptides on PEI-coated microparticles can potentially

allow real-time monitoring of protease activity in the aqueous phase. Based

on findings in Chapter 5, we can immobilize C18-terminated oligopeptides

on microparticles, and subsequent cleavage of oligopeptides by proteases

will alter surface compositions. These changes in surface composition can

potentially result in observable changes in microparticles assembly, since

LC are strongly influenced by C18-terminated oligopeptides. Therefore, we

will be able to carry out real-time protease assay, through monitoring of

microparticles assembly. Additionally, in Chapter 6, the formation of

circular SDS domains in the presence of PEI-coated microparticles is an

interesting finding that requires in-depth studies to identify the driving force

for such phase separations.

(4) To leverage on the facile signal readout and high sensitivity of LC-based

protease assays, efforts should be devoted to the development of point-of-

care (POC) protease sensors. LC-based sensors can potentially be

miniaturized, operate with minimal instrumentation, and provide facile

signal readout system. These characteristics fulfill the basic requirements of

a POC sensor. Furthermore, this thesis showed that the LOD of the LC-

based protease assays is comparable (or even better) to other detection

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techniques reported. Thus, LC-based protease assays can be potentially

developed into a POC protease sensor.

(5) It is important to carry out assays based on biologically-relevant (“real”)

samples, where a complex matrix of biomolecules may adversely affect

detection capabilities of the LC-based protease assays. In particular, the

presence of amphiphilic biomolecules like proteins and lipids may non-

specifically adhere to surfaces of glass and microparticles, and this will

prevent the interactions between peptide-functionalized surfaces and target

proteases. Furthermore, since LC responses are very sensitive to the

presence of amphiphilic molecules, their presence will likely interfere and

result in false assay results. To circumvent this, several strategies can be

employed. First, a pretreatment step can be included to remove some of the

complex materials present in the real samples. However, it is important to

ensure that the activities of the target proteases are not affected by the

pretreatment steps. This will be applicable for both end-point measurement

assays (in Chapters 3 to 5) and potential real-time monitoring assays based

on work presented in Chapter 6. Next, for end-point measurement LC-based

assays, a stringent washing protocol will be required to remove non-

specifically bounded materials on the surfaces. However, to ensure an

accurate assay result, it will be important to ensure that the peptide-

functionalized surfaces are not affected by the stringent washing. Finally, it

is important to have a built-in control mechanism to ensure that results of

144

CHAPTER 7 the assays are solely due to the presence of targets. For example, in the end- point measurement assays, peptide-functionalized surfaces should include control peptides that are non-cleavable by any protease present in the real samples. This will ensure that assay results are reliable, and not due to the unexpected removal of immobilized-peptides in the real samples. Similar built-in control should also be included for potential real-time monitoring protease assays based on work presented in Chapter 6.

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171

APPENDICES

Appendix A

A.1 Effect of Coating Duration on Water Contact Angle of DMOAP-Coated

Surface

Water Contact angle were measured by VCA Optima goniometer from AST

Products (Billerica, U.S.A.). For each measurement, 1 μl of deionized water was introduced to the DMOAP-coated surfaces, and the water contact angle was measured using the software provided by the manufacturer. Each data point presented was determined based on the average of 10 measurements. In these measurements, 0.1% (v/v) DMOAP coating solution was used, and the coating duration was varied from 0 to 60 min.

In order to study the homogeneity of the DMOAP coating, water contact angle measurements were carried out with increasing DMOAP incubation durations from

0 min to 60 min. From Figure A.1, it is supported that after 5 min of incubation, the resulting water contact angle was 82.0±2.4°. Further increase in incubation period to 60 min only resulted in a slight increase of water contact angle to 91.3±1.0°.

Therefore, this results supports that a 5 min DMOAP incubation period is required for a homogenous coverage of the glass surfaces. This conclusion is also supported by previous studies,93 it was suggested that a coating duration of 5 min allows the formation of a uniform monolayer of DMOAP.

172

APPENDICES

Figure A.1. Effect of DMOAP coating duration on the water contact angles.

173

APPENDICES

A.2 Effect of UV Irradiation on Immobilization of P2 and P3

For oligopeptides P2 and P3, variations of UV irradiation duration did not produce fluorescence intensities that were similar to P1, P4 and P5 (Figure A.2). This was likely attributed to the lack of –SH groups present in P2 and P3 for immobilization.

Figure A.2. Fluorescence intensity of immobilized P2 and P3 for different UV durations.

174

APPENDICES

A.3 Determination of Oxidation of Free Thiols

175

APPENDICES

Appendix B

B.1 Surface Amine Density Calculations

To calculate the surface amine densities, fluorescein 5-isothiocynate (FITC) was used to react with surface amines on poly(ethylenimine) (PEI)-coated microparticles and fluorescence intensities were recorded. Next, standard calibration curve was obtained for fluorescence intensities of FITC solutions of varying concentrations (0, 10, 25, 50, 75, 100 and 150 ng/ml) in a rectangular capillary tube (height × width = 0.2 mm × 2 mm). Relationship between surface densities of FITC, , and FITC in solution were obtained using the following equation:252

 (B.1)

Where C is the concentration of FITC in solution, h is the height of rectangular

6 9 capillary tube (0.2 × 10 nm), Mr is the molecular weight of FITC (389.4 × 10

23 ng/mol) and NA is Avogadro’s number (6.02 × 10 1/mol).

The standard calibration plot of fluorescence intensities vs. surface densities was obtained, and this was used to quantify the surface density of amine groups on PEI- coated microparticles.

176

LIST OF PUBLICATIONS

1. Ong, L. H., Ding, X. Yang, K. L., Mechanistic Study for Immobilization of

Cysteine-Labeled Oligopeptides on UV-Activated Surfaces. Colloids Surf. B

2014, 122, 166-174.

2. Ong, L. H., Yang, K. L., Surfactant-Driven Assembly of Poly(ethylenimine)-

Coated Microparticles at the Liquid Crystal/Water Interface. J. Phys. Chem. B

2016, 120, 825-833.

3. Ong, L. H., Yang, K. L., Liquid Crystal-Based Protease Assays with Dual

Cysteine-Labeled Peptides for Improving Peptide Stability. Manuscript submitted.

4. Ong, L. H., Yang, K. L., Recent Developments in Functional Protease Assays and Sensors. Manuscript under preparation.

5. Ong, L. H., Yang, K. L., Distortion of Liquid Crystal Phase by Microparticles

Functionalized with Octadecyl-Terminated Oligopeptides. Manuscript under preparation.

177