A Dissertation Entitled Selection and Characterization of Ssdna Aptamers for Salivary Peptide Histatin 3 and Their Application T

A Dissertation

entitled

Selection and Characterization of ssDNA Aptamers for Salivary Peptide Histatin 3 and

Their Application Towards Assay and Point-of-Care Biosensing

Yagya Raj Ojha

Submitted to the Graduate Faculty as partial fulfillment of the requirements for the

Doctor of Philosophy Degree in

Biomedical Engineering

______Brent D. Cameron, PhD, Committee Chair

______David R. Giovannucci, PhD, Committee Member

______Dong-Shik Kim, PhD, Committee Member

______Scott M. Pappada, PhD, Committee Member

______Casey W. Pirnstill, PhD, Committee Member

______Amanda C. Bryant-Friedrich, PhD, Dean College of Graduate Studies

The University of Toledo December 2019

This document is copyrighted material. Under copyright law, no parts of this document may be reproduced without the expressed permission of the author. An Abstract of

Selection and Characterization of ssDNA Aptamers for Salivary Peptide Histatin 3 and Their Application Towards Assay and Point-of-Care Biosensing

Yagya Raj Ojha

Submitted to the Graduate Faculty as partial fulfillment of the requirements for the Doctor of Philosophy Degree in Biomedical Engineering

The University of Toledo December 2019

The development of detection methods for the novel biomarkers can have a significant impact on the research and clinical applications such as drug discovery, disease diagnosis, and treatment monitoring. Histatin 3 (H3) is an antimicrobial salivary peptide that possesses the capability of being a therapeutic agent against oral candidiasis and has recently been linked to acute stress as a potential novel biomarker. Stress biomarkers reflect the physical and cognitive performance of an individual, and their monitoring in real-time is of vital importance for the high-risk jobs, including military, pilot, and surgeon, where higher vigilance is required for an extended period. The salivary levels of H3 also have been correlated with the HIV-infection and associated oral candidiasis. Therefore, monitoring H3 levels in human saliva can provide essential information about an individual’s health status, including HIV-infection, oral candidiasis, and acute stress.

Additionally, H3 detection could serve as therapeutic drug monitoring if H3 can be established as an alternative therapeutic agent.

The currently available detection techniques for H3 are gel chromatography, high- performance liquid chromatography (HPLC), mass spectrometry (MS), and antibody-

iii based immunoassays. The Chromatographic and mass-spectroscopic methods are laborious, utilize expensive instrumentation, require trained personnel, and time- consuming. Whereas antibody-based immunoassays are not widely validated, expensive, sensitive to temperature, and have a short lifespan. This void in analytical methods is not just for H3 but also applies to several other biomarkers in saliva. Even though saliva is considered as an optimal biofluid, several limitations are impeding its use in diagnostic and research. The major hurdles include the deficient concentration of biomarkers, need of laboratory-based preprocessing to remove mucin and interfering particulate matters, and lack of standard sample collection methods. Also, due to the absence of easily accessible and cost-effective analytical methods, it is challenging to figure out the circadian variation of biomolecules and consequently establish biomolecules as biomarkers. All these necessities mandate the development of alternative techniques that offer more accessible, cost-effective, rapid, real-time, and probably the point-of-care (POC) testing of salivary biomarkers. The aptamers, synthetic oligonucleotides functionally similar to antibodies but with several advantages, have shown promising possibilities to fulfill the requirements in such applications. However, no aptamer for H3 and several other salivary biomarkers exist in literature.

This dissertation presents a widely applicable aptamer selection method for the novel biomarkers and the application of the identified aptamers with various platforms for the detection application in human saliva. In order to identify H3 aptamers, a library immobilization version of an iterative in vitro process known as the systematic evolution of ligands by exponential enrichment (SELEX) was established. This method does not require target immobilization and offers the isolation of signaling aptamers for a target in

iv its unmodified form. Also, the signaling, structure-switching upon target binding, is a highly desired attribute of the aptamers for the biosensing application. Within the SELEX process, counter-selection steps were instituted to stop the evolution of nonspecific sequences that can bind to the molecules closely related to the target. Through the repetitive rounds of selection and counter-selection, four unique aptamer candidates sharing a consensus sequence were identified. The obtained aptamers were analyzed via online servers to predict the stability and structural conformations. A superior candidate in terms of stability and structural complexity was picked for the visual confirmation of binding.

The magnetic bead-based affinity capture combined with sodium dodecyl sulfate- polyacrylamide gel electrophoresis (SDS-PAGE), confirmed the selective binding between the selected aptamer candidate and target H3, indicating the successful aptamer selection process.

Following the preliminary studies, more detailed characterizations were performed by developing a direct format enzyme-linked aptamer sorbent assay (ELASA). The aptamers were compared for their binding affinity and specificity to the target molecule

H3. Also, the effect of primer binding sites on the binding affinity was probed. Selected clones exhibited affinity and specificity to H3, and the sequence with the highest affinity exhibited enhanced binding following the removal of primer binding sites. Based on these results, the truncated aptamer sequence was chosen for further application with other assay platforms.

Next, the signaling feature of the aptamer was employed to explore the prospect of gold nanoparticles (AuNPs)-based colorimetric assays. Two different formats of the colorimetric assays were developed. After optimizing various parameters, both assay

v schemes were able to detect H3 within a salivary level of a healthy individual. Besides, with the assay-format-2, the overall assay time was reduced to just 12 minutes.

Furthermore, surface-enhanced Raman scattering (SERS)-based magnetic aptasensor was developed for the ultrasensitive detection of H3. By optimizing various parameters, SERS-based atpasensor was established. Additionally, the lateral flow test strip to support POC measurements of H3 was demonstrated by utilizing the optical properties of the AuNPs and the specificity of the identified H3 aptamer. The captured

AuNPs on the test line of the lateral flow device provided qualitative analysis within 5 minutes for visual observation. Finally, human saliva and the artificial saliva samples were tested with the developed assay formats to investigate the challenges and the complications associated with the saliva-based measurements.

In conclusion, the work presented in this dissertation lays the foundation for the identification of the signaling aptamers and their subsequent implementation in several platforms for the detection of salivary biomarkers. Further improvements and optimizations of the assay methods will eventually lead to a fast, reliable, cost-effective, ultrasensitive, and POC detection methods for the salivary biomarkers.

To my parents, Dhananjay and Indu, brother Dipak, and wife Dikshya for their persistent love and encouragement.

vii

Acknowledgements

First and foremost, I would like to take this opportunity to express my deepest gratitude to my advisor Dr. Brent D. Cameron for being a great mentor. This work has only been possible with his continuous support, scientific insight, and supervision. I feel truly fortunate to have had him as my advisor. Also, my sincere thanks go to the committee members, Dr. David R. Giovannucci, Dr. Dong-Shikh Kim, Dr. Scott M. Pappada, and Dr.

Casey W. Pirnstill, for their invaluable words of advice and thoughtful comments. It is my great pleasure to express my appreciation to Tamara Phares for her unconditional support throughout all these years. Further, I am thankful to the faculty, staff, and my friends,

Yongsoon Hwang, Niraj K. Gupta, Mahmoud Eladawi, Ujjwal Shrestha, and Nav Raj

Paneru, for providing friendly advice and useful comments.

I’m deeply indebted to my father; with his support and inspiration, I have been able to make it to this point. Special thanks go to my mom, brother, wife, and other family members for caring, loving, and believing in me. I am also thankful to my relatives in my home country, Nepal, for their love, support, and encouragement throughout my journey.

I would also like to thank the Air Force Office of Scientific Research (AFOSR) for sponsoring the project (grant contract FA955-13-1-0187) included in this dissertation.

Finally, I would like to acknowledge the Center for Materials and Sensor Characterization

(CMSC) and Electron Microscope Facility (Dr. W.T. Gunning) at the University of Toledo for the instrumentation and testing support. viii

Table of Contents

Abstract…………………………………………………………………………………iii

Acknowledgements ...... viii

Table of Contents ...... ix

List of Tables ...... xvi

List of Figures ...... xvii

List of Abbreviation ...... xxii

1 Introduction ...... 1

1.1 Background and significance ...... 1

1.2 Research objectives ...... 7

2 Literature Review...... 10

2.1 Aptamers ...... 10

2.2 Generation of aptamers by systematic evolution of ligands by exponential

enrichment (SELEX) ...... 14

2.2.1 Aptamer library ...... 14

2.2.2 Selection ...... 16

2.2.3 Amplification and strand separation ...... 22

2.2.4 Cloning and sequencing ...... 24

2.3 Limitations of SELEX ...... 25

2.4 Aptamer characterization ...... 26

2.5 Applications of aptamers in various assay platforms ...... 27

2.5.1 Magnetic bead (MB)-based affinity capture coupled to sodium

dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) ...... 27

2.5.2 Enzyme-linked aptamer sorbent assay (ELASA) ...... 30

2.5.3 Colorimetric assays based on unmodified gold nanoparticles

(AuNPs) and aptamers ...... 34

2.5.4 Surface-enhanced Raman scattering (SERS)-based magnetic

aptasensor ...... 38

2.5.5 Aptamer-based lateral flow test strip ...... 46

3 Materials and Methods ...... 50

3.1 Selection and preliminary characterization of structure-switching ssDNA

aptamers for salivary peptide histatin 3 (H3) ...... 50

3.1.1 Aptamer selection by immobilizing ssDNA library sequences on

the MBs ...... 51

3.1.1.1 Oligonucleotides for the SELEX ...... 51

3.1.1.2 Preparation of ssDNA library-immobilized MBs ...... 51

3.1.1.3 In vitro selection procedure ...... 52

3.1.1.4 Polymerase chain reaction (PCR) amplification ...... 55

3.1.1.5 Transformation, cloning, and sequencing ...... 55

3.1.2 Analysis of aptamer sequences, structural folding, and stability .. 56

3.1.3 Demonstration of binding through MB-based affinity capture assay

coupled to SDS-PAGE ...... 56

3.2 Integration and demonstration of the H3-selective aptamers into ultrasensitive assay formats to allow for the identification and quantification in complex media 58

3.2.1 Development of ELASA for the additional investigation into the

binding between the selected aptamer candidates and H3 ...... 59

3.2.1.1 Aptamers ...... 59

3.2.1.2 Binding analysis using ELASA ...... 59

3.2.1.3 Binding kinetics study...... 61

3.2.1.4 Determination dissociation constant ( Kd ) ...... 61

3.2.2 Development of AuNPs-based colorimetric assays based on the

structural feature of the identified H3 aptamers ...... 62

3.2.2.1 Assay format-1 ...... 62

3.2.2.2 Assay format-2 ...... 63

3.2.3 Exploration of SERS-based magnetic aptasensor for the

ultrasensitive detection of H3 ...... 63

3.2.3.1 Oligonucleotides ...... 63

3.2.3.2 Synthesis of signal probes ...... 64

3.2.3.3 Preparation of capture probes ...... 65

3.2.3.4 Histatin 3 detection ...... 66

3.2.3.5 SERS measurement ...... 66

3.2.4 Demonstration of an aptamer-based lateral flow test strip for on-

site detection of H3 ...... 67

3.2.4.1 Aptamer and other oligonucleotides ...... 67

3.2.4.2 Preparation of AuNPs-aptamer conjugates ...... 67

3.2.4.3 Preparation of test and control zones ...... 68

3.2.4.4 Assembly of the lateral flow test strip ...... 69

3.2.4.5 Detection procedure ...... 70

3.3 Investigation of the use and challenges associated with saliva-based

measurement ...... 70

3.3.1 Preparation of artificial saliva samples ...... 71

3.3.2 Application with ELASA ...... 71

3.3.3 Collection and processing of human saliva ...... 72

3.3.4 Application of saliva with AuNPs-based colorimetric assay ...... 72

4 Results and Discussion ...... 73

4.1 Selection and preliminary characterization of structure-switching ssDNA

aptamers for salivary peptide H3 ...... 73

4.1.1 Aptamer selection by immobilizing ssDNA library ...... 74

4.1.1.1 Selection strategies...... 74

4.1.1.2 Real-time PCR ...... 76

4.1.1.3 Double-stranded DNA to ssDNA conversion ...... 77

4.1.1.4 Transformation, cloning, and sequencing ...... 78

4.1.2 Analysis of aptamer sequences, structural folding, and stability .. 84

xii

4.1.3 Demonstration of binding through MB-based affinity capture assay

coupled to SDS-PAGE ...... 88

4.2 Integration and demonstration of the H3-selective aptamers into ultrasensitive assay formats to allow for the identification and quantification in complex media 90

4.2.1 Development of ELASA for the additional investigation into the

binding between the selected aptamer candidates and H3 ...... 90

4.2.1.1 Binding analysis using ELASA ...... 90

4.2.1.2 Binding kinetics studies ...... 92

4.2.1.3 Determination of Kd ...... 93

4.2.2 Development of AuNPs-based colorimetric assays based on the

structural feature of the identified H3 aptamers ...... 94

4.2.2.1 Assay format-1 ...... 94

4.2.2.1.1 Principle of the colorimetric detection ...... 94

4.2.2.1.2 Spectral characteristics...... 96

4.2.2.1.3 Optimization of experimental conditions...... 100

4.2.2.1.4 Calibration modeling for H3 ...... 106

4.2.2.1.5 Selectivity ...... 108

4.2.2.2 Assay format-2 ...... 108

4.2.2.2.1 Principle of the colorimetric assay ...... 108

4.2.2.2.2 Calibration modeling for H3 ...... 110

4.2.2.2.3 Selectivity ...... 112

xiii

4.2.3 Exploration of SERS-based magnetic aptasensor for the

ultrasensitive detection of H3 ...... 112

4.2.3.1 Principle of the detection method ...... 112

4.2.3.2 Characterization and optimization of the signal probe ...... 114

4.2.3.3 Characterization of the capture probe ...... 118

4.2.3.4 Optimization of the detection parameters ...... 119

4.2.3.5 Histatin 3 detection ...... 120

4.2.4 Demonstration of an aptamer-based lateral flow test strip for on-

site detection of H3 ...... 124

4.2.4.1 Principle of the aptamer-based lateral flow test strip...... 124

4.2.4.2 Confirmation of aptamer immobilization on AuNPs ...... 125

4.2.4.3 Optimization of the performance of the test and control lines . 127

4.2.4.4 Identification of appropriate binding conditions between AuNPs-

aptamer and H3 ...... 136

4.2.4.5 Detection of H3 ...... 137

4.3 Investigation of the use and challenges associated with saliva-based measurements ...... 139

4.3.1 Analysis of H3 doped artificial saliva samples with ELASA ..... 139

4.3.2 Implementation of human saliva with AuNPs-based colorimetric

assay format-2 ...... 141

4.3.2.1 Selectivity ...... 141

4.3.2.2 Application with human saliva and artificial saliva ...... 143

xiv

5 Conclusions and Recommendations ...... 151

5.1 Conclusions of this research project ...... 151

5.2 Future research recommendations ...... 155

5.2.1 Aptamer selection and modifications to improve affinity ...... 155

5.2.2 Gold nanoparticles-based colorimetric assay ...... 156

5.2.3 Improvement on SERS aptasensor...... 156

5.2.4 Lateral flow test strip ...... 158

5.2.5 Continuous, real-time measurements of salivary biomarkers ..... 159

References ...... 162

Appendix A Experimental protocol for library immobilization SELEX ...... 196

Appendix B Agarose gel electrophoresis protocol for PCR end product verification .... 203

Appendix C Running SDS-PAGE ...... 206

List of Tables

Table 2.1: Antibodies vs. aptamers ...... 13

Table 3.1: Oligonucleotides used for the SELEX process...... 51

Table 3.2: Biotinylated aptamers used in ELASA...... 59

Table 3.3: Oligonucleotides used for the lateral flow test strip development ...... 67

Table 4.1: Selection rounds, respective concentrations of H3 and H8, and capture probes

used in the SELEX process...... 76

Table 4.2: Aptamer sequences obtained from 14 rounds of SELEX for H3. The primer

binding regions are underlined...... 86

Table 4.3: Gibbs free energy change, G-score, and TM of the aptamers...... 88

Table 4.4: Analytical results for H3 in human saliva samples ...... 150

xvi

List of Figures

Figure 2-1: Schematic illustration of the aptamer-target recognition ...... 11

Figure 2-2: Chemical structure of caffeine and theophylline ...... 12

Figure 2-3: Schematic diagram of an ssDNA library sequence...... 16

Figure 2-4: Schematic of SELEX process ...... 18

Figure 2-5: Schematic representation of different ELASA formats...... 34

Figure 2-6 : Schematic illustrating the oscillations of surface plasmons in a spherical

metal nanoparticle ...... 36

Figure 2-7: Energy-level diagram displaying the states involved in Rayleigh and Raman

scattering...... 40

Figure 2-8: The EM field distribution of a AuNP and “hot spot” between two aggregated

AuNPs ...... 42

Figure 2-9: Typical configuration of the LFA strip ...... 48

Figure 3-1: Schematic diagram of SELEX for H3 aptamer selection ...... 54

Figure 3-2: Magnetic bead-based affinity capture coupled to SDS-PAGE for visual

monitoring of binding...... 58

Figure 3-3: Schematic illustration of direct format ELASA...... 61

Figure 3-4: Synthesis of signal probe molecules...... 65

Figure 3-5: Schematic illustration of the capture probe...... 65

xvii

Figure 3-6: Formation of the test and control lines on the nitrocellulose membrane...... 69

Figure 3-7: Top and side views of the assembled lateral flow test strip before AuNPs-

aptamer conjugate loading...... 70

Figure 4-1: Real-time PCR curves for the samples having sufficient and insufficient

ssDNA sequences for amplification and PCR product verification in agarose

gel electrophoresis ...... 77

Figure 4-2: Agarose gel electrophoresis of the asymmetric PCR product...... 78

Figure 4-3: Schematic illustration of the transformation and cloning process...... 80

Figure 4-4: The map of pCRTM 2.1 TOPO® vector displaying the sequences

surrounding the TOPO® Cloning site ...... 81

Figure 4-5: A portion of a petri dish containing the transformed bacterial colonies with

blue-white screening ...... 82

Figure 4-6: Typical sequencing result from Functional Biosciences with the potential

aptamer inserted into the TOPO® vector ...... 83

Figure 4-7: Sequencing result with the reverse insertion of the aptamer sequence into the

TOPO® vector...... 84

Figure 4-8: Predicted secondary structures of aptamer using mFold ...... 87

Figure 4-9: The SDS-PAGE image showing a size ladder, the control sample, and eluents

from the H3 and H8 incubated aptamer modified MBs...... 89

Figure 4-10: Relative binding affinity and cross-reactivity of the aptamers ...... 91

Figure 4-11: Binding kinetics of LI-H3-APT-3 aptamer to H3 ...... 93

Figure 4-12: Equilibrium dissociation curve for the aptamer LI-H3-APT-3 ...... 94

xviii

Figure 4-13: Schematic illustration of the colorimetric assay format-1 using H3 binding

aptamer and AuNPs ...... 96

Figure 4-14: Ultraviolet-Visible absorption spectra of AuNPs, AuNPs with NaCl, AuNPs

containing aptamer and NaCl, and AuNPs with aptamer, H3, and NaCl .... 98

Figure 4-15: Transmission electron microscopy images of citrate-stabilized AuNPs,

AuNPs in the presence of NaCl, aptamer adsorbed AuNPs in NaCl, and

aptamer adsorbed AuNPs in the presence of H3 and NaCl ...... 99

Figure 4-16: Effect of NaCl concentration on the aggregation of the citrate-stabilized

AuNPs ...... 101

Figure 4-17: Effect of the aptamer concentration on the absorbance ratio of AuNPs .... 103

Figure 4-18: Incubation study to optimize the time required for aptamers to stabilize the

AuNPs ...... 105

Figure 4-19: Effect of H3 incubation time on the absorbance ratio ...... 106

Figure 4-20: Detection of H3 using AuNPs-based colorimetric assay format-1 ...... 107

Figure 4-21: Selectivity of the assay format-1 against H8 ...... 108

Figure 4-22: Schematic illustration of the colorimetric assay format-2 ...... 110

Figure 4-23: Colorimetric detection of H3 using AuNPs-based assay format-2 ...... 111

Figure 4-24: Selectivity of the assay format-2 against H8 ...... 112

Figure 4-25: Schematic illustration of the SERS-based magnetic aptasensor for the

detection of H3 ...... 113

Figure 4-26: Raman spectrum of 4-NTP powder, and SERS spectrum of 2 µM 4-NTP

conjugated AuNPs, and Raman spectra of AuNPs and 2 µM 4-NTP

dissolved in ethanol showing no detectable signal...... 115

xix

Figure 4-27: Surface-enhanced Raman scattering intensity of AuNPs-NTP conjugate at

various concentrations of 4-NTP ...... 117

Figure 4-28: Aptamer immobilization on the AuNPs-NTP conjugates functionalized with

various concentrations of 4-NTP ...... 117

Figure 4-29: Verification of aptamer immobilization on AuNPs-NTP conjugates ...... 118

Figure 4-30: Absorbance of cDNA solution before and after immobilization on MBs at

260 nm...... 119

Figure 4-31: Optimization of signal probes for 2 µl of 10 mg/ml capture probe ...... 120

Figure 4-32: Dried samples on the glass slide and 50x magnified portion with SERS

measurement spots of the sample...... 121

Figure 4-33: Response of the SERS-based magnetic aptasensor to H3 samples ...... 122

Figure 4-34: Schematic representation of the aptamer-based lateral flow test strip for the

detection of H3...... 125

Figure 4-35: Ultraviolet-Visible spectra of citrate-stabilized AuNPs and aptamer-

immobilized AuNPs with and without 240 mM NaCl ...... 127

Figure 4-36: Capture of AuNPs-aptamer complex by Probe-TL-1, Probe-TL-2, and

Probe-CL sequences at room temperature ...... 129

Figure 4-37: The most stable secondary structures of the aptamer, Probe-TL-1, and

Probe-TL-2 at 137 mM [Na+] and 5 mM [Mg2+] at room temperature. .... 131

Figure 4-38: Capture of AuNPs-aptamer complex by Probe-TL-1, Probe-TL-2, and

Probe-CL sequences at 70 °C ...... 133

Figure 4-39: The most stable secondary structures of the aptamer, Probe-TL-1, and

Probe-TL-2 at 137 mM [Na+] and 5 mM [Mg2+] at 70 °C...... 134

Figure 4-40: Test strips with 2.5 and 5 µl of AuNPs-aptamer after blocking conjugate pad

with 10 mM PBS containing 1% ovalbumin, 0.25% Tween 20, 2% sucrose,

and 0.2% sodium azide...... 136

Figure 4-41: Identification of appropriate binding conditions between the AuNPs-aptamer

and the target molecule H3 ...... 137

Figure 4-42: Response of lateral flow test strip for 0 (control) and 20 µg/ml H3 prepared

in the conjugate pad blocking buffer at room temperature...... 138

Figure 4-43: The binding performance of the aptamer LI-H3-APT-3 to the H3 doped in

artificial saliva samples ...... 141

Figure 4-44: Selectivity of the AuNPs-based assay format-2 against various components

of saliva ...... 142

Figure 4-45: Absorbance ratios of the blank and H3 (10 µg/ml) doped human saliva,

artificial saliva, and buffer samples...... 143

Figure 4-46: Response to externally added H3 (10 µg/ml) in 1 mg/ml mucin and 10

mg/ml CMC...... 145

Figure 4-47: Interaction of mucin, CMC, and aptamer with citrate-stabilized AuNPs. . 146

Figure 4-48: Response to 10 µg/ml H3 spiked in various concentration of mucin and

CMC samples...... 147

Figure 4-49: Response to H3 doped artificial saliva sample without CMC...... 148

Figure 4-50: Matrix effect of human saliva in the assay ...... 149

Figure 5-1: Proposed improvement to the SERS based magnetic assay ...... 158

xxi

List of Abbreviation

4-ATP ...... 4-aminothiophenol 4-MBA ...... 4-Mercaptobenzoic Acid 4-MPY...... 4-mercaptopyridine 4-NTP ...... 4-Nitrothiophenol

AIDS ...... Acquired Immunodeficiency Syndrome AuNPs ...... Gold Nanoparticles

B&W ...... Binding and Washing BB ...... Binding Buffer BSA ...... Bovine Serum Albumin cDNA ...... Complementary Deoxyribonucleic Acid CE ...... Chemical Enhancement CMC ...... Sodium Carboxymethyl Cellulose

DL ...... Detection Limit DNA ...... Deoxyribonucleic Acid DTNB ...... 5,5-Dithiobis-2-Nitrobenzoicacid

EDTA ...... Ethylenediaminetetraacetic Acid ELASA ...... Enzyme-Linked Aptamer Sorbent Assay EM...... Electromagnetic

FDA...... Food and Drug Administration

HEPES ...... 4-(2-Hydroxyethyl)-1-Piperazineethanesulfonic Acid Histatin 3 ...... H3 Histatin 8 ...... H8 HIV ...... Human Immunodeficiency Virus HPLC ...... High Performance Liquid Chromatography

IB...... Immobilization Buffer kDa ...... Killodaltan LFA ...... Lateral Flow Assay

xxii

MALDI-MS ...... Matrix-Assisted Laser Desorption/Ionization-Mass Spectrometry MB ...... Magnetic Bead MGITC ...... Malachite Green Isothiocyanate

PBS ...... Phosphate Buffered Saline PCR ...... Polymerase Chain Reaction pM ...... Picomolar PMF...... Protein Mass Fingerprinting POC ...... Point-of-Care

QGRS ...... Quadruplex Forming G-Rich Sequences

R6G ...... Rhodamine 6 G RNA ...... Ribonucleic Acid

SDS-PAGE ...... Sodium Dodecyl Sulfate–Polyacrylamide Gel Electrophoresis SELEX ...... Systematic Evolution of Ligands by Exponential enrichment SERS ...... Surface-Enhanced Raman Scattering S-HRP ...... Streptavidin- Horseradish Peroxidase SPR ...... Surface Plasmon Resonance ssDNA ...... Single-Stranded Deoxyribonucleic Acid

TCEP ...... Tris(2-carboxyethyl)phosphine

UV-Vis ...... Ultraviolet-Visible

WHO ...... World Health Organization

xxiii

Chapter 1

1 Introduction

1.1 Background and significance

Biomarkers are measurable variables that can indicate normal biological processes, diseased state, and physiological response to a therapeutic intervention [1]. The ability to rapidly and accurately monitor biomarkers could provide significant advantages to research and clinical applications, such as disease diagnosis, drug development, treatment monitoring, and identifying the physiological state [2, 3]. A variety of biomolecules have been validated as the indicator of different biological conditions, and numerous investigations are ongoing in search of novel biomarkers. Usually, the biosample assessed for identifying or detecting biomarkers is blood, which is considered the gold standard [4].

However, the invasive method of blood sampling is painful, uncomfortable, involves the risk of infection, requires trained personnel, and limits the sampling interval [5].

Alternatively, in recent years, saliva has garnered popularity in search of novel and useful biomarkers due to several advantages compared to other body fluids. For instance, the collection of saliva is easy, non-invasive, inexpensive, and does not require trained personnel. Unlike the blood specimen, saliva does not clot and is easier to handle. For the patients who need frequent sampling, the non-invasive sample collection procedure

1 reduces their anxiety and discomfort. Also, the risk of infection to patients and the exposure of healthcare professionals to blood-borne pathogens is minimal [6].

With technological advances in detection and quantification methods, once merely a digestive juice, saliva is now being considered a mirror of an individual’s health status

[7]. It is a complex biological fluid containing a variety of proteins, enzymes, hormones, nucleic acids, cytokines, antibodies, peptides, polynucleotides, electrolytes, and other components [8-10]. In humans, saliva is secreted from 3 pairs of the major (parotid, sublingual, and submandibular) and numerous minor salivary glands (labial, buccal, lingual, and palatal) [6, 11]. The blood-derived components enter saliva through the transcellular or paracellular routes and influence the molecular constituency [11]; thereby, making saliva a source of biomarkers not just for oral diseases but also for systemic disorders [12]. Furthermore, the salivary glands are innervated by the parasympathetic and sympathetic branches of the autonomic nervous system that controls the salivary secretions according to the physiological conditions [11]. Over the past few years, saliva has been used to detect growing number of diseases including periodontal diseases [13-15], oral cancer [16-18], dental caries [19] and systemic disorders, such as hepatitis [20], human immunodeficiency virus (HIV) [21], diabetes [22-24], breast cancer [25, 26], cardiovascular disease [27, 28], renal diseases [29] , and stress-related disorders [30-35].

By considering the advantages of the saliva over other diagnostic fluids, a collaborative team of researchers at The University of Toledo, Marvin et al., embarked on a project to identify salivary biomarkers that correlate with stress and cognitive performance in human saliva [36]. The study also aimed to develop newer analytical methods for the identified biomarkers. Acute stress results in increased activity of the sympathetic branch of the autonomic nervous system. Exposure to acute stress for a prolonged 2 time may result in chronic stress and fatigue, which might compromise the physical and cognitive performance of an individual [4, 37]. This is of particular interest to professionals in demanding occupations, such as military, healthcare, and law enforcement [4]. Real- time monitoring of stress biomarkers could prove beneficial to obtain optimal performance and prevent damage that may result from the impaired ability.

To identify the novel stress biomarkers, Marvin et al. analyzed the saliva samples collected from the medical residents that were placed into a brief stressed condition by performing emergency medicine simulations. The comparison was made between the saliva collected before simulation, after simulation, and the next morning upon waking.

Following the collection, the samples were analyzed via sodium dodecyl sulfate- polyacrylamide gel electrophoresis (SDS-PAGE), mass spectrometry, and densitometry.

The results revealed increased levels of salivary alpha-amylase, cystatins, a~26 kDa band, and a low-molecular-weight protein (<10 kDa) band. The low-molecular-weight band was identified as containing histatin 3 (H3) by various methods, such as peptide mass fingerprinting (PMF), matrix-assisted laser desorption/ionization-mass spectrometry

(MALDI-MS), and SDS-PAGE. Based on those results, they postulated that the change in

H3 concentration after the simulation studies could be a possible indicator of acute stress

[36].

Histatin 3 is a member of the histatin family, which are low-molecular-weight, cationic, histidine-rich peptides found in the saliva of humans and higher primates. The histatins have been considered the vital components of the non-immune oral host defense system due to their antimicrobial and antifungal properties [38-40]. More specifically, H3 has demonstrated antifungal activity against Candida albicans, which is the most common

3 causative agent of oral candidiasis [41]. The oral candidiasis is commonly associated with

HIV-infection, and it occurs in 62–75% of the human immunodeficiency virus (HIV)- infected patients during the disease progression to acquired immunodeficiency syndrome

(AIDS) [42]. In such patients, the concentration of histatins has been reported significantly lower compared to healthy adults [43]. To prevent further complications of oral candidiasis, the patients need early and effective treatment, which might otherwise lead to oropharyngeal and esophageal candidiasis [44, 45]. The most commonly used therapeutic agents for the treatment are antifungal drugs such as fluconazole or itraconazole. Due to the excessive use of these drugs, there has been an increased incidence of azole-resistance species of Candida, mandating the need for new therapeutic agents [46, 47]. The H3 has antifungal properties and a different mechanism of antifungal activity than the azole-based antifungal drugs; therefore, it has the potential to become an alternative therapeutic agent for oral candidiasis. Overall, the measurement of H3 levels in saliva could provide special insight into human health conditions, including stress, HIV-infection, and associated oral candidiasis. Also, H3 detection might be an essential aspect of therapeutic drug monitoring if H3 can be established as an alternative therapeutic agent.

Several techniques for the detection of H3 exist, including gel chromatography, high-performance liquid chromatography (HPLC), mass spectrometry, and antibody-based immunoassays [36, 40]. These methods, however, have some issues, and few of them were mentioned by Marvin et al. in their study. In an effort to further confirm the presence of

H3 in the low-molecular-weight band, Marvin et al. experienced difficulties with the antibody-based analytical method. In western blot experiments, neither monoclonal (4G9,

Novus Biologicals, Centennial, CO) nor the polyclonal (H-40, Santa Cruz Biotechnology,

Dallas, TX) H3 antibodies detected salivary or the synthetic H3. Marvin et al. pointed out certain issues they had with antibodies, which included the poor binding to H3, limited commercial availability, and lack of adequate validation [36]. The antibodies, in general, are sensitive to temperature and have a short lifespan. Also, the chromatographic methods available for H3 detection are laborious, utilize expensive instrumentation, and time- consuming. Due to the problems and complexities of the currently available H3 detection methods, there is a clear need for an alternative technique. Aptamer-based assays and sensing platforms have the potential to be viable alternatives in such applications.

Aptamers are synthetic oligonucleotides that can selectively bind to a specific target molecule with high affinity [48, 49]. These affinity ligands are functionally similar to the antibodies; however, the aptamers offer several advantages over antibodies. Aptamers are inexpensive to produce and can be developed for almost any target by in vitro iterative process known as Systematic Evolution of Ligands by Exponential Enrichment (SELEX)

[50]. The discovery time is short, and, once identified, aptamers can be synthesized chemically with minimal batch-to-batch variability. The aptamers are stable at extreme heat and pH conditions, and their denaturation is reversible [51-53]. Also, the SELEX process offers flexibility to manipulate the selection parameters, such as temperature, pH, and ionic strength, to match the final application conditions [54]. Moreover, the sequences that may bind non-specifically to other molecules can be removed by the incorporation of negative or counter-selection steps in the selection process [55]. More importantly, the aptamer-based diagnostic assays have demonstrated the potential to fulfill the World

Health Organization's (WHO’s) “ASSURED” criteria for point-of-care (POC) applications. The “ASSURED” stands for: affordable, sensitive, specific, user-friendly,

5 robust, equipment-free, and deliverable to end-users [56]. Despite these benefits, only a few aptamer-based technologies are commercially available today. Since their discovery in 1990, only one aptamer based therapeutic product (Macugen/Pegaptanib sodium) has

FDA clearance for clinical use. For the diagnostic application, a few commercial products, such as OTA-sense, AflaSense, AptoCyto, AptoPrep, SOMAscan, OLIGOBIND, and Taq

DNA polymerase, are available [57]. The sluggish progress in aptamer technology can be contributed to the following issues: Even though the SELEX process is less time- consuming and cost-effective than antibody production, there is no universal method applicable to each target, and the process is still tedious and complicated. The success rate of aptamer selection is also low [58]. The big companies in the biotech industry are still investing their financial resources in the well-established antibody-based technologies

[57]. Also, the research labs are picking the already validated aptamers, such as thrombin aptamers, for the demonstration of novel platforms rather than selecting aptamers for novel biomarkers. The aptamer community has described this issue as the “thrombin problem.”

[59]. All these factors are severely impeding the development and application of aptamers.

Another aspect of this project is the implementation of the developed aptamers for the sensing application in saliva. As mentioned above, saliva is a promising biofluid offering several advantages over other biosamples. The use of saliva, however, is limited due to the methodological and technological obstacles associated with it. Many informative biomarkers generally are present in the lower concentration than in serum. The saliva samples need processing such as freeze-thawing and centrifugation to eliminate the matrix effect arising due to food particles and the mucin. Also, there is currently no standardized technique for sample collection, which is the potential source of the variability in the saliva-

6 based measurements [11]. Further, the circadian variation of the analytes needs to be understood before relying on the results [60]. Due to these limitations, salivary diagnostics are limited to the centralized laboratory and requires time-consuming assays, expensive instruments, and a large volume of samples. The overall cost associated with these methods due to testing supplies, transportation, storage, and labor makes these tests expensive [61].

1.2 Research objectives

The objective of the proposed research is to demonstrate the feasibility of a widely applicable aptamer selection method for novel biomarkers and the implementation of the identified aptamers with different platforms for the detection application in human saliva.

To accomplish this key objective, a core set of three specific aims were defined:

1. Selection and preliminary characterization of structure-switching ssDNA aptamers for

salivary peptide H3.

A variant of the SELEX method in which the library sequences are immobilized for the purpose of aptamer selection was implemented to identify the binding ligands that transform structural conformation upon binding to the target molecule H3. During the

SELEX method, the selection conditions were tuned, and counter-selection steps were incorporated to generate aptamers with higher affinity and selectivity. Following the aptamer selection, the identified aptamers were first analyzed by two different computational tools, mFOLD and quadruplex forming G-rich sequences (QGRS) mapper, to predict the structural conformations and stability. Based on these analyses, the most promising candidate was recognized and evaluated to visually confirm the binding by combining magnetic-bead based affinity capture and SDS-PAGE. This task gave us an

7 indication of whether the aptamer selection process was successful or not before spending financial resources for larger-scale experiments for characterizations.

2. Integration and demonstration of the H3-selective aptamers into ultrasensitive assay

formats to allow for the identification and quantification of H3 in complex media.

The research activities performed in specific aim 1 ascertained the successful aptamer selection process. Within this specific aim, the selected aptamers were further characterized and implemented on various assay platforms for detection applications.

Firstly, a more detailed study on binding affinity and specificity was performed by developing enzyme-linked aptamer sorbent assay (ELASA). Various candidates were compared, and the best performing candidate was identified for further application with other assay platforms. Next, by employing the structural feature of the aptamer, two different formats of the gold nanoparticles (AuNPs)-based colorimetric assays were developed. Numerous parameters were optimized, and the assay time was reduced to 12 minutes. These colorimetric assays are facile and do not require any modifications, washing, and pretreatment steps and even offer visual detection. Afterward, a surface- enhanced Raman scattering (SERS)-based magnetic aptasenor was investigated in order to address the unmet need of the highly sensitive assay to detect the deficient levels of biomarkers in human saliva. Finally, to address the necessity for an alternative tool for saliva processing that can replace the current laboratory-based methods and offer point-of- care testing, a lateral flow test strip was demonstrated.

3. Investigate the use and challenges associated with saliva-based measurements

Most of the sensing platforms or the assays work well in the buffer medium; however, when the biological fluids are applied, these assays fail to detect the required

8 target molecule due to various reasons. In this specific aim, the application of human saliva and artificial saliva samples with the developed assay platforms have been explored.

Chapter 2

2 Literature Review

2.1 Aptamers

Aptamers are nucleic acid ligands that bind to various target molecules with a high affinity and specificity due to their capability of folding into specific three-dimensional complex structures [48, 49]. Three-dimensional shapes of aptamers are characterized by stems, loops, bulges, hairpins, pseudoknots, triplexes, quadruplexes, and their combinations [55]. These conformations allow aptamers to interact uniquely with a wide variety of targets through intermolecular interactions like electrostatic forces between charged groups, hydrogen bonds, stacking of aromatic rings, and the complementary three- dimensional shape or a combination of these effects [55, 62]. An example of aptamer- target interaction mechanism is shown in Figure 2-1.

Figure 2-1: Schematic illustration of the aptamer-target recognition. Reproduced with permission from reference [55].

For the first time in 1990, Tuerk and Gold reported the selection of RNA molecules having binding affinity to T4 DNA polymerase, gp43, and named the selection method

SELEX [50]. At the same time, Ellington and Szostak also described the SELEX method for identifying RNA molecules binding to the organic dyes like Cibacron Blue and

Reactive Blue 4. The Szostak lab termed these binding ligands as “aptamers,” referring to the Latin word “aptus”- which means fitting and the Greek word “meros,” meaning particle

[63]. Two years later, Ellington and Szostak demonstrated the selection of high-affinity aptamers by using the ssDNA library [64], which later became a preferred option over

RNA library because of DNA’s greater inherent stability and easy amplification step.

However, due to the greater diversity in 3D structures of RNA sequences, RNA aptamers are still being selected [65]. Since their discovery in 1990, aptamers have gained increasing attention and consequently have been selected for a wide array of targets ranging from ions

[66, 67], small molecules [68], peptides [69, 70], proteins [71, 72], toxins [73], vitamins

[74, 75], to even whole cells [76, 77], viruses [78-80], or microorganisms [81].

Aptamers are considered as an excellent alternative or substitutes to antibodies due to various advantages over antibodies. In terms of binding affinity, these binding ligands are comparable with monoclonal antibodies and sometimes even better, having the

11 dissociation constants in the range of sub-pM to µM range. They are highly specific and can differentiate the molecules with subtle structural differences such as the molecules having the same structure but different functional groups. An example would be the aptamer against theophylline developed by Jenison et al., which had 10,000 times higher affinity to theophylline than for caffeine [82]. As shown in Figure 2-2, the structure of caffeine and theophylline differs by only a single functional group (methyl) at nitrogen atom N-7. In addition, aptamers offer several other advantages over antibodies, such as a cost-effective method of production via chemical synthesis, shorter discovery time, in vitro method of selection, minimal batch to batch variability, stability in harsh conditions, and reversible denaturation [51-53]. The detailed comparison between the antibodies and aptamer is presented in Table 2.1. Due to these exciting features, aptamers have found a wide range of applications in various areas including therapeutics, diagnostics, drug delivery, bio-imaging, purification and separation, biomarker discovery, food inspection, and environment monitoring [53].

Figure 2-2: Chemical structure of caffeine and theophylline. The difference between these two molecules is highlighted in red circle.

Table 2.1: Antibodies vs. aptamers

Antibodies Aptamers Production Production  Requires use of animals  Produced in vitro (no animals  Expensive reactors required used)  Takes several months  Selection can be performed under  Poor batch-to-batch reproducibility a variety of conditions to match  Viral or bacterial contaminations the final application platform possible  Takes several weeks  Low batch-to-batch variability  Viral and bacterial contamination not problematic Targets Targets  Must provoke a strong immune  A wide range target molecule response including non-immunogenic and  Must be non-toxic toxic molecules Stability Stability  Irreversible denaturation at high  Thermally stable, reversible temperatures denaturation  Sensitive to pH  Tolerant to pH changes

Modifications Modifications  Limited modifications  Wide variety of chemical modifications for diverse applications Shelf-life Shelf-life  Limited (6 months)  Long (several years if frozen)  Single use  Can be reused in some applications Immunogenicity Immunogenicity  Can cause immune response  No intrinsic immune response

Target sites Target sites  Determined by animal immune  Can be determined by investigator system Binding affinity Binding affinity  Usually ranges from nanomolar to  Usually ranges in low nanomolar picomolar to picomolar range

Downstream applications Downstream applications  Will work only under near  Functional under a broader range physiological conditions of conditions

2.2 Generation of aptamers by SELEX

Aptamers for a target are selected by an iterative in vitro process of 8 to 15 rounds known as SELEX. Subsequent selection and amplification of the target-binding sequences from the combinatorial oligonucleotide library lead to the generation of highly specific aptamers. The entire process can be divided into following four major steps: (I) library generation, (II) selection (binding, separation, and elution of target-bound sequences), (III) amplification and strand separation, and (IV) cloning and sequencing.

2.2.1 Aptamer library

The first task for starting SELEX is to design the randomized oligonucleotide library, which is composed of approximately 1013-1015 different single-stranded sequences.

Each molecule (70-120 bases length) in a library is a unique sequence consisting of a central randomized region of 20-80 bases flanked by the defined primer binding sites on both ends (17-21 bases) (Figure 2-3). The fixed regions on both sides of random sequence are required for the amplification of the strands via PCR [83].

Various important aspects, such as the length of the random region, size of the target molecule, and the degree of randomization should be considered while designing a

SELEX library. The diversity of the library is dependent on the length of the random region, and the number of individual/unique strands can be estimated by the formula 4n, where 4 stands for nucleotides A, T, G, and C and n is the length of the random region. In general, the length of the random region can range from 20-80 nucleotides for SELEX.

Mostly after selection, the aptamers are truncated down to a minimal functional sequence.

The study by Bock et al. has reported the preserved functionality of the thrombin aptamer just within 15 nucleotide sequence following the truncation of 96-mer aptamer [84],

14 suggesting that the short randomized regions are enough for strong binding. Also, short library sequences are easier to analyze and cost-effective [55]. However, the longer random region provides the libraries with greater structural complexity and may offer opportunities for the identification of higher affinity aptamers [85]. Nevertheless, care should be taken as very large random regions can interact with the flanking primer binding sites and impair the PCR amplification. Additionally, the structural complexity of the library can also be enhanced by inserting the modified bases in the library sequences. Modified bases introduce new functional groups providing new possibilities for the interaction and might also enhance stability against nuclease degradation [86].

The next parameter that should be considered while designing the library is the size of the target molecule. To increase the chance of binding between the library sequences and target, a random region with comparable molecular weight to the target is recommended. Furthermore, two different types of library sequences, having either a completely random or partially doped central region, can be considered based on the availability of the existed aptamers. The entirely random library is the first choice if a target molecule is new, and no aptamer existed before. However, if a target molecule has already known aptamers, then, a partially doped library by keeping some known motifs responsible for binding can be implemented to re-select aptamers with improved binding characteristics

[87]. In addition to these factors, the primers should be designed carefully to eliminate the possibility of forming self-dimers and heterodimers [87, 88].

Figure 2-3: Schematic diagram of an ssDNA library sequence.

2.2.2 Selection

The selection step of the SELEX procedure can be divided into three different steps; binding of the target molecules with the oligonucleotide library, partitioning of the unbound sequences, and elution of the target-bound oligonucleotides. The aptamer selection process is illustrated in Figure 2-4. For the first step of the SELEX cycle, the randomized library is incubated with the target molecule under the desired set of environmental conditions. Various parameters, such as the buffering system, pH, and temperature can be tuned to match the final application conditions [89]. Careful consideration should be taken while choosing buffering conditions. Since DNA is a polyanionic molecule, the negative–negative charge repulsion may inhibit the formation of complex structures, hindering the binding with the target. The use of counter ion such as

Na+ in the binding buffer can neutralize negative charges of the DNA backbone and enhance the formation of complex structures. Also, the addition of a small amount of divalent cation Mg2+ is recommended as it blocks the negative charge repulsion and might enhance the binding between the target and aptamers [90].

During the incubation, some of the sequences in the library bind to the target, forming aptamer-target complexes. The unbound sequences are removed, and the target- bound sequences are eluted, which are then amplified by PCR. The amplified sequences

16 are strand-separated to generate a new ssDNA library for the subsequent SELEX cycle. By iterative rounds of selection and amplification under the desired conditions, the initial random oligonucleotide pool is reduced to relatively few sequence motifs with the highest affinity and specificity to the target. Following the final round of selection, the aptamer sequences are cloned and sequenced. The rounds of SELEX are typically repeated multiple times in order to obtain the enriched pool of the high-affinity binders. Depending on the type of the target molecule, implementation of counter-selection steps, and stringency of the selection conditions, the selection process typically repeated for 8-15 cycles [48].

Figure 2-4: Schematic of SELEX process, including negative-selection and counter- selection steps. The first step of SELEX is the incubation of the random library with the target molecules. After letting the binding happen, the unbound sequences are separated from aptamer-target complexes and discarded. The sequences bound to the target are then eluted and amplified by PCR. The PCR product, which is dsDNA, is converted into ssDNA before applying it to the subsequent round of selection. For negative-selection, the library is incubated with the support materials and the sequences binding to the support material are discarded. The unbound library sequences are recovered and applied for aptamer selection. In order to perform counter- selection, the eluted sequences are incubated with the potential interfering molecule, and the binding sequences are discarded whereas the nonbinding candidates are recovered for further amplification.

The efficient partitioning of target binding sequences from unbound or weakly bound sequences is a crucial step for obtaining highly selective and high-affinity aptamers.

Most commonly for protein targets, such separation is achieved by utilizing nitrocellulose membrane filtration; in which the ssDNA or RNA library is incubated with the protein target and the mixture is filtered through the nitrocellulose film. The proteins with bound oligonucleotides adsorb onto the membrane surface and retained while the unbound molecules are washed off [50, 91, 92]. Even though aptamers for several target molecules have been identified by using nitrocellulose membrane, the method is limited to the large size proteins as it cannot retain small molecules and peptides [91]. Alternatively, electrophoresis methods such as gel and capillary electrophoresis have been successfully employed to separate the aptamer-target complexes from unbound sequences. These techniques separate ionic species based on their charge, frictional forces, and hydrodynamic radius under the influence of applied potential [53]. As the mixture of target and library sequences pass through the gel or capillary, the target-aptamer complexes demonstrate the mobility shift compared to the free target and can be retained. One major advantage of these methods is that the aptamer selection can be achieved in very few rounds, generally 2-4, compared to other methods. Using electrophoresis-based techniques, aptamers for neuropeptide Y [93] and Immunoglobulin E [94] were obtained after only four rounds of selection. However, these methods also fail when the target is a small as the mobility shift between the aptamer-target complex and the target is relatively small and difficult to separate.

Another widely used portioning method is the target immobilization on solid support. For selecting aptamers, the target of molecule is immobilized on a solid support,

19 such as magnetic particles or the other column materials including, sepharose and agarose beads. Following the target fixation, the library sequences are reacted, and the nonbinding sequences are discarded by washing the support materials several times. The higher affinity sequences bound to the target are then retained for further processing [95, 96]. A major limitation of these methods is the need to immobilize the target. The target immobilization can hide the binding pocket of the small molecules. Also, immobilization is not applicable to the molecules without a suitable functional group for immobilization. For such targets, fixation needs an addition of a proper linker, which may sometimes change the native conformation. The structural change might lead to the selection of the aptamers that might not possess affinity and or selectivity for the target in its native form [97, 98]. This problem can be solved by an alternative approach in which the ssDNA library sequences can be immobilized on a solid support via complementary capture probes instead of the target molecule. Upon binding to the target, the ssDNA sequences undergo a conformational change and get released from the support. These released sequences are then collected and utilized for a further round of SELEX. The aptamers selected by this mechanism are referred to as structure-switching aptamers [99, 100]. This technique eliminates the need for target fixation, and aptamers can be selected for the target in its unmodified form.

Besides, the structure-switching upon target binding is a highly desired attribute of the aptamers for the biosensing application. The structure-switching can ensure high detection specificity as the aptamers switch structural conformation only when it interacts with the specific target molecule. Also, the structure-switching feature is compatible with a variety of signal transduction mechanisms for a signal generation [101].

Overall, for partitioning, the target molecule is either remains in a free state or immobilized on the solid support. Apart from the techniques mentioned above, several additional methods, such as flow cytometry [102], affinity tags [103], surface plasmon resonance (SPR) [104], or centrifugation [105] have been reported for aptamer selection.

While using solid support for aptamer selection, it is possible that the sequences binding to the support material (e.g., affinity chromatography column, magnetic beads, nitrocellulose filters, reaction tubes, etc.) may also evolve during the selection process. In order to avoid such nonspecific binders, a negative-selection (Figure 2-4) step comprising pre-incubation of library sequences with the support material without the target can be performed. The sequences binding to the support material are discarded, and the non- binders are implemented for the subsequent selection process [64]. Similarly, to avoid cross-reactivity and select highly specific aptamers, the counter-selection steps may be carried out against the potentially interfering molecules [106]. As depicted in Figure 2-4, the enriched pool is incubated with the interfering molecule and the sequences interacting with the interfering molecules are excluded, while recovering the unbound sequences. It is possible to implement multiple counter-selection steps within a SELEX method to enable a more specific selection of aptamer candidates for the desired target.

After partitioning the unbound oligonucleotides, the next step in the SELEX cycle is to elute the bound sequences from aptamer-target complexes. As the interaction between the target and binding sequences is non-covalent, relatively mild conditions can be used to separate two species. Several methods have been reported for the aptamer elution including, heat treatment, change in ionic strength or the pH, and use of denaturing substances, such as SDS, ethylenediaminetetraacetic acid (EDTA), and urea [107-110].

Additionally, some researchers have carried out affinity elution by using pure target [111,

112] or competitive binders [113]. The concern with the affinity elution is the cost associated with obtaining the pure target and the solubility of the target molecule, which might not usually be feasible to prepare at the concentration required for competitive elution. It is also worth mentioning that the complete elution of the sequences may not be achieved as very high-affinity sequences are often difficult to elute and therefore are lost during elution. Besides, it is also possible to process aptamer-target complexes without separating, if the target molecules do not interfere with the PCR amplification [114].

2.2.3 Amplification and strand separation

The next step in the SELEX process is the amplification of the eluted oligonucleotides. After discarding the non-binding sequences from the library pool, only a small population of oligonucleotides is retained. In order to enable selective evolution, the retained sequences are amplified via PCR to make millions of copies. The amplification is the only step that is different for ssDNA or RNA based SELEX. While using an RNA library, the ssDNA library is first transformed into an RNA by using a sense primer on the

5' end containing the T7 promoter sequence along with the antisense primer. In every round, following the elution, the RNA has to be reverse-transcribed back into DNA via reverse transcription-PCR (RT-PCR). The generated DNA sequences are again transcribed back into RNA with T7 RNA polymerase for the subsequent round of selection. In comparison to RNA, DNA amplification is straight forward and requires only one step of amplification, which makes DNA a more attractive ligand for SELEX [55].

In order to obtain efficient amplification, various parameters, including primer concentration, annealing temperature, and the number of amplification cycles, needs to the

22 optimized. Any mismatch in these parameters will result in the byproduct formation and inefficient PCR amplification, limiting the evolution of the high-affinity aptamers [115].

The optimization experiments by using the traditional amplification systems are often time- consuming as they require product verification via agarose gel electrophoresis after changing each variable. The problem with traditional amplification systems can be solved by using real-time PCR, which uses a fluorescent dye such as SYBR green to track the amount of dsDNA being produced during each PCR cycle in real-time. The tracking of dsDNA during PCR cycles helps to identify the appropriate cycles of PCR required to obtain sufficient amplification and prevent overamplification. Also, the melting curves obtained through real-time PCR can be utilized to predict enrichment progression [116]. In addition to the real-time PCR, some other PCR techniques such as emulsion PCR [117] and high-fidelity digital-PCR [118] have been applied in the SELEX procedure.

Besides the enrichment of target binding sequences, PCR also offers the opportunity for the modification of the oligonucleotides. The primers can be modified with the desired functional groups such as biotin or fluorescence molecules for immobilization or detection, respectively. Biotin modification is commonly used to immobilize dsDNA on the streptavidin beads for the strand separation; whereas, the fluorescence modifications are used for monitoring the selection enrichment [55]. Also, modified bases can be introduced to increase the diversity in the structures of PCR amplicons; thereby, enhancing the aptamer-target interactions.

Following the PCR amplification, the selection process requires the conversion of dsDNA to ssDNA after each round for starting the subsequent SELEX cycle. It is a critical and time-consuming step in SELEX, as the amount and purity of ssDNA can have a

23 significant impact on the success of the aptamer selection [119]. Various methods have been reported for the conversion of dsDNA into ssDNA, such as separation with streptavidin-coated beads [120, 121], lambda exonuclease digestion [122-124], strand separation using urea–polyacrylamide gel [110, 125], and asymmetric PCR [126, 127].

Comparing all of the above methods, asymmetric PCR is the most cost-effective and least time-consuming. In asymmetric PCR, unequal concentrations of forward and reverse primers are used to amplify the desired strand of the template DNA more than the other.

The overall process includes the two phases of amplification known as the exponential amplification phase (produces dsDNA), which is followed by the linear phase that generates ssDNA. Single-stranded DNA generation continues until the reaction is limited by the amount of enzyme present in the reaction mixture [115]. The final product of asymmetric PCR contains the mixture of ssDNA and dsDNA, which can either be purified or can be directly applied to the next round of SELEX.

2.2.4 Cloning and sequencing

After getting enough enrichment of the target-specific oligonucleotides, the last round of SELEX is stopped following the PCR amplification. The PCR products are then cloned into a bacterial vector and sequenced to obtain the individual clones. However, the recent technology of next-generation sequencing (NGS) combined with bioinformatic analysis has replaced the cloning step. Following each round of SELEX, the product can be sequenced with NGS, and through the bioinformatic analysis, the enrichment of the specific binders can be monitored [99]. The SELEX process ends with the cloning and sequencing step.

2.3 Limitations of SELEX

Despite various advancements and optimizations in the selection strategies, the

SELEX process is not free of limitations. The overall success rate of aptamer selection has been reported to be about 50% [128]. Theoretically, SELEX can be performed for any target; however, not all the molecules are suitable for the in vitro selection method. The target molecules should be available in the highest purity and sufficient quantity. Slightly impure targets may produce aptamers with nonspecific binding characteristics. The molecules having large hydrophobic groups and highly negatively charged species are challenging due to electrostatic repulsion with the oligonucleotides. Each target molecule needs different selection conditions; therefore, there is no standardized SELEX protocol.

The protocols depend on the type of target (size, purity, and charge), desired features of the aptamers, and application conditions. As each target requires a unique approach to achieve an appropriate selection environment, it might take weeks to months to complete the aptamer selection process [55, 129].

Another limitation of the SELEX is the chemical diversity of the starting library pool. The diverse library has a higher possibility of containing the aptamers with high affinity and specificity. Increasing the length of the random region might facilitate the crosstalk between the flanking primer binding sites and random region. The modification of the library sequences is another technique to increase structural diversity; however, only the modifications that are compatible with PCR amplification can be used. Also, the price associated with the modification is higher, and the associated analysis is time-consuming.

An additional limiting factor is the loss of potential binders during elution of the target bound sequences from aptamer-target complexes. Also, the PCR may introduce the

25 mutations in the sequences resulting in the loss of some potential binders. The further constraint of the SELEX process is the enrichment of the nonspecific sequences due to the interaction with the tubes or the solid surfaces utilized in the selection process. As already explained in section 2.2.3, additional steps of negative-selections within the SELEX process are required to eliminate the evolution of nonspecific binders. Additionally,

SELEX might require counter-selections with various molecules that share structural similarity with the target. Additional steps, such as counter and negative selections, make the aptamer selection process more complicated, expensive, and time-consuming. It is very important to understand the limitations of the SELEX process, as some limitations can be overcome by modifying the SELEX conditions.

2.4 Aptamer characterization

Once the aptamers are identified by sequencing, the next step is to find the best candidate with the highest affinity and specificity. Before performing experimental studies, the identified sequences can be analyzed in silico by using analytical tools such as Clustal

Omega [130]or the MEME Suite [131] to find the homologous sequences and consensus motifs that are thought to play an essential role in affinity and specificity to the target.

Secondary structure analysis also gives important information about structures of the sequences in given environmental conditions that are relevant for binding. The online server mFold developed by Zuker et al. is used to determine the putative 2D structure [132].

Also, the likelihood of forming G-Quadruplexes (G4) can be predicted by using the online tool QGRS mapper [133].

Following the in silico evaluations, the binding studies are performed to determine the specificity and affinity of the selective aptamer candidates. The 퐾푑 is a reliable measure

26 to represent the binding affinity. A small 퐾푑 value indicates a high affinity to the target. To determine 퐾푑 , several biophysical instruments and techniques, such as SPR, isothermal titration calorimetry (ITC), microscale thermophoresis (MST), backscattering interferometry (BSI), flow cytometry, ELASA, fluorescent and radioactive binding assays, and equilibrium filtration or dialysis, have been reported in the literature [134].

2.5 Applications of aptamers in various assay platforms

As mentioned previously, aptamers possess various advantages over antibodies and are an excellent alternative as the biorecognition element for various diagnostic and analytical applications. In the following sub-sections, an overview of some of the assay approaches that can be realized using aptamers as a biorecognition element is given.

2.5.1 Magnetic bead-based affinity capture coupled to SDS-PAGE

The isolation or the purification of a single protein out of the complex biological media is a challenging task [135]. Traditional purification methods, such as liquid chromatography, centrifugation, filtration, and other equipment, are expensive, require multiple steps, and there is a loss of protein due to the involvement of multiple steps. By using MBs for separation, purification can be reduced to a single step, and the entire step can be performed in a single vial, which results in higher efficiency and reduced risk of protein loss [136].

Magnetic beads are superparamagnetic particles composed of a magnetic core and a polymeric surface coating containing a reactive group that allows their functionalization.

Besides offering functionalization, the polymeric coating also provides the physical and chemical stability to the MBs [137]. As solid support, MBs possess a large surface-to- volume ratio, providing relatively large numbers of binding sites for affinity ligands. The

27 higher number of binding ligands offer higher efficiency of interactions between the ligand and target, resulting in faster assay kinetics [138]. The superparamagnetic property of the

MBs facilitates an easy separation from complex matrices by using an external magnet, eliminating the need for the tedious cleanup process.

In order to provide specificity, the MBs can be modified with affinity ligands, such as antibodies, peptides, and aptamers. As aptamers are superior biorecognition elements compared to its counterparts, the combination of MBs and aptamer for the affinity separation, purification, and analytical application is promising. Various strategies have been utilized to immobilize the aptamers on the MBs. The most commonly used techniques include the conjugation of amine-terminated aptamer with carboxyl functionalized MBs via EDC/NHS chemistry and coupling of biotin functionalized aptamer with streptavidin- coated MBs. Both of these methods provide directional immobilization of the aptamer, which is vital to develop a highly sensitive affinity capture platform [137].

The application of aptamer for affinity purification was first reported by Romig et al. in 1999 for the separation of an L-selectin-immunoglobulin fusion protein form Chinese hamster ovary cell-conditioned medium. In their study, Romig et al. designed the aptamer affinity chromatography column by immobilizing biotinylated aptamers on a streptavidin linked resin. The captured protein was eluted and analyzed via SDS PAGE, protein blot analyses, and ELISA [139]. Over the past few years, simpler separation methods utilizing

DNA aptamer and MBs have been developed for many other proteins, including thyroid transcription factor [140], Thermus aquaticus DNA polymerase [141], His-tagged proteins from E coli. lysates [140], and also for vascular endothelial growth factor [142].

Following the affinity separation, the captured protein molecules are eluted and verified by performing SDS-PAGE or other detection methods. In SDS-PAGE, the protein molecules are differentiated according to their electrophoretic mobility. The SDS, an anionic detergent, binds to the polypeptide chain of the proteins, providing a negative charge in proportion to the mass and destroys the complex structures of proteins. As a result, negatively charged proteins migrate towards the positive electrode and separated by the approximate size during electrophoresis [143].

The SDS-PAGE gels are formed by polymerizing acrylamide monomers with a bisacrylamide crosslinker [144]. In order to control the pore size, the concentration of acrylamide in the gel can be varied. The gels with higher concentrations of acrylamide

(e.g., 15%) will have smaller pores and offer better separation of low molecular weight proteins. Conversely, the gels with lower concentrations (e.g., 8%) will have bigger pore sizes that allow better resolution for high molecular weight proteins [143].

The samples for the SDS-PAGE are prepared by two different methods. In reducing

SDS-PAGE, the samples are treated with reducing agents, such as urea, dithiothreitol, or

2-mercaptoethanol, which completely unfold the proteins by reducing the disulfide bridges.

Another method is known as non-reducing PAGE, in which the use of reducing chemicals is omitted; thereby preventing the disulfide cleavage and protecting the complete unfolding of the proteins [143].

Different protein stains can be used to visualize the protein bands in the gel. The most commonly used stain is Coomassie blue; it is capable of detecting approximately 1

μg of protein in a band. Another protein stain is a silver stain, which used to detect the deficient amount of protein. The silver stain is 100 times more sensitive than the traditional

Coomassie blue stain [145]. The fluorescent stains are alternative to silver stain and offer a larger dynamic range [146]. Following the staining, further confirmation can be done by performing western blotting or the mass spectrometry of the desired bands.

2.5.2 Enzyme-linked aptamer sorbent assay

Enzyme-linked immunosorbent assay (ELISA), first developed by Engvall and

Perlmann in 1971 [147], is a well-established technology and a vital part of the biomedical research and clinical applications today. It relies on the principle of antigen-antibody interactions in combination with the enzymatic systems that develop signals corresponding to the binding results. The colorimetric signals developed in ELISA can easily be read by a standard spectrophotometric detection, eliminating the need for sophisticated and expensive equipment. Enzyme-linked immunosorbent assay reactions are usually performed in microtiter plates, which allows testing multiple samples simultaneously. It is a highly sensitive method and can detect ultralow concentrations of antigens, such as hormones, metabolites, proteins, peptides, or antibodies, in a fluid sample [148, 149].

The ELISA methods, however, have some downsides relating to the antibodies. As mentioned in section 2.1, the antibodies are expensive and time-consuming to produce, unstable at room temperature, have batch-to-batch variability, offer limited chemical modifications, and challenging to produce against non-immunogenic and toxic molecules.

The emergence of aptamers as the superior recognition element with various advantages over antibodies has demonstrated the potential to replace or complement the antibodies in several analytical applications, including ELISA [150, 151]. Besides the advantages of aptamers mentioned in section 2.1, the aptamers can be immobilized with a controlled orientation by attaching a biotin or amine functional group at the terminal. The controlled

30 orientation enables high target binding efficiency. Also, an oligonucleotide spacer can be added to the terminal functional group to create flexibility in order to enhance target binding and improve detection levels. The ELISA technique utilizing aptamer as recognition element is known as enzyme-linked aptamer sorbent assay (ELASA), and other variations of the term 'ELASA' are enzyme-linked oligonucleotide assay (ELONA), enzyme-linked aptamer assay (ELAA), and aptamer-linked immobilized sorbent assay

(ALISA) [151].

Similar to ELISA, different configurations for ELASA exist depending on the availability of aptamers or other recognition elements that can bind to the different regions of the target. Basically, the ELASA configurations can be divided into the direct, indirect, sandwich, and competitive formats.

In direct ELASA (Figure 2-5), the target molecule is first immobilized into the microtiter wells, and the remaining sites are blocked with a blocking agent. After that, the biotinylated aptamer is reacted with the immobilized target, and unbound aptamer sequences are washed. Following the washing, streptavidin-conjugated enzyme, mostly horseradish peroxidase (HRP), is introduced, followed by the addition of an appropriate

HRP substrate. Commonly used substrates are 3,3′,5,5′-tetramethylbenzidine (TMB), 2,2'- azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS), and 3,3'-Diaminobenzidine

(DAB). Among three substrates, TMB is most widely used due to its high signal to noise ratio. The direct format ELASA is the simplest form of ELASA and mainly used to compare the affinities of aptamer candidates and estimate the 퐾푑 value of the aptamer- target complexes. Also, when only one aptamer is available, direct ELASA is the preferred method. The direct format ELASA has been utilized to characterize aptamers against

31 various targets, including acetylcholine [152], human growth hormone [153], arbovirus

[154], and anthrax protective antigen [155]. In this study, direct format has been implemented to characterize the aptamers and determine the 퐾푑 value.

For the indirect format (Figure 2-5), the target is immobilized on the well surface, and the capture aptamer/antibody is introduced so that it binds to the target. Next, an enzyme-labeled secondary antibody/aptamer that is specific to capture affinity molecule is utilized. A substrate for the enzyme is introduced to generate the signal. An example of this format was reported for the detection of Leishmania infantum H2A antigen. After immobilizing the antigen on the plate, the digoxigenin-labeled aptamer was used to capture the target. The HRP labeled anti-digoxigenin antibody was implemented as the additional affinity molecule [156]. This format of the ELASA is not very popular as target immobilization is nonspecific, and also requires additional recognition element.

In sandwich ELASA, the target molecule is sandwiched in between the capturing and reporting affinity molecules (Figure 2-5). This format requires a set of aptamers or aptamer and antibody that can bind to two different sites of the target molecule. The capturing aptamer/antibody is immobilized on the well surface, and the sample containing target is introduced. Following the target capturing, another aptamer/antibody labeled with an enzyme is added. Subsequently, the signal is generated by introducing the substrate. For the first time in 1996, Drolet et al. reported the use of aptamer in ELASA by demonstrating a sandwich format utilizing aptamer and antibody for detecting vascular endothelial growth factor [157]. In their configuration, the monoclonal antibody was immobilized to capture the target, and fluorescein tagged RNA aptamer was utilized as the reporting affinity molecule (antibody-target-aptamer). The antibody-target-aptamer format of the sandwich

32 assay was also developed for the detection of aflatoxin B1 [158]. Other demonstrated configurations of the sandwich ELASA include aptamer-target-aptamer [159-162] and aptamer-target-capture antibody-detection antibody [163]. Sandwich ELASA is considered as a better version of ELASA compared to direct and indirect formats as it does not require target immobilization.

Another format of the ELASA is known as competitive ELASA. In this configuration, a competitive reaction between the target in the sample and target immobilized in microtiter plate occurs for binding to capture aptamer (Figure 2-5). In a typical assay, the capture aptamer is incubated with the sample first, and the resulting sample is added to the well that already contains immobilized target molecules. The higher the target concentration in the sample, the more capture aptamer binds to the target, leaving a smaller amount of capture aptamer available to bind to the target coated on the well. The corresponding signal is then generated similarly to the direct ELASA. This technique was developed for the detection of tetracycline [164], oxytetracycline [165], and antibodies to

Mycoplasma bovis [166].

Figure 2-5: Schematic representation of different ELASA formats.

2.5.3 Colorimetric assays based on unmodified AuNPs and aptamers

Gold nanoparticles have attracted tremendous attention in biosensing and other biomedical applications due to their unique attributes, such as tunable optical properties, facile synthesis methods, large surface-to-volume ratio, easy and well-established functionalization procedures, multiplexing abilities, stability, and biocompatibility [167].

For the first time in 1857, Michael Faraday chemically synthesized AuNPs by reduction of a chloroauric solution with phosphorus dissolved in carbon disulfide [168]. Since then, several other approaches have been developed for the synthesis of AuNPs in various sizes and shapes. In general, the synthesis methods chemically reduce chloroauric acid (HAuCl4) in the presence of capping reagents. Among the reported techniques, the most frequently utilized is the citrate method reported by Turkevitch et al. in 1951 [169]. In this process, the preheated sodium citrate solution is added to boiling water containing HAuCl4 salt to reduce Au (III) ions to metallic Au (0). The Au (0) species are thermodynamically unstable and act as the center of nucleation for further reduction. The citrate molecules in the reaction act as both reducing and stabilizing agents. In order to control the size of AuNPs

(up to 150 nm), the amount of sodium citrate is varied [170]. Other commonly used synthesis techniques include Brust-Schiffrin method, which is utilized to achieve the synthesis of small AuNPs (1-5 nm) [171] and seed-mediated growth methods to produce nonspherical shapes like rods [172], prisms [173], cubes [174], stars [175], and shells

[176].

The optical and electronic properties of the AuNPs arise due to localized surface plasmon resonance (LSPR). When the incident photons strike the surface of AuNPs or other metal nanostructures (푅/휆<0.1, where 푅 is the radius of nanoparticle and 휆 is the wavelength of the incident light), the electromagnetic wave induces collective oscillation of surface-confined conduction electrons. The collective oscillation of the electrons displaces the electron cloud from its equilibrium position, which is reinstated by coulombic restoring forces forming an oscillating dipole (Figure 2-6). When the frequency of the incident light matches the frequency of the electron oscillation, the phenomenon is known

35 as LSPR, and it leads to the strong scattering and absorption of the incident electromagnetic wave. The extinction cross-section (퐶푒푥푡) of the spherical AuNPs can be obtained by Mei’s solution to Maxwell’s equations [177, 178].

2 3 3/2 24휋 푅 휀푚 푁 휀푖(휆) 퐶푒푥푡= 2 2 휆ln (10) (휀푟(휆) + 2휀푚) + 휀푖(휆)

Where 휀푚 is the surrounding medium’s dielectric constant, 휀푟(휆) and 휀푖(휆) are the real and imaginary component of the nanoparticle’s dielectric constant, respectively, and

푁 is the electron density. When the condition 휀푟(휆) = −2휀푚 is satisfied, the electromagnetic field is enhanced relative to the incident field, and LSPR peak is observed.

In the case of AuNPs, LSPR peaks are present in the visible region of the spectrum, making these nanoparticles ideal colorimetric reporters for biological analysis. The resonance condition of AuNPs depends on the size, shape, surface chemistry, dielectric environment, electron density, and the aggregation state [178, 179]. This means that highly sensitive colorimetric sensors can be designed by employing different mechanisms to control the resonance parameters of the AuNPs.

Figure 2-6 : Schematic illustrating the oscillations of surface plasmons in a spherical metal nanoparticle. Adopted with permission from reference [180].

The colorimetric assays utilizing AuNPs and aptamer are based on the change in optical properties corresponding to the aggregation state or inter-particle distance. Small

AuNPs (~30 nm diameter) absorb light in the blue-green portion of the spectrum and reflect the red light, displaying a vibrant red color. When AuNPs come into close proximity with each other (below 2.5 times the diameter of nanoparticle), near-field electromagnetic coupling takes place as the conduction electrons on each particle surface collectively oscillate. The strong enhancement of the localized electric field within the interparticle spacing causes the plasmon resonance wavelength of the coupled particles to red-shift to longer wavelengths (lower energies) in order to lower the energy state of the oscillating electrons. The red light is then adsorbed, and the blue light is reflected. Therefore, the dispersed AuNPs (~30 nm diameter) appear red, while the aggregation is triggered, AuNPs display a purple to blue color depending on the degree of aggregation, which is also visible to the naked eye [167].

The implementation of color change associated with the inter-particle distance for biosensing was first reported by Mirkin et al., in which they demonstrated the detection of a DNA duplex by using thiolated ssDNA modified AuNPs [181]. In their subsequent study,

Mirkin’s group also detected polynucleotides by using AuNPs modified with mercaptoalkyloligonucleotide [182]. Mirkin’s method, however, required immobilization of oligonucleotides, which involved thiol or mercaptan modification of oligonucleotides and additional steps for the immobilization. These extra steps were rather time-consuming and relatively costly.

In 2006, Li and Rothberg made an exciting discovery that the ssDNA and dsDNA have different binding affinities for AuNPs in colloidal solution. The ssDNA can adsorb on the surface of citrate capped AuNPs and stabilize against salt concentrations that would ordinarily screen the repulsive interactions of the citrate ions, whereas dsDNA cannot adsorb and consequently unable to protect AuNPs against salt-induced aggregation. By using this difference in electrostatic interaction of ssDNA and dsDNA with AuNPs, the Li and Rothberg’s group was able to discriminate single base mismatch between the probe and target in a hybridization assay. The strategy was simple, cost-effective, and less time consuming as it did not require modification and immobilization steps [183, 184]. Since Li and Rothberg’s discovery, the unmodified AuNP-based method has been reported for the colorimetric detection of various targets, including proteins [185], small molecules [186,

187], drugs [188-190], inorganic ions [191, 192], and even cells [193, 194]. All these assays were based on the principle that in the absence of target analytes, ssDNA aptamers adsorbed to the AuNPs and stabilized at high salt concentrations. In contrast, upon addition of target molecules, the aptamers switched structural conformation from random coil structures to more rigid structures and formed aptamer-target complexes, which did not adsorb to the AuNPs, causing the aggregation while adding salt.

2.5.4 Surface-enhanced Raman scattering (SERS)-based magnetic aptasensor

Raman scattering is the inelastic scattering of light, and this phenomenon was first discovered in 1928 by C. V Raman his student K. S. Krishnan [195], for which C. V.

Raman was awarded The Nobel Prize in Physics in 1930. When light interacts with matter, the light is scattered in two different ways. Most of the photons are scattered elastically, commonly known as Rayleigh scattering. While a few, approximately one in a million, are

38 scattered inelastically and referred to as Raman scattering [196]. Figure 2-7 displays both phenomena using energy level diagrams. Following the interaction with an incident photon, the molecule in the ground electronic state (V=0) or excited phonon state (V=1) absorbs the energy of the incident photon and gets excited to a virtual energy state. The excited molecule instantly relaxes back to the ground state and can have three different potential outcomes. First, the emitted photon can have equal energy to that of the incident photon, which is an elastic process and referred to as Rayleigh scattering. The second possible outcome is the emission of lower energy photon than the incident photon; this occurs when the excited photon relaxes back to the phonon state (V=1), and the process is called Stokes

Raman scattering. The third possibility is that the emitted photon has higher energy to that of the incident photon and the scattering is known as Anti-Stokes Raman scattering, which occurs when a molecule from a phonon state (V=1) is excited to a higher virtual state and then relaxes to the ground electronic state (V=0). In most of the Raman scattering, the

Stokes scattering is more desirable than the Anti-Stokes scattering, because, at room temperature, the ground vibrational level (V = 0) is highly populated than the excited phonon state (V=1) [197].

Figure 2-7: Energy-level diagram displaying the states involved in Rayleigh and Raman scattering. As the energy shift of the emitted photon results from the interaction of light with specific molecular vibrations that are unique to a molecule's chemical bonds and symmetry,

Raman scattering can be used as a molecular fingerprint of a molecule. Also, the method of obtaining a Raman spectrum is fast, non-destructive, not interfered by water, offers facile sample preparation method, and requires a small amount of sample [198]. Despite these advantages, the use of Raman spectroscopy for sensitive detection was limited because of the inherently lower efficiency of Raman scattering. Raman molecules exhibit cross-sections of the order of 10-30-10-27 cm2 molecule-1 [199], whereas the fluorescent molecule’s cross-section is in the order of 10-16 cm2 molecule-1 [200], making Raman spectroscopy inferior to traditional fluorescence spectroscopy. This limitation of Raman spectroscopy can be overcome by taking advantage of the SERS effect.

Surface-enhanced Raman scattering is a phenomenon that enhances the inelastic scattering of incident light by employing plasmonic nanostructures. It was first discovered in 1974 by Fleischmann et al., while measuring Raman signals for pyridine adsorbed on the roughened silver electrodes [201]. This historical discovery led to the development of

SERS as a powerful analytical technique offering all the merits of classical Raman spectroscopy and additionally providing sensitivity and selectivity for detection. It has found application in various analytical fields, including trace detection in food safety, environmental monitoring, diagnostics, pharmaceutical, forensic science, clinical chemistry, material science, and biomedicine [196, 202].

Typically, plasmonic nanostructures based on silver and gold have been most commonly used for the SERS application. Silver has the highest SERS enhancement capability for the identical structure due to its favorable dielectric function, while gold has better chemical stability and biocompatibility. However, both metal nanostructures have been extensively used for SERS measurement [202].

Surface-enhanced Raman scattering is a highly sensitive technique that can detect up to a single molecule adsorbed on the metallic nanostructures [203, 204]. The enhancement of the Raman signal can be explained by two mechanisms, namely electromagnetic (EM) enhancement and chemical enhancement (CE). Electromagnetic enhancement is typically induced by the enhanced EM fields, localized within a few nanometers of metallic nanostructures, formed by SPR (Figure 2-8 (A)). If an analyte is located in the enhanced EM field, it experiences the much-enhanced intensity of incident light, and subsequently, the Raman scattering of the molecule is also magnified [205]. In general, EM enhancement is the major component of the total SERS enhancement, and

41 enhancement can range from 104 to 107 times. The mechanism of CE is still under debate; however, it is believed to be due to resonant charge transfer between the chemically adsorbed molecule and nanostructure. The contribution of chemical enhancement is only

10-100 times. Usually, the EM enhancement and CE coexist and work together to yield the overall SERS enhancement effect [202, 206]. The areas of extremely large SERS enhancement are defined as “hot spots,” which occur within the interstitial crevices present in metallic nanostructures or within the small gaps (<10 nm) between the nanoparticles

(Figure 2-8 (B)). When analyte molecules are adsorbed on “hot spots,” SERS signals of the analyte can be enhanced up to 1011 , achieving up to a single molecule detection [207,

208].

Figure 2-8: The EM field distribution of a AuNP (A) [209] and “hot spot” between two aggregated AuNPs (B) [210]. Adopted with permission from respective references.

Overall, SERS-based sensing approaches can be classified into two categories, namely the label-free and label-based methods. The label-free assays directly measure the intrinsic SERS spectrum of an analyte, which is either directly adsorbed [211, 212] or captured by recognition elements [213, 214] on to the SERS substrate. This method fails when the target molecule has weak intrinsic Raman signals. By contrast, the label-based

42 approaches employ a SERS label, usually called SERS-tag, that utilizes Raman signature peaks of a reporter molecule attached to metal nanostructures for the detection of analytes.

The SERS tags are usually composed of plasmonic nanoparticle core modified with Raman reporter molecule, biorecognition element, and sometimes contains a protective outer layer for enhancing stability and biocompatibility [205]. The plasmonic core amplifies the

Raman signal of the reporter molecule attached to it upon excitation with the laser. A wide array of plasmonic nanostructures, like gold and silver nanospheres, bimetallic nanoparticles, nanocubes, nanorods, nanoflowers, and several other nanostructures, have been implemented as SERS substrates [210]. The Raman reporter molecules are required to possess some characteristics, such as a large Raman cross-section for producing intense spectral signals, unique Raman peaks not non-overlapping with those of other biomolecules, high affinity toward metallic surfaces, and stability against the robust external environment. Various Raman molecules have been reported in the literature, including 4-mercaptobenzoic acid (4-MBA), 4-aminothiophenol (4-ATP), rhodamine 6 G

(R6G), malachite green isothiocyanate (MGITC), 4-mercaptopyridine (4-MPY), and 5,5- dithiobis-2-nitrobenzoicacid (DTNB). The choice of Raman molecule also depends on its solubility. While applying SERS tags in biological samples, the water-soluble molecules may degrade over time, compromising the stability and reproducibility of the SERS measurement. Therefore, the labels with low water solubility such as MGITC or 4-NTP are preferred over water-soluble molecules [209]. Usually, antibodies and aptamers are implemented in the SERS tag to provide specificity. The use of antibodies in SERS assays is less preferred due to the large size of the antibody, which adds distance between the target and substrate, making it challenging to enhance the Raman signal. Compared to the

43 antibodies, the aptamers have a smaller size and several other advantages, which make them better affinity molecules for SERS assay [215].

The fabrication of a high-performance SERS substrate is an essential factor for the biosensing application. Different types of SERS substrates have been constructed generally by using metal nanoparticles in suspension or solid-phase nanostructures. The solution- based nanoparticles can induce high SERS enhancement by forming “hot spots” through aggregation in solution. However, the particle aggregation in the liquid is difficult to control, making it challenging to obtain better reproducibility. Also, the washing step in the solution phase is time-consuming as it utilizes centrifugation. These problems with the solution-phase nanoparticle substrate can be mitigated by implementing a solid phase substrate with ordered nanostructures [202]. Although solid-phase substrates offer easier detection methods than solution-phase nanoparticles, the use of a solid-phase surface also possesses some drawbacks. The fabrication of ordered nanostructures is a tedious process and utilizes sophisticated instruments, making these substrates expensive [216]. The loading density of aptamer molecules on the solid substrate is limited because of the small surface-to-volume ratio. Also, the binding steps are time-consuming due to the slower kinetics resulting from limiting the molecular diffusion rate near the solid surface. All of these problems can be resolved by using MBs as capturing substrate to fabricate a quick and reproducible SERS-based assay. Magnetic beads are three-dimensional microspheres and have a higher surface-to-volume ratio to support the increased loading of aptamers. As the binding reactions take place in the solution phase, the use of MBs also overcomes the slow reaction problems caused by diffusion-limited kinetics on the solid substrate. In

44 addition, the separation of MBs form solution is facile and can be obtained with an external magnet, making washing processes very simple [217, 218].

Several assays have been reported by using MBs as a substrate for SERS. Gong et al. in 2007 demonstrated the SERS-based analytical method for α-fetoprotein by combining SERS tag fabricated with polyclonal antibody, Raman reporter molecule, and

Ag/SiO2 nanoparticle, and capturing substrate made up of monoclonal antibody modified silica-coated magnetic nanoparticles. In their study, Gong’s group was able to obtain a detection limit of 11.5 pg/ml [219]. A similar technique by using antibodies as affinity molecule was reported by Chon et al. for detecting carcinoembryonic antigen. Chon et al. utilized hollow AuNPs for fabricating SERS tag and were able to obtain a detection limit of 1-10 pg/ml, which was 100-1000 times more sensitive than the ELISA [217]. This assay format was later implemented with aptamer for the detection of thrombin. Two different aptamers that bind to the two different epitopes of thrombin were utilized to fabricate the

SERS tag and MB-based capture substrate, obtaining the detection limit of 0.27 pM [218].

Recently, another study for the detection of prostate-specific antigen was reported by Yang et al. To perform the detection, the assemblies between the aptamer-modified MBs and complementary sequence functionalized AuNPs were formed first. The assemblies were then incubated with the sample containing the target molecule. When the target was present in the sample, the AuNPs/SERS tags were released form the MBs due to the higher affinity of the aptamer to the target. The supernatant containing the released SERS tags was tested for correlating with the target concentration. The assay displayed an excellent detection limit of 5.0 pg/ml [220]. In our study, we employed an aptamer and its complementary sequence for assay development; however, the implementation was slightly different from

Yang et at. The aptamer was immobilized on the SERS tags/AuNPs, which was fabricated by using AuNPs and 4-NTP, and the complementary sequence was functionalized on the

MBs as a capture substrate. Instead of monitoring supernatant, the assemblies formed between the aptamer containing SERS tag and complementary sequence immobilized MBs were tested.

2.5.5 Aptamer-based lateral flow test strip

Lateral flow test strips, also known as lateral flow assay (LFA) strips, present a promising alternative to traditional laboratory-based diagnostic tests by allowing the simple, rapid, and low-cost diagnostic services in POC settings. In terms of application, the end-users, which could be non-specialized personnel, only have to add the small amount of crude sample on the device and read the test results visually after a couple of minutes [221, 222]. One excellent example of the LFA strip is a urine-based pregnancy test kit, which can be bought over-the-counter in various stores for testing pregnancy at home.

The LFAs also support the testing of a variety of other biological samples, including sweat

[223], saliva [224, 225], and whole blood [226, 227] without requiring any preprocessing.

Besides diagnostics, LFA-based technologies have found extensive applications in several other sectors, such as product safety and food inspection [228], environmental monitoring

[229], veterinary medicine [230], and quality control [231], among others.

The concept of LFA is to generate a visible signals/lines on the chromatographic membrane by using the interaction of the analyte with molecular recognition element while the sample moves laterally through the chromatographic system via capillary force [232].

An LFA test strip (Figure 2-9) consists of a sample pad, conjugate pad, nitrocellulose membrane, and absorbent pad mounted on an adhesive backing that provides mechanical

46 support. In a typical test, the sample is applied to the sample pad, which filters some components of the crude sample, distributes the sample uniformly across the strip, and adjusts the assay conditions (pH and ionic concentrations) based on its treatment with buffer salts and surfactants. The sample then migrates to the conjugation pad, where it rehydrates detection labels impregnated and dried on the conjugate pad. The detection labels, also known as conjugates, are composed of the recognition molecules, such as antibodies or aptamers, coupled to the reporter particles like AuNPs, latex microspheres, fluorophores, or quantum dots. The analytes in the sample interact with the recognition elements on conjugates, while the sample, along with interacting conjugates, move forward to the membrane. The membrane, mostly made up of nitrocellulose, contains the test and control zones made up of immobilized capturing molecules, such as antibodies, aptamers, proteins, or oligonucleotides. In the test and control lines, the immobilized capturing molecules bind to the target or the recognition molecules on the detection label; thereby, demonstrating the colored line with variable intensity on the test zone and a clear visible line on the control zone. The response on the test zone indicates the presence or absence of the biomolecule of interest, while the response on the control zone validates the test. The remaining sample flows to the absorbent pad, which wicks the excess reagents and prevents backflow [221, 222]. Mostly the LFAs are qualitative, and the results can be read by the naked eye; however, by using a strip reader [233] or a smartphone [234], the color of the test zone can be quantified, rendering the LFAs quantitative. Some researchers have also reported multiplexing capability of LFAs by fabricating multiple test lines on the membrane [235].

Traditionally, antibodies were extensively used as recognition elements for the development of LFAs; however, recently, aptamers are replacing antibodies in LFAs because of their various advantages over antibodies. Aptamers offer several advantages suitable for LFAs, including lower cost, enhanced stability at a wide temperature range, and easier methods for preparations of aptamer-labeled detection labels [221].

Figure 2-9: Typical configuration of the LFA strip. Adopted from reference [236] with permission.

In terms of format, the aptamer-based LFAs can be divided into the sandwich and competitive formats. A sandwich format is preferred when the target is a large molecule having multiple binding sites or when a pair of aptamers binding to two different epitopes of the target is available. In this configuration, an aptamer-conjugated detection label reacts with the target analyte, and the resulting complex is captured by another aptamer immobilized on the test zone of the nitrocellulose membrane. The appearance of lines at the test zone and control zone represents a positive result, while a single line at the control zone indicates a negative result [237]. By using a pair of aptamers, the LFAs for several target molecules, such as thrombin [238], cancer cells [239], vaspin [240], and tick-borne encephalitis virus [154], have been developed. Additionally, the sandwich format has been

48 demonstrated by using the combination of aptamer and antibody for the detection of salivary α-amylase [241]. Also, by using the splitting aptamer, the sandwich format LFA was developed for the detection of ATP [242].

The competitive configuration is utilized whenever the target molecule is small, with only one epitope or when only one aptamer is available. Two different formats of the competitive configurations have been reported based on the capture molecule on the test zone. The capture molecule on the test zone could be either a target molecule or a partially complementary DNA sequence to the aptamer. Whenever the target is immobilized on the test line, the competition occurs between the target in sample and target immobilized on the test line for binding to the aptamer-modified reporter molecule. While in other configuration, the complementary DNA sequence immobilized on the test line competes with the target molecule for binding to the aptamer-modified detection label. In both cases, the decreasing intensity of the test line color corresponds with the increasing concentration of the target analyte in the sample, while the control line should be visible to validate the test [237]. The competitive format has been employed for the detection and quantification of various target molecules, including β-conglutin [243], ochratoxin A [233, 244], and zearalenone [245].

In this study, the competitive format LFA was developed by implementing the complementary DNA sequence on the test line. The H3 aptamer was immobilized to the

AuNPs as the detection label.

Chapter 3

3 Materials and Methods

To address the goals and specific aims of this project, numerous chemicals and devices were used, and experimental methods were developed. More specifically, aptamers were selected, characterized, and implemented in various assay platforms. This chapter will outline the materials, devices, and detailed procedures used to accomplish the key tasks.

3.1 Selection and preliminary characterization of structure-switching

ssDNA aptamers for salivary peptide H3

This section describes the H3 aptamer selection process for identifying structure- switching aptamers through the library immobilization version of the SELEX method.

Following the aptamer identification, subsequent characterizations using the software and the visual confirmation of binding through MB-based capture assay coupled to SDS-PAGE have been explained.

3.1.1 Aptamer selection by immobilizing ssDNA library sequences on the MBs

3.1.1.1 Oligonucleotides for the SELEX

The ssDNA library, forward and reverse primers, and biotinylated complementary capture probes (Table 3.1) used for aptamer selection were all synthesized by Integrated

DNA Technologies (Coralville, IA). These ssDNA library sequences contain the specific primer binding sites of 20 nucleotides at the 5’ and 3’ end, and in between the primer binding sites, they contain a random region of 44 bases denoted by the term (N44). For the immobilization of ssDNA pool sequences on the MBs, biotinylated capture probes of length 7 (probe 1) and 9 (probe 2) bases, complementary to the 5’ primer binding region of the library sequence were designed.

Table 3.1: Oligonucleotides used for the SELEX process.

3.1.1.2 Preparation of ssDNA library-immobilized MBs

The ssDNA library sequences were immobilized on the Dynabeads M-280

Streptavidin MBs (Thermo Fisher Scientific, Wayne, MI) by utilizing 3’ biotinylated complementary capture probes. Briefly, a 100 µl aliquot of MBs (10 mg/ml) was washed

3 times with 500 µl of the binding buffer (BB; 50 mM Tris, 137 mM NaCl, 5 mM MgCl2, pH 7.4) and reconstituted with 5 nmol of a biotinylated capture probe in 100 µl of the same

51 buffer. After a 30-min incubation period at room temperature on a shaking platform, the beads were washed 4 times with 500 µl of BB to remove the unbound and loosely bound capture probes. The washed beads were then resuspended in 100 µl of BB and stored at 4

°C for further use. In order to immobilize the ssDNA library on the capture probe immobilized MBs, 250 pmol of the library sequences in 100 µl of BB were denatured at

95 °C for 5 minutes, snap cooled on ice, and then equilibrated at room temperature for 10 minutes. The equilibrated library sequences were then incubated with 20 µl of the probe coupled MBs for 30 minutes with gentle agitation. These beads were then washed 5 times to get rid of unbound and loosely bound library sequences and resuspended in 20 µl of BB.

All wash steps consisted of a gentle agitation for 5 minutes and then 2 minutes of incubation of beads with an external magnet to remove the supernatant.

3.1.1.3 In vitro selection procedure

Iterative rounds of in vitro selection were carried out as shown in Figure 3-1. A 20

µl aliquot of library immobilized MBs was mixed with 40 µl of H3 (Genemed Synthesis,

San Antonio, TX) diluted with BB. The reaction was incubated for 30 minutes with gentle agitation. During the incubation, the sequences binding to the H3 were released from the magnetic beads due to conformational change induced by the binding event. The supernatant, containing H3 and H3 binding sequences, was retained and then dialyzed using the Slide-A-Lyzer 20K molecular weight cut-off dialysis cassettes (Thermo Fisher

Scientific, Wayne, MI) to get rid of excess H3. Dialysis was performed according to the manufacturer’s protocol. In short, the supernatant was introduced into the cassette, and the cassette was put into the ultrapure water (Direct-Q® 3 UV water purification system, EMD

Millipore, Darmstadt, Germany) for 2 hours. The ultrapure water was then replaced, and

52 the dialysis was performed for another 2 hours. After replacing the water one more time, the cassette was placed at 4 °C overnight. The dialyzed sample was then used for PCR amplification to generate a new pool of DNA sequences for the subsequent round of

SELEX. Starting from the sixth round, counter-selection with H8 (Genemed Synthesis, San

Antonio, TX) in every other round was executed. For the counter-selection, library immobilized MBs were incubated with H8 before selection with H3. The sequences having an affinity to H8 were released into the supernatant and were discarded, whereas the MBs containing the library sequences were retained for further selection. The selection process was repeated for 14 rounds with varied concentrations of H3 and H8 (Table 4.1). From round 10 onwards, capture probe 1 was replaced by capture probe 2 to increase the stringency of selection.

Figure 3-1: Schematic diagram of SELEX for H3 aptamer selection. The biotinylated capture probes were immobilized on the MBs, and ssDNA library sequences were hybridized to the capture probes (1). After washing multiple times using an external magnet to remove non-bound and loosely-bound ssDNA sequences, the target peptide H3 was introduced to the library immobilized MBs (2). Provided sufficient incubation time, H3 binding sequences were released due to conformational change induced by the target binding (3). The supernatant containing the high-affinity sequences was retained (4) and dialyzed to get rid of excess H3 (5). Following the dialysis, the H3 binding sequences were PCR amplified and converted to ssDNA to generate a new library for the next round of SELEX (6). Counter-selection with H8 was started at round 6 and performed in every other round thereafter. To execute the counter-selection, library sequences from the previous selection round were immobilized on the magnetic beads and were exposed to the H8 (7). The sequences binding to H8 were released from the beads and were discarded by washing several times (8). The MBs containing the remaining library sequences were retained for further selection with H3. Overall, the SLEX process was performed for 14 rounds, and after the final round of SELEX, the high-affinity sequences were identified through sequencing (9).

3.1.1.4 Polymerase chain reaction amplification

The PCR amplification was performed with 25 µl of reaction volume using the

MyiQ Single-Color Real-Time PCR Detection System (Bio-Rad Laboratories, Hercules,

CA). Each reaction contained dialyzed supernatant, 500 nM of each primer, and SYBR

Green Supermix (Bio-Rad Laboratories, Hercules, CA). The PCR was carried out as follows: initial denaturation at 94 °C for 2 minutes followed by 16 amplification cycles of

30 seconds denaturation at 94 °C, 1-minute annealing at 60 °C, 1-minute extension at 72

°C, and the final extension of 7 minutes at 72 °C. After the PCR, the product was verified by running in 2% agarose gel electrophoresis in 1X Tris/borate/EDTA buffer at 100 V, 25 mA for 2 hours (Details for running agarose gel electrophoresis are provided in Appendix

B). The double-stranded DNA sequences were converted to ssDNA by asymmetric PCR using forward primer only. For the asymmetric PCR, the reaction product from the first

PCR was further added with SYBR Green Supermix, forward primer, and ultrapure water.

The amplification protocol was the same as before.

3.1.1.5 Transformation, cloning, and sequencing

To identify the potential aptamer sequences, transformation, cloning, and sequencing were performed. First, the PCR product was ligated into the pCRTM 2.1-

TOPO® vector from a TOPO® TA cloning kit (Invitrogen, Carlsbad, CA). For ligation, the aptamer sequences were 3’ adenylated by using a final extension step of 15 min during

PCR and then mixed with the plasmid vectors for 30 minutes at room temperature. The ligated vectors were then inserted into the One Shot TOP 10 Electrocomp E. coli cells

(Invitrogen, Carlsbad, CA) by electroporation at 1.5 kV (ECM 399 Electroporation system,

BTX, Holliston, MA). Following the electroporation, the cells were plated on the Luria-

Bertani (LB) medium plates containing kanamycin (Fisher Scientific, Pittsburgh, PA) and

X-gal (Indofine Chemical Company, Hillsborough, NJ) and incubated at 37 ºC for 12 hours. The white colonies on the plate were then shipped to Functional Biosciences

(Madison, WI) for sequencing. All the experimental details for transformation and cloning have been explained in Appendix A.7.

3.1.2 Analysis of aptamer sequences, structural folding, and stability

Prediction of lowest free energy secondary structures and Gibbs free energy change

(ΔG) of the aptamer candidates was executed by using the online server mFold

(http://unafold.rna.albany.edu/?q=mfold/DNA-Folding-Form) at 25 °C in 137 mM [Na+] and 5 mM [Mg+2] [132]. The possible G-quadruplexes in the aptamer sequences were predicted using a web-based server, QGRS mapper

[http://bioinformatics.ramapo.edu/QGRS/analyze.php] [133].

3.1.3 Demonstration of binding through MB-based affinity capture assay coupled

to SDS-PAGE

Based on the mFOLD and QGRS mapper analysis, the most promising aptamer sequence was chosen to assess the binding between the target molecule and aptamer. The magnetic bead-based affinity capture was combined with SDS-PAGE for visual selective binding confirmation (Figure 3-2). Briefly, amine-terminated full-length aptamer, LI-H3-

APT-3P (5’-

/AmMC6/GGTGACTGCTACTGTGTTGGACCGGGTGAGGGGGGTCCAGTGTTAG

TAGCGATGGAGGGGTGACCCACACATCCAAGCAGAACC-3’), was immobilized on the Dynabeads M-280 Tosylactivated MBs (Thermo Fisher Scientific, Wayne, MI) using the manufacturer's instructions. Briefly, a 165 µl aliquot of MBs was washed with 1

56 ml of buffer B (0.1 M Na-phosphate buffer, pH 7.4), and the washed MBs were resuspended in 50 µl 100 µM of amine-terminated aptamer solution. The final volume of the sample was made 150 µl by adding 100 µl of buffer B. An additional 100 µl of buffer

C (3 M ammonium sulfate in buffer B) was also supplemented to the sample, which was then incubated on a roller at 37 °C for 12 hours. By using an external magnet, the supernatant was removed, and 1 ml of buffer D (10 mM PBS, pH 7.4 with 0.5% (w/v)

BSA) was added. The sample was then incubated for an hour at 37 °C on a roller. Following the incubation, the MBs were washed 3 times with 1 ml of buffer D and resuspended in the same buffer at a final concentration of 20 mg/ml. Before testing, the buffer D was replaced with H3 BB. For testing, the H3 and H8 samples (500 µl of 20 µg/ml in BB) were incubated with aptamer immobilized MBs (50 µl of 20 mg/ml) for about an hour at 37 ˚C with gentle shaking. Following the incubation, the samples were washed 7 times with BB using an external magnet to pull down the MBs. The reaction tubes were replaced after every two consecutive washes to get rid of the H3 and H8 nonspecifically binding to the tube surface.

Following the washing, the beads were reconstituted in 10 µl of the ultrapure water. To elute the aptamer bound peptides, 10 µl of 2x Laemmli sample buffer containing β- mercaptoethanol (5%) was added to each sample and heated for 10 minutes at 100 ˚C in a dry bath. The supernatants were retained and run in a 12% Mini-PROTEAN TGX Precast

Gel (Bio-Rad Laboratories, Hercules, CA) for 1 hour at 100 V and 25 mA using 1x SDS-

PAGE buffer (25 mM Tris, 0.192 M glycine, and 0.1 % SDS (pH 8.3)). The gel was then stained with Coomassie brilliant blue R250 by incubating in a staining solution (45% methanol, 10% glacial acetic acid, 45% water, and 3 g/L Coomassie brilliant blue R250) for 4 hours (or until the gel had a uniform color). The stained gel was destained for further

4 hours in a destaining solution (5% methanol, 7.5% glacial acetic acid, 87.5% water).

Finally, the destained gel was visualized to observe the protein bands. The detailed protocol for running SDS-PAGE is listed in Appendix C.

Figure 3-2: Magnetic bead-based affinity capture coupled to SDS-PAGE for visual monitoring of binding.

3.2 Integration and demonstration of the H3-selective aptamers into

ultrasensitive assay formats to allow for the identification and

quantification in complex media

Within this section, various assays have been established for the characterization and application of the aptamers. Initially, ELASA was developed, and the multiple aptamers were compared for their affinity to the H3. The best performing candidate was identified and truncated to enhance the binding affinity. The truncated sequence was then

58 employed for the development of various detection schemes including AuNPs-based colorimetric assays, SERS-based magnetic aptasensor, and lateral flow test strip for point- of-care detection.

3.2.1 Development of ELASA for the additional investigation into the binding

between the selected aptamer candidates and H3

3.2.1.1 Aptamers

Three different aptamer sequences, LI-H3-APT-3P, LI-H3-APT-4P, and LI-H3-

APT-3, with 5’ biotin modification were synthesized by Integrated DNA Technologies

(Coralville, IA).

Table 3.2: Biotinylated aptamers used in ELASA.

3.2.1.2 Binding analysis using ELASA

To evaluate relative binding affinity and cross-reactivity of the selected aptamers

(LI-H3-APT-3P, LI-H3-APT-4P, and LI-H3-APT-3), a direct format ELASA (Figure 3-3) was developed using amine-binding, maleic anhydride activated plates (Pierce

Biotechnology, Rockford, IL). Peptides (H3 and H8) were diluted to 40 µg/ml in immobilization buffer (IB, 0.1 M NaH2PO4 and 0.15 M NaCl, pH 7.2), and 100 µl/well of these peptide solutions were incubated for 1.5 hours at 37 ˚C with mild shaking. The blank

59 samples were created by treating wells with 100 µl of the IB without any peptide. After peptide immobilization, the wells were washed twice with 300 µl wash buffer (Thermo

Fisher Scientific, Wayne, MI) diluted 1X with ultrapure water. Following the wash step, blocking was performed with 300 µl of 1% bovine serum albumin (BSA) for an hour at

37˚C. The BSA solution was washed twice with 300 µl of diluted wash buffer as before.

Afterward, 100 pmol biotinylated aptamers in 100 µl BB (previously heated for 5 minutes at 95 ˚C, snap cooled on ice for five minutes, and equilibrated at room temperature for 5 minutes) were added to the respective wells and incubated for an hour at room temperature with gentle agitation. These wells were again washed twice with 300 µl of 1X wash buffer to remove the unbound and loosely bound biotinylated aptamers. Followed by this washing, a 100 µl aliquot of S-HRP conjugate (Invitrogen, Carlsbad, CA) diluted 1:5000 with IB was placed in each well and hatched for an hour at room temperature. The excess

S-HRP was removed by washing three times with the 1X wash buffer as before. After washing S-HRP conjugate, a 100 µl aliquot of 1-Step™ Ultra TMB substrate solution

(Thermo Fisher Scientific, Wayne, MI) was added to each well. The plate was incubated for 12 minutes in the dark at room temperature, after which 70 µl of 2 M H2SO4 was added to stop the reaction. The plate was read at 450 nm with the UV-Vis spectrophotometer

(SpectraMax plus 384 Microplate Reader, Molecular Devices, Sunnyvale, CA).

Figure 3-3: Schematic illustration of direct format ELASA.

3.2.1.3 Binding kinetics study

In order to explore the binding kinetics of the aptamer with H3, an ELASA experiment was designed. The target molecule H3 (4 µg) was coated on the wells, and the remaining sites were blocked with BSA. Each well was then reacted with 100 pmol of the aptamer LI-H3-APT-3 for different time periods (5, 30, 60, and 90 min). The signal was developed using S-HRP and TMB, as explained in section 3.2.1.2.

3.2.1.4 Determination Kd

The was determined only for the best performing aptamer sequence (LI-H3-

APT-3) using an ELASA assay. Briefly, a fixed amount of H3 (4 µg) was immobilized in the amine-binding, maleic anhydride activated plate wells. After performing the washing and blocking steps, 100 µl of different concentrations (1 nM, 10 nM, 100 nM, 500 nM, 1

µM, 5 µM, and 10 µM) of biotinylated aptamer diluted in BB were reacted to the immobilized H3 in the wells. The assay signal was developed using S-HRP conjugate and

TMB substrate, according to section 3.2.1.2. To estimate the Kd , plots of the OD at 450 nm as a function of aptamer concentration were generated using one site-specific binding model in GraphPad Prism 7.02 (San Deigo, CA).

3.2.2 Development of AuNPs-based colorimetric assays based on the structural

feature of the identified H3 aptamers

The structural feature, structure-switching upon target binding, of the identified aptamer was implemented to develop two different formats of the AuNPs-based colorimetric assays for the detection and quantification of H3. The highest affinity aptamer sequence (LI-H3-APT-3) without any modifications (5’-

ACCGGGTGAGGGGGGTCCAGTGTTAGTAGCGATGGAGGGGTGAC-3’;

Integrated DNA Technologies, Coralville, IA), and commercially available citrate- stabilized AuNPs with 15 nm ± 1.3 nm diameter (nanoComposix, San Diego, CA) were used.

In assay format-1, the aptamers were first adsorbed on the surface of the citrate- stabilized AuNPs, and then the target molecules were introduced to interact with the AuNP- aptamer colloid. Various parameters of the assay were investigated to optimize the response in format-1. For assay format-2, free aptamer and target molecule were first reacted to allow the binding to happen, following that the AuNP solution was introduced.

With the format-2, the overall assay time was just 12 minutes.

3.2.2.1 Assay format-1

In a typical experiment, 10 µl of 25 µM aptamer solution in TE buffer was mixed with 200 µl of 2.6 nM AuNPs in 2 mM sodium citrate buffer (molar ratio of AuNP to aptamer is 1:480). After incubating for 4 hours, 10 µl of H3 diluted in H3 BB was added

62 to the AuNP-aptamer solution and further incubated for 30 minutes. Subsequently, the sample was transferred to the 96-well UV-Vis plate, and 22 µl of 453 mM NaCl was added.

Immediately after NaCl addition, UV-Vis absorbance spectra over the wavelength range of 450 nm to 750 nm were recorded with the UV-Vis spectrophotometer (SpectraMax plus

384 Microplate Reader, Molecular Devices, Sunnyvale, CA).

3.2.2.2 Assay format-2

Briefly, 10 µl of 25 µM aptamer solution in TE buffer was mixed with 10 µl of different concentrations of H3 in H3 BB. The mixture was incubated for 5 minutes, and

200 µl of citrate-stabilized AuNP solution (2.6 nM) was introduced. Following 5 min incubation, the mixture was transferred to UV-Vis plate and challenged by 22 µl of 453 mM NaCl. Immediately, after adding NaCl, assay response was monitored by recording

UV-Vis spectra. The entire experiment took about 12 minutes.

3.2.3 Exploration of SERS-based magnetic aptasensor for the ultrasensitive

detection of H3

3.2.3.1 Oligonucleotides

Histatin 3 specific aptamer (LI-H3-APT-3) with 5’ thiol modification (5’-

/ThioMC6-

D/ACCGGGTGAGGGGGGTCCAGTGTTAGTAGCGATGGAGGGGTGAC-3’) and 5’ biotinylated capture sequence (cDNA; 5’-/Biosg/GGACCCCCCTCACCCGGT-3’) that was complementary to the first 18 sequences of the aptamer were fabricated by Integrated

DNA Technologies (Coralville, IA).

3.2.3.2 Synthesis of signal probes

The schematic representation of the signal probe fabrication is shown in Figure 3-

4. Gold nanoparticles (15 ± 1.3 nm) were first modified with the Raman reporter molecule,

4-nitrothiophenol (4-NTP; Sigma-Aldrich, St. Louis, MO). For which, 20 µl of 100 µM 4-

NTP was transferred to 1 ml of 2.6 nM AuNP solution and sonicated for 40 minutes.

Unbound 4-NTP molecules were then removed by centrifuging for 15 minutes at 13,000 rpm (Micro 12TM; EKF Diagnostics, Wales, UK) and washing (resuspension in 1 ml of 5 mM HEPES buffer (pH 7.6) and centrifugation at 13,000 rpm for 15 minutes). The washed

AuNPs-NTP conjugates were resuspended in 1 ml of 5 mM HEPES buffer for further functionalization with aptamers.

To start the aptamer conjugation, the disulfide bond of the aptamer was reduced by reacting 100 µM aptamer solution with an equal volume of freshly prepared 10 mM tris(2- carboxyethyl) phosphine (TCEP; Sigma-Aldrich, St. Louis, MO) solution at room temperature for 1 hour in the dark. Following the activation, 50 µl of aptamer solution was transferred to the above prepared AuNPs-NTP conjugate (1 ml) and incubated for 24 hours on a rotating platform. The solution was then adjusted to obtain a final NaCl concentration of 60 mM with 1 M NaCl (15 μl per hour, 4 times) and reacted for another 24 hours. Next, the excess reagents were removed by centrifugation at 13,000 rpm for 15 minutes with two times washing by HEPES buffer. The washed signal probe (AuNPs-NTP-aptamer) conjugates were resuspended in 1 ml of HEPES buffer and stored at 4 °C for further use.

The conjugation of aptamer was confirmed by the salt-induced aggregation assay and UV-

Vis measurements.

Figure 3-4: Synthesis of signal probe molecules.

3.2.3.3 Preparation of capture probes

The capture probes were prepared by immobilizing biotinylated cDNA molecules on the Dynabeads M-280 streptavidin MBs (Figure 3-5). Briefly, an aliquot of 100 µl of

MBs (10 mg/ml) was washed twice with 1 ml of 1X B&W buffer (2X B&W buffer: 10 mM Tris-HCl (pH 7.5), 1 mM EDTA, and 2 M NaCl) and resuspended in 200 µl of 2X

B&W buffer. For washing, the tube was placed on the external magnet for 1 minute, and the supernatant was discarded. Afterward, 200 µl of 5 µM biotinylated cDNA was added to the washed beads and incubated for 30 minutes using gentle rotation. Following the incubation, excess cDNA sequences were removed by washing three times with 1 ml of

1X B&W buffer using a magnetic separator. Finally, the washed MBs-cDNA conjugates were resuspended in 100 µl of BB and stored at 4ºC for further use. The functionalization of MBs with cDNA molecules was verified by measuring the absorbance of the aptamer solution at 260 nm before and after immobilization.

Figure 3-5: Schematic illustration of the capture probe.

3.2.3.4 Histatin 3 detection

To detect H3, an aliquot of 40 µl of as-prepared AuNPs-NTP-aptamer was mixed with 50 µl of various concentrations of H3 in BB and incubated for 1 hour with gentle shaking. Following this incubation, 2 µl of 10 mg/ml MBs-cDNA was transferred to the mixture and further hatched for 2 hours on the same shaking platform. Afterward, the mixture was separated by using an external magnet, and the supernatant was discarded.

The precipitate was then washed twice with 100 µl of BB. The washed conjugate was finally dispersed in 2 µl of BB and transferred to the glass slide for SERS measurement.

3.2.3.5 SERS measurement

Surface-enhanced Raman scattering signals were collected using HR 800 confocal

Raman spectrometer (Jobin-Yvon-Horiba, Edison, NJ). To obtain the spectrum, 2 µl droplet of the sample was deposited on a clean glass slide and dried. The dried sample was excited with 632 nm He-Ne laser through a 50x objective lens with a confocal pin hole of

600 µm and a 400 µm slit width. The spectrum grating was set 1800 grooves/mm. For each sample, the spectral region between 1000 cm-1 and 1800 cm-1 was retrieved. The number of accumulations was 5, and the exposure time was 1 second. A total of 5 different spots were scanned for each sample, and the average intensity was reported. The LabSpec software package (Horiba/Jobin-Yvon) was used for instrument control and data acquisition.

3.2.4 Demonstration of an aptamer-based lateral flow test strip for on-site detection

of H3

3.2.4.1 Aptamer and other oligonucleotides

Table 3.2 contains the aptamer sequence and other DNA molecules that were used for this study. The aptamer LI-H3-APT-3 was designed to contain T overhang at the 3’ end and thiol modification at the 5’ region. To fabricate the control line, a biotinylated DNA molecule with Poly A bases (Probe-CL) complementary to the T overhang region of the aptamer was utilized. For the test line, two different biotinylated complementary sequences

(Probe-TL-1 and Probe-TL-2) of varying length were examined. All sequences were obtained from Integrated DNA Technologies (Coralville, IA). The Probe-TL-1 is the same sequence as cDNA in SERS-based assay; for the ease of comparison with Probe-TL-2, it was renamed to Probe-TL-1 in this assay.

Table 3.3: Oligonucleotides used for the lateral flow test strip development. Probe-TL-1 is the same sequence as cDNA in SERS-based magnetic aptasensor.

3.2.4.2 Preparation of AuNPs-aptamer conjugates

The conjugation of the aptamer to the AuNPs (15 ± 1.3 nm) was based on the formation of the thiol-gold bond, as explained in section 3.2.3.2. Briefly, before conjugation, 5’ thiol modified aptamer (100 µM) was activated by an equal volume of 67 freshly prepared 10 mM TCEP at room temperature for an hour in the dark. A 50 µl volume of the activated aptamer (50 µM) was then transferred to 1 ml of 2.6 nM citrate-stabilized

AuNP solution. After incubating 24 hours on a slowly rotating platform, 1 M NaCl solution was added to a final concentration of 60 mM and reacted for another 24 hours. The excess reagents were then removed by centrifuging at 13,000 rpm for 15 minutes and further washed twice with 1 ml of 5 mM HEPES buffer (pH 7.6). The washed AuNPs-aptamer conjugates were then resuspended in 100 µl of HEPES buffer and stored at 4ºC for further use. The functionalization of the AuNPs was confirmed via salt-induced aggregation and

UV-Vis measurements.

3.2.4.3 Preparation of test and control zones

Figure 3-6 displays the construction process of the test and control lines on the test strip. Biotinylated probes (Probe-TL and Probe-CL) were first conjugated with the streptavidin and then stamped on the nitrocellulose membrane by using a ceramic disk. At first, 10 µl of 100 µM Probe-TL and Probe-CL were reacted separately with 10 µl of 1 mg/ml streptavidin at room temperature for 60 minutes and then diluted by adding 10 µl of

10 mM PBS (pH 7.4). Next, a 5 µl droplet of the streptavidin-conjugated probe was placed on the glass slide, and one side of the ceramic disk (0.5 mm thick) was soaked into the droplet which was then gently stamped on the nitrocellulose membrane. The distance between the test and the control line was approximately 4 mm. Following the stamping, the nitrocellulose membrane was dried at room temperature for 30 minutes.

Figure 3-6: Formation of the test and control lines on the nitrocellulose membrane.

3.2.4.4 Assembly of the lateral flow test strip

The schematic of the lateral flow test strip is shown in Figure 3-7. It was comprised of a sample pad (Millipore SIGMA, Billerica, MA), conjugate pad (Millipore SIGMA,

Billerica, MA), nitrocellulose membrane (GE Healthcare, Pittsburg, PA), and an absorbent pad (same as sample pad) fixed on a plastic adhesive backing layer (DCN Diagnostics,

Carlsbad, CA). Before assembly, the conjugate pad was blocked with the conjugate pad blocking buffer (10 mM PBS solution (pH 7.4) containing 1% ovalbumin, 0.25% Tween

20, 2% sucrose, and 0.02% sodium azide) and dried overnight at room temperature. All the components were then fixed on the plastic backing layer with 2 mm overlap with each other and cut into 5 mm wide strips with the paper trimmer. The length of the strip was about 58 mm. Later, as explained in section 3.2.4.3, control and test lines were formed on the individual strips, and the conjugate pad was loaded with AuNPs-aptamer conjugates (5 µl).

After drying AuNPs-aptamer conjugates for 20 minutes, the assembled lateral flow strips were stored at room temperature for further use.

Figure 3-7: Top (A) and side (B) views of the assembled lateral flow test strip before AuNPs-aptamer conjugate loading.

3.2.4.5 Detection procedure

The detection of H3 was carried out by applying 150 µl of the sample solution, containing the desired concentration of H3 in the conjugate pad blocking buffer, to the sample pad. After 5 minutes, the results were observed.

3.3 Investigation of the use and challenges associated with saliva-based

measurement

Within this section, the application of saliva with the developed assay platforms was studied. Various types of artificial saliva samples were prepared and tested with the

ELASA assay for its capability to detect H3 in complex media. Additional investigation

70 was performed with the AuNPs-based colorimetric assay for the possibility of its implementation with the human saliva.

3.3.1 Preparation of artificial saliva samples

Two different types of artificial saliva samples were prepared. Group A saliva was the commercially available product, used to get relief from dry mouth symptoms, Biotene

Dry Mouth Oral Rinse (GlaxoSmithKline, Warren, NJ) containing purified water, glycerin, xylitol, sorbitol, propylene glycol, poloxamer 407, sodium benzoate, hydroxyethyl cellulose, methylparaben, propylparaben, flavor, sodium phosphate, and disodium phosphate. Group B saliva was a lab-made solution having similar viscosity and electrolyte concentration to the human unstimulated whole saliva [3], and consisted of 5 mM of sodium chloride (NaCl), 1 mM of calcium chloride (CaCl2), 15 mM of potassium chloride

(KCl), 1.1 mM of potassium thiocyanate (KSCN), 4 mM of ammonium chloride (NH4Cl),

1 mM of citric acid, 5 mM uric acid, 1 mM ascorbic acid, 0.2 mM lactate, and 1% sodium carboxymethyl cellulose (CMC) in ultrapure water.

3.3.2 Application with ELASA

Both of the artificial saliva samples were utilized to demonstrate the target binding capability of the best performing aptamer sequence (LI-H3-APT-3) in the saliva like mediums. To assess the performance of the aptamer, these formulations of artificial saliva were spiked with three different concentrations of H3 (35 µg/ml, 25 µg/ml, and 15 µg/ml), and 100 µl of each sample was incubated on the amine binding wells. Based on the ELASA protocol (section 3.2.1.2), the immobilized H3 was monitored using 500 pmol of the selected biotinylated aptamer. The control samples did not contain any external H3 and were incubated with the same amount of aptamer as other samples.

3.3.3 Collection and processing of human saliva

Unstimulated whole saliva samples were collected from a healthy individual via the passive drool method. Before collection, the mouth was rinsed with the drinking water, and after waiting for 10 minutes, 2 ml of the whole saliva was collected. The collected sample was then filtered via a 0.2 µm syringe filter, aliquoted into small vials, and stored at -80

°C. Just before the application, the sample was thawed, centrifuged at 13,000 rpm for 10 minutes, and the supernatant was aspirated for the experiments. This method of saliva collection and processing provides a reliable performance to remove large biomolecules and food particles from the saliva samples.

3.3.4 Application of saliva with AuNPs-based colorimetric assay

Due to the short retention time, AuNPs-based assay format-2 was chosen for this study. Briefly, H3 doped human saliva samples and group B artificial saliva samples were first inspected, as explained in section 3.2.2.2. Further studies were performed to identify possible interfering components and to resolve the matrix effect of these biological samples. Finally, the application was demonstrated in diluted human saliva samples spiked with external H3.

Chapter 4

4 Results and Discussion

This chapter presents the results obtained through the experimental methods, discussions, and the measures taken to solve the encountered issues during this project. As mentioned in chapter 3, several tasks were accomplished, including aptamer selection, characterization, and implementation of aptamers on various assay platforms for the detection applications. Additionally, the application and the challenges of the saliva as a diagnostic fluid were investigated. All the results and corresponding discussions are outlined in this chapter.

4.1 Selection and preliminary characterization of structure-switching

ssDNA aptamers for salivary peptide H3

Within this section, the results and related discussions regarding the aptamer selection, characterization via software, and application of aptamer for the validation of binding through MB-based capture assay are presented.

4.1.1 Aptamer selection by immobilizing ssDNA library

4.1.1.1 Selection strategies

For the selection of aptamers, ssDNA library sequences were immobilized on the streptavidin-functionalized MBs via biotinylated capture probes that were complementary to the 5’ primer binding region of the library sequences. These capture probes hybridize with library sequences and hold them on the MBs. The basic principle in this type of selection is that the sequences binding to the target undergo a conformational change and get released from the complementary capture probes and, hence, from the MBs. Such detachment occurs only when the affinity between the target and aptamer overcomes the binding force between the capture probe and library sequence. Increasing the length of the complementary sequence increases the hybridization strength or the binding force between the capture probe and the ssDNA library sequence. Thus, by changing the length of the complementary capture probe, the selection stringency can be manipulated [99]. Nutiu et al. first described the idea of selecting structure-switching aptamers in which they implemented a capture probe having 15 bases complementary to the library sequence to select aptamers for nucleoside triphosphates [100]. Based on a similar idea, Morose, in

2007, selected RNA aptamers for aminoglycoside antibiotic tobramycin using a 6-mer probe having 5 bases complementary to the library sequence [246]. Stoltenburg et al., in

2012, utilized 12-mer complementary capture oligo to identify ssDNA aptamers for aminoglycoside antibiotic kanamycin [247]. Following the similar selection principle, cortisol aptamers were developed by using 7, 8, and 9-mer probes having 5, 7, and 8 bases complementary to the library sequence, respectively [99]. In this study, two different complementary capture probes of length 7 (probe 1) and 9 (probe 2) bases having 6 and 8

74 complementary bases to the 5’ primer binding sites, respectively, were designed. Besides the length of the capture probe, the concentration of the target molecule also plays a role in tuning the selection stringency. Increasing the concentration of the target molecule increases the detachment of the ssDNA sequences from the MBs [99]. Another significant aspect of this SELEX process was the counter-selection with H8, a proteolytic fragment of

H3 and hemagglutination-inhibiting peptide in human saliva [248]. Various other peptides in saliva could have been chosen for the counter-selection, but choosing a molecule that is closely related to H3 is essential. So, H8, a peptide sequence within the H3, was considered for the counter-selection. The selection conditions utilized are outlined in Table 4.1. For the initial five rounds of selection, the concentration of H3 was kept constant to 100 µM to retain all possible binding sequences. Once the pool was enriched, the counter-selection was initiated at round 6 and executed in every other round before the selection with H3.

The initial concentration of H8 was kept low (1 µM) at round 6 and gradually increased until round 14 to 60 µM. Increasing the concentration of H8 helped to get rid of sequences having a slight affinity to H8. The loss of the sequences during the counter-selection was compensated by increasing the concentration of H3 from round 6 (100 µM) to round 10

(250 µM). The increased concentration of H3 helped to retain the maximum number of sequences on the MBs after counter-selection. Following round 10, selection stringency was further increased by increasing the length of the capture probe, increasing the concentration of H8, and decreasing the concentration of H3. All three conditions favored the retention of only high-affinity sequences.

Table 4.1: Selection rounds, respective concentrations of H3 and H8, and capture probes used in the SELEX process.

4.1.1.2 Real-time PCR

In this study, real-time PCR was utilized to amplify the sequences and to troubleshoot the SELEX process. For the real-time PCR, a fluorescent dye, SYBR Green

I, was used. SYBR Green I is an intercalating dye that binds between the two strands of dsDNA and strongly flourishes when it is excited in a bound state, allowing this dye to be used as a detector of the amount of double-stranded DNA (dsDNA) in a PCR reaction [249,

250]. As can be seen in Figure 4-1 (A), a successful SELEX round resulted in an exponentially increased fluorescent intensity over the course of the PCR run (a). If there is no amplification, a straight line without any exponential component is observed (b), and it requires the repetition of the SELEX round until the amplification is achieved. Following the PCR amplification, the product was verified with agarose gel electrophoresis. The PCR product at this point must appear as a single band on the gel as shown in Figure 4-1 (B); if

76 multiple bands are detected, the SELEX round should be checked for the contamination and repeated.

Figure 4-1: (A) Real-time PCR curves for the samples having sufficient (a) and insufficient (b) ssDNA sequences for amplification. (B) Polymerase chain reaction product verification in agarose gel electrophoresis. The single band around 100 bp in the right lane is the PCR product after a successful SELEX round.

4.1.1.3 Double-stranded DNA to ssDNA conversion

To convert the dsDNA produced by the real-time PCR to ssDNA, the asymmetric

PCR method was employed. As mentioned in section 2.2.3, among all other techniques, the asymmetric PCR is the most cost-effective and least time-consuming. The final product of asymmetric PCR is the mixture of ssDNA and dsDNA, which can either be purified or can be directly applied to the next round of SELEX. In the library immobilization SELEX, purifying the asymmetric PCR product is not necessary as dsDNA is eventually discarded 77 during the library immobilization to the complementary capture probes. In this study, only the forward primer was used to obtain the unidirectional asymmetric amplification. After the amplification, agarose gel electrophoresis was performed to verify the production of ssDNA. As can be seen in Figure 4-2, there were two separate bands in the right lane representing the dsDNA (upper band) and the ssDNA (lower band), which verified the production of ssDNA in the asymmetric PCR method. The final product was then applied for the next round for SELEX without purification.

Figure 4-2: Agarose gel electrophoresis of the asymmetric PCR product. Two separate bands in the right lane represent the dsDNA and ssDNA.

4.1.1.4 Transformation, cloning, and sequencing

Figure 4-3 illustrates the transformation and cloning process. Overall, the process includes taking a DNA fragment, inserting it into a circular vector, introducing a recombinant vector into a host cell, and replicating into millions of copies through both the

78 multiplication of cells and recombinant vector [251]. In this study, a linearized pCRTM 2.1-

TOPO® vector with a single 3’-thymidine (T) overhang was utilized for the cloning. The linearized vector remains activated due to the covalently bound topoisomerase I until a

DNA fragment is inserted into it. A map of this vector is shown in Figure 4-4. The TOPO® vector contains three main genes of interest that are useful for screening the successful ligation and electroporation. The essential genes are ampicillin resistance, kanamycin resistance, and LacZα. When the adenylated final PCR product is mixed with the linearized

TOPO® vector, a DNA fragment is ligated, leading to the formation of a circular recombinant plasmid vector. The circular plasmid is then inserted into the host cells, electrocompetent E. Coli in this study, through the electroporation process. For the electroporation, the circular plasmids are mixed with the host cells, and a high voltage pulse is applied. The high voltage pulse causes temporary openings within the cell membrane, thus allowing the circular plasmid to enter inside the cell. The cells are then called genetically transformed. Insertion of the plasmid into the cells increases the metabolic load compared to the other cells without the plasmid, which is the disadvantage to the transformed cells as they will be out-competed while growing. In order to grow the genetically transformed cells, a selective pressure must be applied so that the small number of transformed cells will become the majority of the population on the LB plates.

Figure 4-3: Schematic illustration of the transformation and cloning process.

The first selective pressure exploited in this process was the kanamycin resistance.

The transformed cells were grown in an antibiotic, kanamycin, containing LB media plates.

As the plasmid contains the gene for kanamycin resistance, the cells without the inserted

TOPO® vector die, and only the cells with the inserted plasmid survive.

Figure 4-4: The map of pCRTM 2.1 TOPO® vector displaying the sequences surrounding the TOPO® Cloning site [252].

Sometimes, during the ligation process, the linearized vector may also close without an insertion. The host cell transformed with the vector without an insert will still be resistant to the antibiotics. This will lead to the growth of the cells without an actual inset.

To solve this problem, the TOPO cloning kit also provides the blue-white screen technology that allows the detection of successful ligation of DNA fragments into the vector. The gene responsible for this mechanism is LacZα. As can be seen in Figure 4-4, the insertion site of the DNA fragment lies in the middle of the LacZα gene, which codes

81 for a portion of the enzyme β-galactosidase (146 first amino acids). The host E. Coli strains contain lacZΔ15 deletion mutation. When the plasmid vector is inserted into such cells, due to α-complementation, functional β-galactosidase is produced. The enzyme β- galactosidase can hydrolyze 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside (X-gal), resulting in an insoluble blue compound. When the plasmid vector containing the ligated foreign DNA is taken up by the host cells, α-complementation does not occur; therefore, a functional β-galactosidase enzyme is not produced. Thus, if the transformed cells are grown in the presence of X-gal and antibiotic, cells containing the vector without an insert will be blue, and the cells comprising the vector with an actual insert will develop as white colonies [253]. Figure 4-5 depicts the blue-white colonies for the visual confirmation of transformation and cloning from this study.

Figure 4-5: A portion of a petri dish containing the transformed bacterial colonies with blue-white screening. White colonies are the bacterial cells containing the vector with an aptamer insert, whereas the blue colonies are the host cells that have taken up the vector without an insert.

For sequencing, the white colonies were transferred in fresh LB plates and shipped in ice overnight to Functional Biosciences. The sequencing results from the Functional

Biosciences can be obtained in two business days. The results consist of an inserted aptamer fragment flanked by the sequences from the TOPO® vector. A typical example is

82 shown in Figure 4-6. The potential aptamer sequence within the provided sequencing results can be identified by searching the common regions (primer binding sites) within the initial ssDNA library (5’-GGTGACTGCTACTGTGTTGG-N44-

CCACACATCCAAGCAGAACC-3’). If the sequence in between these common regions

(5’-GGTGACTGCTACTGTGTTGG-3’ and 5’-CCACACATCCAAGCAGAACC-3’) has 44 bases, that will be a potential aptamer sequence. nnnnggttcttgggcgatgggccctctgatgcatgctcgagcggccgccagtgtgatggatatctgcagaattcgcccttggtgactgctact gtgttggaccgggtgaggggggtccagtgttagtagcgatggaggggtgacccacacatccaagcagaaccaagggcgaattccagcac actggcggccgttactagtggatccgagctcggtaccaagcttggcgtaatcatggtcatagctgtttcctgtgtgaaattgttatccgctcaca attccacacaacatacgagccggaagcataaagtgtaaagcctggggtgcctaatgagtgagctaactcacattaattgcgttgcgctcactg cccgctttccagtcgggaaacctgtcgtgccagctgcattaatgaatcggccaacgcgcggggagaggcggtttgcgtattgggcgctcttc cgcttcctcgctcactgactcgctgcgctcggtcgttcggctgcggcgagcggtatcagctcactcaaaggcggtaatacggttatccacaga atcaggggataacgcaggaaagaacatgtgagcaaaaggccagcaaaaggccaggaaccgtaaaaaggccgcgttgctggcgtttttcc ataggctccgcccccctgacgagcatcacaaaaatcgacgctcgagtcagaggtggcgaaacccgacaggactataaagataccaggcgt ttccccctggaagctccctcgtgcgctctcctgttccgaccctgtcgcttaccggatacctgtccgcctctctcccttcgggaagcgtggcgctt tctcatagctcacgctgtacgtatctcagttcggtgtaggtcgttcgctccnagctgggctgtgtgcacgaactcatcggtcagctcgaccgct gcgtcttatccgggtactatcgtcttgagtcaatcgcggtagacacgacttatcgnctctggcagcagcgactggtaacaggatttacagagc gatgtatgtaggcgctgctacagagttcttgagtggatggnctactgacgctacactagagaacagtattggcatctgcactctgctgagcca gttactttcgaaaangantcgtagctctgatccggcaactaacacgctggtancggtgggtttttgnttgcagccccaattaaccctgaaaatg atctcggaaatcctggacttttttagggctgagctcagtgaacaaaatcccctaaggtttggcccgaaatttcaaaggaattcctaatcttctataa aaagaatttatttctccaatattagaaattggtgcggcacacggcttgtgggccccct Figure 4-6: Typical sequencing result from Functional Biosciences with the potential aptamer inserted into the TOPO® vector. The common start and end regions are highlighted in red and bright blue, respectively. The aptamer sequence is between the start and end sequences (highlighted in pink).

The ligation into the TOPO® vector, however, may not always be in that order (5’-

GGTGACTGCTACTGTGTTGG-N44-CCACACATCCAAGCAGAACC-3’). It is also possible for the DNA fragment to be inserted in reverse (5’-

GGTTCTGCTTGGATGTGTGG-N44-CCAACACAGTAGCAGTCACC-3’), which is the complementary sequence to the initial library sequence. Due to the single 3’ adenine overhang in the DNA fragment, it can insert in either way. In this case, the common regions will be the 5’-GGTTCTGCTTGGATGTGTGG-3’ and CCAACACAGTAGCAGTCACC-

3’. The reverse complement of the sequence in between these sequences will be a potential aptamer. An example of such insertion is shown in Figure 4-7. nnnggttcttgggcgattgggccctctgatgcatgctcgagcggccgccagtgtgatggatatctgcagaattcgcccttggttctgcttggat gtgtggggtgtcacccacgctgagtcaccgccagttagtctcccagtcccccaacacagtagcagtcaccaagggcgaattccagcacact ggcggccgttactagtggatccgagctcggtaccaagcttggcgtaatcatggtcatagctgtttcctgtgtgaaattgttatccgctcacaattc cacacaacatacgagccggaagcataaagtgtaaagcctggggtgcctaatgagtgagctaactcacattaattgcgttgcgctcactgccc gctttccagtcgggaaacctgtcgtgccagctgcattaatgaatcggccaacgcgcggggagaggcggtttgcgtattgggcgctcttccgc ttcctcgctcactgactcgctgcgctcggtcgttcggctgcggcgagcggtatcagctcactcaaaggcggtaatacggttatccacagaatc aggggataacgcaggaaagaacatgtgagcaaaaggccagcaaaaggccaggaaccgtaaaaaggccgcgttgctggcgtttttccata ggctccgcccccctgacgagcatcacaaaaatcgacgctcaagtcagaggtggcgaaacccgacaggactataaagataccaggcgtttc cccctggaagctccctcgtgcgctctcctgttccgaccctgccgcttaccggatacctgtccgcctttctcccttcgggaagcgtggcgctttct catagctcacgctgtangtatctcagttcggtgtangtcgttcgctccaagctgggctgtgtgcacgaaccccccgttcagcccgaccgctgc gccttattcggtaactatcgtcttgagtccaacccggntagacacgacttatcgccactggcagcagccactggttacaggnatagcagagc gaggtatgtaagcggtgcttcagagttcttgaagtgtgggctaactaccgctaccctagaagaacaggatttggaatctggctctgctgaagc aggtaccttcggaaaagatttgtagctcttgatccggaaacaaacaccgctgtaccggtggtttttttttgcaacaccaattaccgccgaaaaaa gatttcaaagatctttgatttttttcggggctgaactcgggaacgaaactcgttaggatttgggctgaatttcaaagagttcccaatccttttattaa agagtttaaccctaagaatgtggaatgggctgagtccaggtaacggggccttcccgtgtt Figure 4-7: A representative sequencing result with the reverse insertion of the aptamer sequence into the TOPO® vector. The common start and end regions are highlighted in grey and yellow, respectively. The reverse complement of the sequence in between the start and end sequences (highlighted in dark red) is a potential aptamer.

4.1.2 Analysis of aptamer sequences, structural folding, and stability

After 14 rounds of selection, 4 unique aptamer candidates were identified (Table

4.2). The structural folding and stability of the aptamers were predicted using the web- based tool mFold, which predicts the lowest free energy secondary structures, Gibbs free energy change (ΔG), enthalpy change (ΔH), melting temperature (TM), and entropy change

(ΔS) [132]. The parameters of interest in this study were ΔG and TM. Gibbs free energy change provides a measure for the spontaneity of formation and degree of stability. The structures with more negative ΔG value form spontaneously and are more stable. Melting temperature provides insight into whether a structure exists at room temperature. As shown

-1 in Table 4.3, LI-H3-APT-3P had the lowest ΔG value ( -14.86 kcal.mole ) followed by LI-

-1 -1 H3-APT-1P (-13.63 kcal.mole ), LI-H3-APT-2P (-12.24 kcal.mole ), and LI-H3-APT-4P

-1 (-12.17 kcal.mole ). All aptamers exhibited melting temperatures higher than room temperature. The identified sequences exhibited stem and loop structures that are supposed to be critical for specific binding to the target (Figure 4-8). Each of these aptamers exhibited different secondary structures, but all of them contained a consensus region of seven bases

(GGGTGAC). Aptamer LI-H3-APT-3P contained two of these sequences, with one of them missing the base C (GGGTGAG). Interestingly, the consensus region was always on the loops in the secondary structures (Figure 4-8) except for the aptamer LI-H3-APT-4P in which the consensus region was involved in the formation of the stem as well.

Guanine (G)-rich ssDNA aptamers have the propensity to fold into stable tertiary conformation known as G-quadruplex (G4). When two or more planer G-quadrates stack on top of one another, they form a more stable tertiary structure G4. These G4 structures are highly polymorphic and recognize very different protein targets with high affinity [254-

256]. Over the past decade, numerous studies have reported G4 forming aptamers for a wide variety of molecules [257, 258]. In this work, these putative G4 structures of the aptamer candidates were predicted using an online server QGRS mapper. This mapper searches for the occurrence of quadruplex forming G-rich sequences and retains G-score.

The G-score is the likelihood of forming a stable G4; the higher the score, the better the candidate for G4. The highest possible G-score, using the default maximum QGRS length of 30, is 105 [133]. Among 4 candidates, three sequences displayed the propensity to form stable G4 structures except for LI-H3-APT-1P. The aptamer LI-H3-APT-3P had the highest G-score of 51, followed by 31 and 23 for LI-H3-APT-4P and LI-H3-APT-2P, respectively.

Based on these results, the aptamer sequence with the highest G-score and the lowest ΔG value, LI-H3-APT-3P, was chosen for -based capture assay.

The full-length aptamer LI-H3-APT-3P was truncated on both sides to remove the primer binding sites (LI-H3-APT-3) and evaluated (Table 4.3). In QGRS mapper analysis, the truncated sequence retained the same G-score as its parent sequence, suggesting the involvement of only the random region in the formation of the G4 structure. Removal of the primer binding regions increased the ΔG value and decreased the melting temperature of the sequence.

Table 4.2: Aptamer sequences obtained from 14 rounds of SELEX for H3. The primer binding regions are underlined.

Figure 4-8: Predicted secondary structures of aptamer LI-H3-APT-1P, LI-H3-APT-2P, LI- H3-APT-3P, LI-H3-APT-4P, and LI-H3-APT-3 using mFold online program based on a free energy minimization algorithm at 137 mM [Na+] and 5 mM [Mg+2]. The consensus sequence has been highlighted inside the green box.

Table 4.3: Gibbs free energy change, G-score, and TM of the aptamers.

4.1.3 Demonstration of binding through MB-based affinity capture assay coupled

to SDS-PAGE

In this study, the H3 binding aptamer with the lowest ΔG value and highest G- score, LI-H3-APT-3P, was covalently coupled to magnetic beads for generating a functionalized affinity matrix to capture H3. For the conjugation, amino-modified aptamers were covalently coupled to the tosylactivated MBs. In order to minimize the unspecific binding, the unoccupied amine binding sites of the MBs were blocked with BSA. The samples containing H3 and H8 were tested to evaluate the binding affinity and specificity of the aptamer, respectively. Captured peptide molecules were eluted and run into SDS-

PAGE to monitor binding visually.

Figure 4-9 shows the gel image from the SDS-PAGE study. The 1st lane from left is a reference size ladder, where the band for H3 lies roughly around 10 kDa. The 2nd lane is the control sample containing H3 before incubation with aptamer modified MBs; as expected, a strong band at ~ 10 kDa can be seen (C). The 3rd (H3) and 4th (H8) lanes are the samples that were eluted from the aptamer-modified MBs incubated with H3 and H8, respectively. As can be seen in lane 3, the aptamer MBs were able to effectively capture

H3 as indicated by the dark band at ~ 10 kDa. However, there was no such band for the H8

88 sample (lane 4th) at or below 10 kDa, demonstrating that the aptamer did not capture H8.

As counter-selection was performed with H8, a closely related molecule to H3, the aptamers were expected not to bind with this molecule. The smear around 70 kDa was due to the BSA, which was used to block the unoccupied amine binding sites on the MBs.

Although other aptamers have not yet been tested, this experiment indicated the successful

SELEX process to identify H3 selective aptamers.

Figure 4-9: The SDS-PAGE image showing a size ladder, the control sample (c), and eluents from the H3 and H8 incubated aptamer modified MBs.

4.2 Integration and demonstration of the H3-selective aptamers into

ultrasensitive assay formats to allow for the identification and

quantification in complex media

This section presents the results and discussions related to the implementation of the aptamers on various assay platforms such as ELASA, AuNPs-based colorimetric assays, SERS-based magnetic aptasensor, and lateral flow test strip.

4.2.1 Development of ELASA for the additional investigation into the binding

between the selected aptamer candidates and H3

In this subsection, a direct format ELASA was developed to perform the additional characterizations of the aptamer candidates. Based on the computational studies in section

4.1.2, two different aptamers, LI-H3-APT-3P and LI-H3-APT-4P, were chosen for the comparative binding affinity testing. For the most promising candidate, the truncation studies were also performed. The aptamer LI-H3-APT-3P was truncated on both sides (LI-

H3-APT-3) to remove the primer binding regions and tested. Further studies were conducted on the best performing candidate.

4.2.1.1 Binding analysis using ELASA

As explained in section 3.2.1.2, relative binding affinity and cross-reactivity of selected aptamer candidates were assessed using ELASA. As depicted in Figure 4-10, among three aptamers (LI-H3-APT-3P, LI-H3-APT-4P, and LI-H3-APT-3), the truncated sequence, LI-H3-APT-3, had higher affinity compared to the other two full-length aptamers. The full-length version of the truncated sequence, LI-H3-APT-3P, demonstrated intermediate binding followed by another full-length sequence LI-H3-APT-4P. The very

90 high affinity of the truncated sequence indicates that the presence of flanking regions strongly inhibits binding. This reduced binding is thought to emerge from steric hindrance imposed by the stems or loops that are formed by the primer binding constant regions [259].

After removing the 3’ primer binding site of the LI-H3-APT-3P, one of the consensus regions was open; however, there was another conserved region on the second loop, which may have acted as the binding motif.

The binding results were consistent with the G-scores. Comparing the two full- length aptamers, the sequence with higher G-score had a higher affinity, and, in the case of a truncated sequence, increased affinity can be contributed to the reduced steric hindrance after removing flanking regions on both sides. In combination with mFold analysis, QGRS mapper proved to be a useful tool for preliminary analysis of aptamer sequences.

m n

LI-H3-APT-3P

0 5

4 2 LI-H3-APT-4P

O LI-H3-APT-3

0 3 8 k H H n la B

Figure 4-10: Relative binding affinity and cross-reactivity of the aptamers. The H3 and H8 sample wells were coated with the respective peptides, whereas the blank samples were deprived of any proteins. All wells were reacted with 100 pmol of respective aptamers.

All aptamers had a significantly higher binding signal to H3 compared to H8 and blank samples that were of similar intensity. From these results, it is evident that none of the aptamers cross-reacted with H8. These results supported the findings of the MB-based affinity capture assay and further validated the SELEX process, including the counter- selection that was executed to get rid of the unwanted sequences from the selected candidate pool. Based on these results, the best performing truncated aptamer sequence LI-

H3-APT-3 was chosen for further characterization.

4.2.1.2 Binding kinetics studies

Using the ELASA assay, the time required to achieve optimal binding between aptamer (LI-H3-APT-3) and immobilized H3 was determined. Results in Figure 4-11 indicate that the aptamer binds to H3 in a time-dependent manner, reaching a plateau after

60 minutes of incubation. Even 5 minutes of incubation was sufficient to demonstrate the binding event. This fast response time is beneficial for the development of the diagnostic assays/sensors for real-time monitoring. Based on these results, the incubation time of 60 minutes was employed for other experiments to obtain the optimal binding in ELASA.

Figure 4-11: Binding kinetics of LI-H3-APT-3 aptamer to H3. Histatin 3 immobilized wells were applied with 100 pmol of LI-H3-APT-3 for various incubation times (0, 5, 30, 60, and 90 min).

4.2.1.3 Determination of Kd

To determine the , several concentrations (10 nM – 10000 nM) of biotinylated aptamer, LI-H3-APT-3, were used while keeping the H3 amount constant (4 µg/well) as explained in section 3.2.1.4. The data were analyzed using non-linear regression analysis based on the one-site binding model, 푌 = 퐵푚푎푥 × 푋/(퐾푑 + 푋), where 퐵푚푎푥 is the maximum specific binding, and 퐾푑 is the concentration of ligand needed to achieve a half- maximum binding at equilibrium. The approximated 퐾푑 value for LI-H3-APT-3 thorough

ELASA was 1.97 ± 0.48 µM, and the regression coefficient (R2) associated with this model was 0.9604. Figure 4-12 shows the corresponding curve generated using the one-site binding model. This aptamer had slightly better 퐾푑 value compared to the aptamers developed by using similar selection techniques. The best 퐾푑 values for tobramycin,

93 kanamycin, and cortisol aptamer generated using a similar method were 16 µM [246], 16.1

µM [247], and 3.9 µM [99], respectively.

Figure 4-12: Equilibrium dissociation curve for the aptamer LI-H3-APT-3. Amine-binding wells were coated with H3, and a concentration series in a range of 10–10000 nM of biotinylated aptamer LI-H3-APT-3 was applied for binding. The data were fitted to a one-site specific binding model.

4.2.2 Development of AuNPs-based colorimetric assays based on the structural

feature of the identified H3 aptamers

As mentioned in section 3.2.2, two different formats of colorimetric assays were developed by combining AuNPs and the H3 specific aptamer. The structural feature of the aptamer (i.e., conformational change when binding to H3) was exploited in these assays.

4.2.2.1 Assay format-1

4.2.2.1.1 Principle of the colorimetric detection

Schematic representation of the colorimetric assay format-1 is shown in Figure 4-

13. The AuNPs were stabilized in a sodium citrate solution by the adsorbed citrate anions

94 on the surface. The surface adsorbed negative ions exerted a strong repulsive force preventing the aggregation caused by van der Waals attraction. In this dispersed state, the color of the AuNP solution appeared red-wine. While adding high salt concentration (NaCl solution), the NaCl molecules neutralized the negative charge of the citrate anions leading to the aggregation. The aggregated AuNPs changed the color from red-wine to blue (Figure

4-13 (1)). Interestingly, randomly coiled ssDNA aptamers adsorb onto the surface of citrate-stabilized AuNPs via coordination interaction between the exposed nitrogen bases of the DNA molecules and AuNPs. The adsorbed ssDNA molecules add the negative charge to the AuNPs and increase their repulsion, which leads to the enhanced stability against NaCl-induced aggregation [183]. Thus, in the presence of aptamers, the AuNP solution remained red when challenged by the high-salt conditions (Figure 4-13 (3)).

However, when the target molecule was introduced to the aptamer-stabilized AuNPs, the aptamer underwent a conformational change from a random coil structure to a more rigid tertiary structure like G-quadruplex forming an aptamer-target complex. When the aptamers transform into the rigid tertiary structures, they cannot physically absorb on the surface of AuNPs and thus lose the ability to protect the salt-induced aggregation [260,

261]. Hence, in the presence of the target molecule H3, the addition of NaCl solution led to the aggregation of the AuNPs (Figure 4-13 (4)).

Figure 4-13: Schematic illustration of the colorimetric assay format-1 using H3 binding aptamer and AuNPs. Bare citrate-stabilized AuNPs aggregate (color of the solution changes from red to blue) in the presence of salt (1). After mixing aptamers with the AuNPs, aptamers adsorb on the surface of AuNPs and prevent salt-induced aggregation (3). In the presence of H3, the aptamers dissociate from the surface and bind to the H3, resulting in salt-induced aggregation of AuNPs (4).

4.2.2.1.2 Spectral characteristics

In order to investigate the spectral characteristics of the assay mechanism, AuNP solutions with different experimental conditions were prepared. Figure 4-14 displays UV-

Vis spectra and the corresponding photographs of AuNP solutions in different conditions.

The citrate-stabilized AuNPs of 15 nm diameter exhibited a surface plasmon resonance

(SPR) absorbance peak around 520 nm and red color (Figure 4-14 (a )). Upon addition of

NaCl, the absorbance peak around 520 nm decreased, and a new peak at about 690 nm appeared, which was accompanied by the color shift from red to blue representing the aggregation of AuNPs (Figure 4-14 (b)). This phenomenon could simply be monitored by

96 a UV-Vis spectrophotometer or observed by the naked eye. While in the presence of aptamer, the peak at 520 nm was intact, and the AuNP solution was still red, indicating that the added salt was unable to induce the aggregation (Figure 4-14 (c)). In this case, the randomly coiled ssDNA aptamers were wrapped around the AuNPs and protected the

AuNPs from salt-induced aggregation. The slightly decreased peak at 520 nm can be attributed to the effect of TE buffer that the aptamers were resuspended. Slower kinetics of the aptamer adsorption compared to the interaction between the ions of TE buffer and citrate anion can cause such effect. Moreover, upon addition of H3 to AuNP-aptamer mixture, it was observed that the SPR peak at 520 nm decreased, and a new peak at 640 nm appeared depicting the aggregation of AuNPs in the presence of H3. Correspondingly it can be seen that the color of the solution changed from red to purple (Figure 4-14 (d)).

Thus, binding of aptamer with H3 transformed random coil aptamer structure to G- quadruplex; this structure could not adsorb on the AuNPs and lost the capability to protect the AuNPs from salt-induced aggregation.

Figure 4-14: Ultraviolet-Visible absorption spectra of (a) AuNPs, (b) AuNPs with NaCl, (c) AuNPs containing aptamer and NaCl, and (d) AuNPs with aptamer, H3, and NaCl. (Note: 200 µl of 2.6 nM AuNPs, 1.03 µM aptamer, 10 µl of 5 µg/ml H3, and 41.18 mM NaCl).

Further characterization in the morphology of AuNPs in these conditions was performed by using an FEI Tecnai G2 Spirit transmission electron microscope (TEM; FEI

Company, Hillsboro, OR) at an operating voltage of 80 kV. As depicted in Figure 4-15

(A), the citrate-stabilized AuNPs were in a monodispersed state, and following the addition of NaCl, the AuNPs aggregated as represented by the large clump in Figure 4-15 (B). The presence of aptamer in the AuNP solution prevented the salt-induced aggregation (Figure

4-15 (C)). Most of the AuNPs were in a dispersed state except for a couple of dimers, which may be due to the interaction of the TE buffer ions with citrate anions on the AuNP surface.

The observed dimers also explain the slightly decreased absorbance at 520 nm in UV-Vis spectra for AuNPs and aptamer mixture in Figure 4-14. Furthermore, the addition of target molecule H3 to aptamer-stabilized AuNPs resulted in the aggregation (Figure 4-15 (D)).

All these results were in good agreement with our assumption that the aptamer binding to

H3 induced a conformational change, leading to the salt-induced aggregation of AuNPs.

Figure 4-15: Transmission electron microscopy images of (A) citrate-stabilized AuNPs, (B) AuNPs in the presence of NaCl, (C) aptamer adsorbed AuNPs in NaCl, and (D) aptamer adsorbed AuNPs in the presence of H3 and NaCl. (Note: 200 µl of 2.6 nM AuNPs, 1.03 µM aptamer, 10 µl of 5 µg/ml H3, and 41.18 mM NaCl).

4.2.2.1.3 Optimization of experimental conditions

Various parameters, such as the concentration of salt, concentration of aptamer, and incubation time, have a significant effect on the sensitivity of the colorimetric assay [189,

260, 261]. Hence, a series of experimental conditions were optimized to obtain the optimal assay conditions. The absorbance ratio between A640 and A520 was used to evaluate the degree of aggregation, where a lower value indicates a lesser degree, and a higher value depicts a greater degree of aggregation.

4.2.2.1.3.1 Effect of NaCl concentration

Firstly, the effect of NaCl concentration on the citrate-stabilized AuNPs was studied over the concentration range of 20.66-123.96 mM NaCl. To perform this experiment, 200 µl of 2.6 nM AuNP samples were taken into the 96-well plate, in which

10 µl of TE buffer and 10 µl of H3 BB were added to compensate the salt content of aptamer solution and the H3 BB (that will later contain the target molecule), respectively.

After that, the NaCl solution was added to each sample so that the final concertation was

0, 20.66, 41.32, 61.98, 82.644, 103.3, and 123.96 mM, respectively. The total volume of each sample was 242 µl. Immediately followed by the NaCl addition, UV-Vis spectra were recorded, and photos were taken. As can be seen in Figure 4-16, the AuNPs started aggregating from the 20.66 mM of NaCl as represented by the increase in the absorbance ratio (A640/A520), which can also be observed by the corresponding color change in the sample (Figure 4-16 (inset)). For further experiments, the slightly lower concentration of

NaCl (41.32 mM) than the concentration that completely aggregated the AuNPs (60.98 mM) was chosen as the starting salt concentration.

100

Figure 4-16: Effect of NaCl concentration on the aggregation of the citrate-stabilized AuNPs. The absorbance ratio of A640/A520 vs. the NaCl concentration. (Note: 200 µl of 2.6 nM AuNPs, 10 µl TE buffer, 10 µl BB, and various concentrations (0, 20.66, 41.32, 61.98, 82.644, 103.3, and 123.96 mM) of NaCl.

4.2.2.1.3.2 Effect of aptamer concentration on citrate-stabilized AuNPs

Besides the NaCl concentration, the aptamer concentration also affects the analytical performance of an assay. To optimize the aptamer concentration, the amount of aptamer required to prevent the aggregation induced by the 41.18 mM NaCl was determined first. For this, 200 µl of 2.6 nM AuNP solutions were incubated with 10 µl of

0, 6.25, 12.5, 25, 50, and 100 µM aptamer samples in TE buffer, which made final concentration of aptamer in AuNP solutions as 0, 0.26, 0.52, 1.03, 2.06, and 4.13 µM, respectively. After incubating for 1 hour at room temperature, 10 µl of BB was mixed to compensate for the ionic concentration of the target solution. Afterward, 22 µl of 453 mM

NaCl (41.32 mM) was added to each sample. As can be seen in Figure 4-17 (A), the aptamer concentrations below 2.06 µM were unable to prevent the aggregation as

101 represented by the increasing absorbance ratio (A640/A520) and corresponding color change from purple to blue with decreasing concentration of aptamer. At 2.06 µM and 4.12 µM, the AuNPs were entirely protected by the aptamers, which was indicated by the saturated absorbance ratio and red color of the samples.

Moreover, additional NaCl was added to each sample to observe their response

(Figure 4-17 (B)). The samples with 2.06 and 4.13 µM aptamer concentration did not experience any further change in absorbance ratio, whereas, for other samples, the absorbance ratio increased with increasing salt concentration. The excessive number of aptamers in the assay reduces the sensitivity as free aptamers in the mixture compete with the adsorbed aptamers for the target binding [262]. Therefore, 1.03 µM aptamer concentration (molar ratio of AuNPs to aptamer 1:480) was chosen as the appropriate concentration for sensitive assay development.

102

Figure 4-17: (A) Effect of the aptamer concentration on the absorbance ratio (A640/A520) of AuNPs in the presence of 41.32 mM NaCl (Inset: color change of the AuNP solutions corresponding to 0, 0.13, 0.26, 0.52, 1.03, and 2.06 µM aptamer concentration from left to right). (B) Effect of higher salt concentrations on the samples containing AuNPs and 0, 0.26, 0.52, 1.03, 2.06, and 4.13 µM aptamer.

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4.2.2.1.3.3 Effect of incubation time

Two different incubation times, incubation of aptamers with AuNPs and incubation of target molecule with aptamer-protected AuNPs, were studied. At first, the reaction time between the aptamer and AuNPs was investigated to identify the minimal time required to obtain the optimal binding between the aptamer and AuNPs. It was performed by incubating 10 µl of 25 µM aptamer with 200 µl of 2.6 nM AuNP solutions for various time periods (0.167, 0.5, 1, 2, 4, and 8 hours). Following the incubation, 10 µl of BB and 22 µl of 453 mM NaCl were added, and the absorbance ratio (A640/A520) was calculated. The resultant graph in Figure 4-18 shows that the amount of aggregation was dependent on the reaction time. When the incubation time was less than 4-hour, the binding between the aptamer and AuNPs was not strong enough to completely protect the AuNPs against salt- induced aggregation as represented by the higher absorbance ratio for lesser incubation times. The difference between the ratio was unapparent when the reaction time was more than 4 hours, which was probably because the binding between the aptamer and AuNPs was comparatively strong to the binding between the AuNPs and neutralizing ions of the salt. Hence, the incubation time of 4 hours was chosen as the appropriate time.

104

0.40

0.35

5 A

/ 0.30

6 A 0.25

0.20 0 2 4 6 8 10 Incubation Time (hour)

Figure 4-18: Incubation study to optimize the time required for aptamers to stabilize the AuNPs. The absorbance ratio was monitored by adding 10 µl of various aptamer concentrations and 10 µl of BB to 200 µl of AuNPs at 41.18 mM NaCl.

Subsequently, the H3 incubation time was also studied to identify the optimal time required for the aptamer to bind to the target in the defined assay conditions. To carry out this experiment, 200 µl of AuNP solutions were hatched with 10 µl of 25 µM aptamer for

4 hours at room temperature. Then, 10 µl of 5 µg/ml H3 in BB was added to the aptamer- stabilized AuNP solutions and incubated for various time periods (5, 15, 30, and 60 min).

Following this incubation, 22 µl of 453 mM NaCl was supplemented, and the results were recorded. As shown in Figure 4-19, the aggregation gradually increased with increasing incubation time and reached a plateau after 30 minutes. When the incubation time was less than 30 minutes, the target did not have enough time to dissociate aptamer from AuNPs surface; therefore, the color of the solution remained red, and consequently, the absorbance ratio was small. For the incubation time of 30 minutes or more, H3 was bound with more aptamers leading to the desorption of a large number of aptamers from AuNPs surface,

105 increasing the aggregation (higher absorption ratio). Based on these results, an incubation time of 30 minutes was chosen as the optimum time for further experiments.

0.45

0.40

A /

0 0.35

6 A 0.30

0.25 0 20 40 60 80 Incubation Time (min)

Figure 4-19: Effect of H3 incubation time on the absorbance ratio. The signal was obtained by using 200 µl of AuNPs, 10 µl of aptamer solution, 10 µl of 5 µg/ml H3 in BB, and 41.18 mM NaCl.

4.2.2.1.4 Calibration modeling for H3

As explained in section 3.2.2.1, a series of different concentrations of H3 (0, 1.5,

2.5, 5, 10, 20, 40, and 80 µg/ml) was tested, and their UV-Vis spectra were recorded. As expected, in the presence of H3, the aptamers captured H3 to form aptamer-H3 complexes, resulting in different extents of AuNPs aggregation. As shown in Figure 4-20 (A), with increasing concentration of H3, the absorbance at 520 nm decreased, and the absorbance at 640 nm increased. Also, the absorption ratio (A640/A520) gradually increased with increasing concertation of the H3 up to 80 µg/ml (Figure 4-20 (B)). Moreover, this ratio was linear in the concentration range of 1.5-20 µg/ml (Figure 4-20 (C)), which clearly included the normal range of H3 in human saliva, 1.7-11.8 µg/ml [263]. The regression

106 equation associated with the linear region was A640/A520 = 0.0279 * concentration of H3 in

µg/ml + 0.2384, with a correlation coefficient (R) of 0.993. The detection limit (퐷퐿) was calculated to be 0.199 µg/ml by using 퐷퐿 = 3푆퐷퐵/푚, where 푆퐷퐵 is the standard deviation of the blank sample and 푚 is the slope of the calibration curve.

Figure 4-20: Detection of H3 using AuNPs-based colorimetric assay format-1. (A) Ultraviolet-visible absorption spectra following incubation with various H3 concentrations. (B) Absorbance ratio (A640/A520) as a function of H3 concentration: concentrations were 0, 1.5, 2.5, 5, 10, 20, 40, and 80 µg/ml H3. (C) Calibration curve for H3 in the 0-20 µg/ml range.

107

4.2.2.1.5 Selectivity

To examine the selectivity of this assay, response to 5 µg/ml H8 was obtained and compared with the 5 µg/ml H3 under the optimized conditions. As shown in Figure 4-21, only the addition of H3 resulted in an apparent change in UV-Vis absorption spectra or the absorbance ratio, whereas no difference was observed for H8 and blank. All these results further validated the outcomes from the MB-based capture assay and ELASA assay and demonstrated the selectivity of the assay format-1 to the target molecule H3.

0.5

0.4

0 2

5 0.3

0 4

6 0.2 A

0.1

0.0 H3 H8 Blank

Figure 4-21: Selectivity of the assay format-1 against H8. Absorbance ratio was obtained by testing 5 µg/ml of H3 and H8. For blank sample, BB was used.

4.2.2.2 Assay format-2

4.2.2.2.1 Principle of the colorimetric assay

The overall strategy for this assay format-2 is also similar to the format-1. The major differences are the order at which the components are mixed, and the time taken to complete the assay. In assay format-1, the aptamer and AuNPs are mixed first, and then the target molecule is introduced. The entire assay procedure using the optimized

108 parameters takes about 4 hours and 30 minutes. However, in assay format-2, the aptamer and target molecules are mixed first, and after letting the binding happen, the citrate- stabilized AuNP solution is added. The binding between the target molecule and aptamer happens in the absence of any external competitor. As depicted in Figure 4-22, in the absence of H3, the aptamers adsorb onto the surface of AuNPs and increase the repulsive force, thus preventing the aggregation when challenged by the high salt conditions.

However, if the sample contains H3, the aptamers undergo a conformational change and form the aptamer-H3 complex. Due to the rigid three-dimensional structure, the aptamer-

H3 complex cannot adsorb on the surface of AuNPs and, when challenged by the salt, the

AuNPs aggregate. In this format, the amount of aptamer, NaCl, and AuNPs were the same as assay format-1. Only the incubation times were reduced to 5 minutes to reduce the assay time.

109

Figure 4-22: Schematic illustration of the colorimetric assay format-2. Aptamers and the target molecules are mixed first, and then AuNPs are added. In the absence of the target, the aptamers protect citrate-stabilized AuNPs against salt-induced aggregation, and the solution remains red. In the presence of H3, the aptamers change conformation and form aptamer-H3 complexes, which cannot protect the salt-induced aggregation, and the solution changes color to blue.

4.2.2.2.2 Calibration modeling for H3

Under the defined conditions (section 3.2.2.2), the response of the assay format-2 to various concentrations of H3 (0, 1.25, 2.5, 5, 10, 20, 40, 80, 160 µg/ml) was investigated.

As shown in Figure 4-23 (A), with an increase in H3 concentration, the broad absorbance band at 640 nm gradually increased. Similar to the format-1, the absorbance ratio of

A640/A520 was employed to measure the assay response quantitatively. Figure 4-23 (B) shows that the absorbance ratio exhibits a dynamic range of the assay from 0-160 µg/ml

H3. A linear relationship was found between 0 to 20 µg/ml (Figure 4-23 (C)), which was similar to the assay format-1. The calibration equation for the linear range was 푦 =

0.01794푥 + 0.1925, where 푦 is the ratio of A640/A520 and 푥 is the H3 concentration in

110

µg/ml. The R value associated with this model was 0.97. The 퐷퐿 calculated by using 퐷퐿 =

3푆퐷퐵/푚 was 0.505 µg/ml. Comparing to format-1, the slope of the assay was slightly lower, indicating the reduced sensitivity, which can be attributed to the reduced incubation times.

Figure 4-23: Colorimetric detection of H3 using AuNPs-based assay format-2. (A) Ultraviolet-visible absorption spectra following incubation with various concentrations of H3. (B) Absorbance ratio (A640/A520) as a function of H3 concentration: Concentrations were 0, 1.5, 2.5, 5, 10, 20, 40, 80, and 160 µg/ml. (C) Calibration curve for H3 in the 0-20 µg/ml range.

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4.2.2.2.3 Selectivity

Similar to the format -1, the selectivity of assay format-2 against 5 µg/ml H8 was also evaluated. As can be observed in Figure 4-23, upon the addition of H3, there was a noticeable change in the adsorption ratio, while no such effect in the blank or H8 sample occurred. The adsorption ratio value, A640/A520, in the presence of H3 was considerably larger than those of blank or the H8, which were of similar intensity. All results indicate that this assay approach has a specificity to H3.

0.4

0.3

A /

0 0.2

6 A 0.1

0.0 H3 H8 Blank

Figure 4-24: Selectivity of the assay format-2 against H8. Absorbance ratio was obtained by testing 5 µg/ml of H3 and H8. For blank sample, BB was used.

4.2.3 Exploration of SERS-based magnetic aptasensor for the ultrasensitive

detection of H3

4.2.3.1 Principle of the detection method

Figure 4-25 depicts the schematic illustration describing the mechanism of the developed method. The AuNPs modified with 4-NTP, and thiolated H3 aptamers were used as the signal probe (AuNPs-NTP-aptamer). Capture probes, MBs-cDNA, were prepared

112 by immobilizing the biotinylated cDNA on streptavidin functionalized magnetic beads. For the assay, the AuNPs-NTP-aptamer conjugates were first reacted with the H3 samples to let the binding happen between the aptamers and target molecules. In the absence of H3, the aptamers on the AuNPs-NTP-aptamer complex remained unoccupied, and when reacted with MBs-cDNA, the unoccupied aptamers hybridized with the cDNA sequence.

After washing the sample by using an external magnet to pull down the conjugates, the resulting complex, AuNPs-NTP-aptamer_MBs-cDNA displayed high SERS activity.

Conversely, when the target molecules were present, the aptamers on the AuNPs-NTP- aptamer were bound to H3 and were unavailable to hybridize with the MBs-cDNA.

Following the magnetic separation, the occupied signal probes were discarded, and the resultant complex displayed depleted SERS intensity. The decrease in the SERS signal was proportional to the concentration of the H3 in the sample.

Figure 4-25: Schematic illustration of the SERS-based magnetic aptasensor for the detection of H3. In the absence of H3, the aptamers on the AuNPs-NTP- aptamer conjugate remain unoccupied and can bind to the MBs-cDNA, resulting in high SERS intensity. Whenever H3 is present, the aptamers on the signal probe are occupied and cannot hybridize to the capture probe. The resulting complex displays lower SERS intensity.

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4.2.3.2 Characterization and optimization of the signal probe

In this study, 4-NTP was chosen as the Raman reporter molecule. Thiol functional group of 4-NTP facilitate the binding to the SERS substrate (AuNPs) via Au-S covalent bond. Due to the nitro functional group and aromatic ring structure, the Raman scattering cross-section of this molecule is relatively large, and at certain excitation wavelengths, resonance Raman scattering can be obtained [264]. To verify the functionalization of

AuNPs with 4-NTP, Raman and SERS scans were performed. Initially, a Raman scan of

4-NTP powder was obtained, and as depicted in Figure 4-26 (A), three sharp characteristic vibrational peaks at 1101, 1334, and 1599 cm-1 were observed. After that, SERS spectra for 2 µM 4-NTP immobilized on AuNPs was acquired. Similar to the 4-NTP powder, the

SERS scan of AuNPs-NTP had three peaks; however, the peaks were at slightly different positions: 1079, 1328, and 1569 cm-1 (Figure 4-26 (B)) corresponding to C-S, N-O, and C-

C stretching vibrations [265, 266]. Out of the three peaks, the most prominent one at 1328 cm-1 was considered for the assay development. After knowing the Raman active peaks,

Raman scan of 2 µM 4-NTP dissolved in ethanol was taken to see if that concentration of

4-NTP gives off a detectable signal. As shown in Figure 4-26 (B), there was no detectable signal from 2 µM 4-NTP solution, verifying that the 4-NTP peaks can be obtained only when it is conjugated to the AuNPs surface. Also, the Raman signal for AuNPs was tested, and as can be seen in Figure 4-26 (B), no such peaks were observed. All these results verify that the 4-NTP was immobilized on the AuNPs.

The SERS enhancement factor (퐸퐹) was calculated by using the equation 퐸퐹 =

(퐼푆퐸푅푆 × 퐶푅푎푚푎푛)/(퐼푅푎푚푎푛 × 퐶푆퐸푅푆) [267], where 퐶푆퐸푅푆 and 퐶푅푎푚푎푛 are the concentrations of the Raman reporter molecule for SERS and normal Raman 114 measurements, respectively, and 퐼푆퐸푅푆 and 퐼푅푎푚푎푛 are the corresponding SERS and normal intensities. To calculate the EF, Raman spectra of 5 mM 4-NTP dissolved in ethanol was used to compare with the SERS intensity of 2 µM 4-NTP modified AuNPs. Through the calculation, EF was found to be 105.

Figure 4-26: (A) Raman spectrum of 4-NTP powder, and (B) SERS spectrum of 2 µM 4- NTP conjugated AuNPs, and Raman spectra of AuNPs and 2 µM 4-NTP dissolved in ethanol showing no detectable signal.

The amount of Raman active molecule immobilized on the AuNPs affects the sensitivity and signal intensity of the SERS aptasensor [268, 269]. In order to optimize the

4-NTP concentration, various amounts (0, 1, 4, 10, 20, and 40 µl) of 100 µM 4-NTP were added to the 200 µl of AuNPs so that the final concentration was 0, 0.5, 2, 5, 10, and 20

µM in the AuNP solution. The samples were sonicated for 40 minutes to enhance the conjugation, and then the particles were washed by centrifuging (13000 rpm/15 minutes) and resuspending the precipitate in HEPES buffer. Next, the washed AuNPs-NTP conjugates were redispersed in 20 µl of HEPES and 1 µl was transferred on the glass slide to obtain the SERS spectra. As can be seen in Figure 4-27, SERS intensity increased with increasing concentration of 4-NTP until 5 µM and then decreased for higher

115 concentrations. Similar trends have been reported in other studies for functionalizing nanoparticles with Raman molecules [268-270]. The reduced intensity at low concentration of 4-NTP was due to the lesser number of reporter molecules immobilized on the AuNPs, while the suppressed intensity at higher concentration was thought to emerge from the aggregation of nanoparticles caused by the very high concentration of 4-NTP [270].

Based on these results, 5 µM 4-NTP concentration would have been optimum; however, the amount of 4-NTP on the surface of AuNPs also determines the quantity of aptamer immobilized, as the remaining surface would be available for aptamer immobilization. The amount of aptamer immobilized determines the stability against the salt-induced aggregation in the buffer mediums and thus eliminates the nonspecific signals arising from the aggregated capture probes. Therefore, all of the above AuNPs-NTP conjugate samples were immobilized with the thiol modified aptamers to test their ability to prevent the salt-induced aggregation. To functionalize with aptamers, the AuNPs-NTP solutions were made 190 µl by adding 171 µl of HEPES buffer so that the final concentration was 2.6 nM, which is the same as the unmodified AuNP solution. Afterward,

12 µl of aptamer solution (50 µM) previously activated with TCEP was added to each

AuNPs-NTP samples and incubated for 24 hours on a shaking platform. Following the 24- hour incubation, 60 mM NaCl was added and further incubated for another 24 hours.

Afterward, the color and the UV-Vis spectra of the samples were monitored to test their aggregation. As shown in Figure 4-28, the sample without 4-NTP had the lowest absorbance ratio (A640/A520), representing the smallest amount of aggregation. The samples with 0.5 µM and 2 µM 4-NTP had similar and slightly higher absorbance ratio than the control (0 µM) sample. However, for other samples including 5 µM, the aggregation

116 increased with increasing concentration of 4-NTP as represented by the higher absorbance ratio and corresponding color change. Based on these results, 2 µM 4-NTP was identified as the optimum concentration for the SERS aptasensor.

2000

y 1500

t n

I 1000

R E

S 500

0 0 5 10 15 20 25 4-NTP Concnetration (M)

Figure 4-27: Surface-enhanced Raman scattering intensity of AuNPs-NTP conjugate at various concentrations of 4-NTP (0, 0.5, 2, 5, 10, and 20 µM).

Figure 4-28: Aptamer immobilization on the AuNPs-NTP conjugates functionalized with various concentrations of 4-NTP. The samples in Figure 4-27 were further functionalized with thiol modified aptamers.

117

Following the optimization of the 4-NTP concentration, thiol modified aptamers were immobilized by the salt-aging method. The conjugation of the aptamer was verified by performing salt-induced aggregation assay and taking UV-Vis measurements. For executing the assay, 200 µl of AuNPs-NTP-aptamer was taken, and 240 mM NaCl was added. As depicted in Figure 4-29, the addition of NaCl did not induce any change in the absorbance spectra and color. These results verified the immobilization of aptamers on the

AuNPs-NTP conjugates.

Figure 4-29: Verification of aptamer immobilization on AuNPs-NTP conjugates. The NaCl solution (240 mM) was added to the prepared AuNPs-NTP-aptamer conjugates (200 µl).

4.2.3.3 Characterization of the capture probe

Biotinylated cDNA sequences were immobilized on the streptavidin modified magnetic beads via biotin-streptavidin bonding, and the functionalization was confirmed by measuring the absorbance of cDNA solution before and after immobilization at 260 nm.

For 200 µl of aptamer solution, the absorbance before immobilization was 0.274, and

118 following the conjugation, the supernatant had the absorbance of 0.187 (Figure 4-30). The decrease in absorbance value was due to the immobilization of DNA sequences on the magnetic beads.

0.3

m 0.2

O 0.1

0.0 n n o o ti ti za za il li b i o ob m m im m i re r o te ef f B A

Figure 4-30: Absorbance of cDNA solution before and after immobilization on MBs at 260 nm.

4.2.3.4 Optimization of the detection parameters

For the aptasensor, the amount of MBs-cDNA was fixed to 2 µl (10 mg/ml), and the quantity of AuNPs-NTP-aptamer required to saturate the MBs-cDNA was identified.

Before mixing, the AuNPs-NTP-aptamer molecules were centrifuged and resuspended in the same volume of BB. To the 2 µl MBs-cDNA, various amounts (10 µl, 20 µl, 40 µl, and

80 µl) of AuNPs-NTP-aptamer were added and reacted for two hours on a shaking platform. The unbound AuNPs-NTP-aptamer conjugates were discarded via washing two times with BB using a magnetic separator. The washed precipitate was then redispersed in

119

2 µl of BB and deposited on the glass slide for SERS measurement. As can be seen in

Figure 4-31, SERS intensity increased corresponding to the higher amounts of AuNPs-

NTP-aptamer and reached saturation for 40 µl. Hence, for 2 µl of MBs-cDNA, 40 µl of as- prepared AuNPs-NTP-aptamer was identified as the optimal amount.

Figure 4-31: Optimization of signal probes for 2 µl of 10 mg/ml capture probe. Various amounts of the signal probe (10 µl, 20 µl, 40 µl, and 80 µl) were reacted with the fixed amount of capture probe.

4.2.3.5 Histatin 3 detection

For the quantitative evaluation, following the deposition of the samples on the glass slide, the spots of the high SERS intensity were identified, and the signals from five identical spots (Figure 4-32) were acquired. The background signals of each spectrum were subtracted, and the average intensity of the most prominent peaks at 1328 cm-1 was considered. Through this technique, the relative standard deviation (RSD) for repetitive measurement was reduced to 15%.

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Figure 4-32: (A) Dried samples on the glass slide and (B) 50x magnified portion with SERS measurement spots of the sample.

To evaluate the feasibility of the SERS aptasensor for detecting H3, two different concentrations of H3 (1 µg/ml and 5 µg/ml) in BB were tested. Figure 4-33 illustrates the obtained data from this experiment. In the absence of H3, the SERS aptasensor has the highest intensity, indicating the maximum binding between the AuNPs-NTP-aptamer and

MBs-cDNA. Conversely, when there was H3 in the sample, the assay signal depleted, corresponding to the increasing concentration. In this case, the H3 molecules occupied the aptamers on the signal probes, and during washing, the occupied molecules were discarded.

The resultant complex had lesser signal probes, which caused the loss in signal intensity.

These results indicate that the proposed aptasensor can differentiate various concentrations of H3.

121

1500

t i

s 1000

R 500

E S

0 l o 3 3 tr H H n l l o /m /m C g g µ µ 1 5

Figure 4-33: Response of the SERS-based magnetic aptasensor to 0, 1, and 5 µg/ml H3 samples.

Although the developed aptasensor was able to discriminate various concentrations of H3, there were some issues that still need attention. Firstly, a relatively large concentration (in µg/ml range) of the target was needed to induce a significant spectral change. Even though it would work in the salivary range of H3 (1.7-11.8 µg/ml) in a healthy individual, a slightly better sensitivity for SERS aptasensor probably in the ng/ml range was expected. The lower assay range is particularly advantageous while working with viscous samples like saliva. It allows dilution of samples, which might sometimes help to reduce the matrix effect. The reason for this assay not to respond to lower H3 concentrations may be due to the adsorption of H3 on the tubes. During the experiments, it was observed that the H3 solution diluted in BB became sticky to the surface of the polypropylene vials. There are various reports about the nonspecific binding of the peptides to the various surfaces, including polypropylene [271]. Therefore, it can be speculated that

122 there might be nonspecific adsorption of H3 to the tube surface. This may not be a problem in some instances; however, while working with small amounts of the peptide, this type of adsorption may reduce the available quantity to significantly low. To lessen the nonspecific interactions and improve the 퐷퐿, various techniques such as adding Tween-20, high salt concentrations, BSA, or coating tubes with polyethylene glycol can be implemented [271].

Additionally, the low protein binding tubes that are commercially available can be used.

Also, the AuNPs tend to aggregate during various steps of the functionalization due to the ion-induced displacement of the citrate-capping. When AuNPs functionalized with

Raman reporter molecule aggregate, they form an area of very high SERS intensity know as hot spots [272]. The presence of such aggregated conjugates in the signal probe impairs the reproducibility of the assay by resulting in the strong SERS spectra for some spots

[273]. Therefore, it is critical to identify the appropriate conditions to precisely control functionalization, resuspension, and washing steps to minimize aggregation. In this study, using ultrapure water, 10 mM PBS, or BB for the washing and redispersion resulted in aggregation. Finally, the use of 5 mM HEPES (pH 7.6) buffer prevented such problems, and the aggregation was minimized. In the case of HEPES, the piperazine moiety acted as a stabilizing agent for the AuNPs [274].

Overall, this study was able to overcome some of the issues and demonstrate the

SERS-based aptasensor for H3 detection. By carefully optimizing some other parameters or changing the format of the assay, a more sensitive SERS-based assay can be designed.

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4.2.4 Demonstration of an aptamer-based lateral flow test strip for on-site detection

of H3

4.2.4.1 Principle of the aptamer-based lateral flow test strip

The principle of the aptamer-based lateral flow test strip was based on the competitive binding reaction between the H3 in the sample and the immobilized complementary DNA sequence, Probe-TL, on the test line for binding to the AuNPs- aptamer conjugates (Figure 4-34). Typically, the sample solution to be tested was applied to the sample pad. Subsequently, the sample migrated by capillary action and rehydrated the AuNPs-aptamer complexes on the conjugate pad. The rehydrated AuNPs-aptamer conjugates reacted with the target molecule in the sample and at the same time migrated forward to the nitrocellulose pad, where the competitive reaction took place at the test zone.

In the absence of H3, the aptamers on the AuNPs were unoccupied and hybridized with the

Probe-TL on the test line and the Probe-CL on the control line via complementary base pairing, resulting in two red lines on the strip. When H3 was present in the sample, aptamers on the AuNPs preferred to bind with the H3, decreasing the AuNPs-aptamer conjugates that could hybridize to the Probe-TL on the test line causing the red color intensity to become weaker. In general, for the higher concentration of H3, the weaker intensity of the test line was expected. The capture of AuNPs-aptamer by the Probe-CL on the control line remained the same regardless of the presence of H3, ensuring the validity of the detection. No visible line at the test line was taken as an invalid test. The excess

AuNPs-aptamer conjugates and the sample solution migrated further and finally reached the absorbent pad. The entire detection process can be achieved within 5 minutes.

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Figure 4-34: Schematic representation of the aptamer-based lateral flow test strip for the detection of H3.

4.2.4.2 Confirmation of aptamer immobilization on AuNPs

To verify the conjugation of thiol modified aptamers to AuNPs, a salt-induced aggregation test was performed. In this test, 100 µl of unmodified AuNPs and the same

125 amount of aptamer-immobilized AuNPs (10 µl of as-prepared AuNPs-aptamer diluted with

90 µl of HEPES buffer) were treated with 240 mM NaCl. As can be seen in Figure 4-35

(A), citrate-stabilized AuNPs aggregated in the presence of salt as represented by the diminished peak at 520 nm and increased absorbance band in the higher wavelength range.

The corresponding color was changed to blue. For the aptamer-immobilized AuNPs, the addition of salt did not cause any change in the color and absorbance spectra, indicating protection by the immobilized aptamers (Figure 4-35 (B)).

126

Figure 4-35: Ultraviolet-Visible spectra of (A) citrate-stabilized AuNPs and (B) aptamer- immobilized AuNPs with and without 240 mM NaCl. Inset: Photographs of corresponding samples with (+NaCl) and without (-NaCl) 240 mM NaCl.

4.2.4.3 Optimization of the performance of the test and control lines

The nitrocellulose membranes are the widely used signal pad materials in lateral flow devices as they support adsorption of the proteins to create the test and control lines

127 and offer a sufficient sample flow rate [244]. In this project, a nitrocellulose membrane specially designed to handle the viscous samples such as saliva or serum was obtained. In order to immobilize the control and test probe DNA sequences on the nitrocellulose membrane, streptavidin was utilized as the bridging molecule. The DNA fragments do not bind to the nitrocellulose membrane and may get washed away if sprayed directly.

Therefore, a large molecular weight protein molecule, streptavidin, which can be immobilized on the nitrocellulose membrane by electrostatic adsorption and can hold the biotin-modified DNA molecules was utilized [244, 245].

To create the test line, two different complementary sequences, Probe-TL-1 and probe-TL-2, were designed. The Probe-TL-1 comprised of 18 bases long sequence complementary to the 5’region (thiol modified end) of the H3 aptamer, whereas the Probe-

TL-2 was complementary to the entire aptamer except for the T overhang segment. Both of the sequences were immobilized on the individual test strip to compare their performance. Initially, experiments were performed on the strips without blocking any components and using 2.5 µl of AuNPs-aptamer conjugates. The running buffer was 150

µl of H3 BB. As can be seen in Figure 4-36, Probe-TL-1 did not capture any AuNPs- aptamer complex, whereas Probe-TL-2 displayed a faint band depicting the weak interaction between the aptamer and the Probe-TL-2. At the same conditions, the control line composed of poly A’s, complementary to the T overhang segment of the aptamer, had a bright red line.

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Figure 4-36: Capture of AuNPs-aptamer complex by Probe-TL-1, Probe-TL-2, and Probe- CL sequences at room temperature. (Note: The conjugate pad was not blocked, and the amount of AuNPs-aptamer complex was 2.5 µl).

The results obtained for the test line were unanticipated, as we were expecting a better affinity of the AuNPs-aptamer to the test line compared to the control line. The A pairs with T by forming 2 hydrogen bonds, and G-C pair with 3 hydrogen bonds [275].

Therefore, in theory, the base pairing between the complementary DNA sequences made out of A, T, G, and C should be stronger (between the aptamer and Probe-TL-1 and Probe-

TL-2) than the hybridization between the oligo containing only A and T (between T overhang and Probe-CL). However, in this experiment, the T overhang region had a better affinity for poly-A bases than the affinity of the remaining sequence to its complementary sequence. To further investigate the reason behind these results, the secondary structures of the DNA sequences were predicted by using the mFOLD software. The most stable secondary structures at the 137 mM [Na+] and 5 mM [MG2+] at room temperature are shown in Figure 4-37. The aptamer sequence had three separate stem-loop structures, and

129 the T overhang segment at 3’ end was linear. The Probe-TL-1 exhibited the stem and loop made out of almost the entire sequence. Whereas, the Probe-TL-2 displayed a stem-loop structure at the 3’end, which is made up of essentially the bases of Probe-TL-1, and a linear region at the 5’ end. The Probe-CL sequence made out of poly A’s could not form any complex shapes and was linear. The observed results on the test strip were in agreement with the secondary structures of the oligonucleotides. In the case of the control line, the complementary sequences were linear and displayed a higher affinity to each other.

However, for the test line, the aptamer and the Probe-TL sequences formed stem-loop structures, and these conformations hindered the hybridization between them. The slightly better affinity of AuNPs-aptamer to Probe-TL-2 compared to Probe-TL-1 was due to the binding of the aptamer to the linear segment of the Probe-TL-2. As there was no linear segment in Probe-TL-1, it was unable to capture the AuNPs-aptamer complex.

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Figure 4-37: The most stable secondary structures of the aptamer, Probe-TL-1, and Probe- TL-2 at 137 mM [Na+] and 5 mM [Mg2+] at room temperature.

To further confirm these analyses, the tests were repeated at an elevated temperature (70 °C). At the higher temperature, the secondary structures were expected to degrade and thus enhance the hybridization between the complementary sequences to

131 capture the AuNPs-aptamer complexes. The theoretical melting temperatures (provided by

Integrated DNA Technologies) of the aptamer, Probe-TL-1, Probe-TL-2, and Probe-CL were 70.6 °C, 65.2 °C, 73.3 °C, and 34.4 °C, respectively. As shown in Figure 4-38, the

Probe-TL-2 was able to capture the AuNPs-aptamer complex similar to the Probe-CL; however, Probe-TL-1 still did not show any signal even at 70 °C. In order to understand the observed results, the secondary structures at 70 °C were predicted (Figure 4-39).

Among the three stem-loop structures of the aptamer at room temperature, two were destroyed by the heat, whereas one stem-loop conformation at the 5’end was still intact.

Similarly, the stem-loop structures of the Probe-TL-1 and Probe-TL-2 at 70 °C were undamaged. The observed results were in agreement with these secondary conformations.

Due to the melting of two stem-loop conformations of the aptamer, Probe-TL-2 was able to capture the AuNPs-aptamer, improving the intensity of the red band. The Probe-TL-1 and the complementary part of the aptamer did not bind due to the steric hindrance imposed by the undamaged stem-loop conformations. Based on these results, Probe-TL-2 was chosen for further experiments, as it will require mild conditions compared to Probe-TL-1 to make it bind to the AuNPs-aptamer complex at room temperature. As we were able to acquire the test signal at 70 °C, the next challenge was to obtain the test line signal at room temperature.

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Figure 4-38: Capture of AuNPs-aptamer complex by Probe-TL-1, Probe-TL-2, and Probe- CL sequences at 70 °C. (Note: The conjugate pad was not blocked and the amount of AuNPs-aptamer complex was 2.5 µl).

133

Figure 4-39: The most stable secondary structures of the aptamer, Probe-TL-1, and Probe- TL-2 at 137 mM [Na+] and 5 mM [Mg2+] at 70 °C.

One major component of the lateral flow test strip development is the selection of an appropriate blocking buffer. Various components of the test strip such as sample pad, conjugate pad, and nitrocellulose membrane can be blocked with various buffers to obtain the signal, enhance sensitivity, minimize nonspecific binding, optimize flow rate, and improve reproducibility [276]. In our current study, we started the blocking process with

134 the conjugate pad. The conjugate pad was saturated with the 10 mM PBS containing 1% ovalbumin, 0.25% Tween 20, 2% sucrose, and 0.2% sodium azide. These components stabilize, facilitate the release, and reduce the nonspecific interaction of AuNPs-aptamer conjugates [276]. More specifically, sucrose and sodium azide act as a preservative. The other components, Tween 20 and ovalbumin, reduce the nonspecific interaction of AuNPs- aptamer on the nitrocellulose membrane without requiring additional blocking. Tween 20, being a detergent, also facilitates the fast migration of the conjugates. In this experiment, the blocking buffer mainly helped to obtain the signal at the test line at room temperature.

As can be seen in Figure 4-40, following the conjugate pad blocking, the signal intensity similar to the one at 70 °C (Figure 4-38) for the test line was obtained. This can be attributed to the use of 0.25% Tween 20, which might weaken the secondary structures and promote the capture at the test line.

Another factor impacting the sensitivity is the amount of the AuNPs-aptamer conjugates [245, 277]. For comparison, two different volumes, 2.5 and 5 µl, of the AuNPs- aptamer complex were tested. The 5 µl of AuNPs-aptamer had a better signal intensity for both test and control lines (Figure 4-40). Thus, for further experiments, 5 µl of AuNPs- aptamer complex was used.

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Figure 4-40: Test strips with 2.5 and 5 µl of AuNPs-aptamer after blocking conjugate pad with 10 mM PBS containing 1% ovalbumin, 0.25% Tween 20, 2% sucrose, and 0.2% sodium azide.

4.2.4.4 Identification of appropriate binding conditions between AuNPs-aptamer and H3

Before testing H3 samples with the test strip, an additional experiment to identify the appropriate binding conditions between the AuNPs-aptamer and the H3 was performed.

For this, H3 (1 mg/ml) was deposited on the nitrocellulose membrane by the stamping process and dried for an hour at room temperature. No Probe-TL-2 and Probe-CL lines were created. Next, the AuNPs-aptamer conjugates (5 µl) were deposited on the conjugate pad and dried for 10 minutes. Subsequently, the prepared strips were tested with 150 µl of

H3 BB and the same volume of conjugate pad blocking buffer as running sample buffer.

Neither of these solutions had H3. As can be seen in Figure 4-41, the strip tested with conjugate pad blocking buffer had the brighter red band depicting the higher amount of

136 binding between the AuNPs-aptamer and immobilized H3. Utilization of BB as the sample buffer resulted in the faint band or reduced binding on the lateral flow test strip. In the case of the earlier strip, the blocking buffer might have helped to reduce the nonspecific binding and enhanced the release of more AuNPs-aptamer conjugates. Thus, for H3 on the test strips, the samples were prepared in the conjugate pad blocking buffer.

Figure 4-41: Identification of appropriate binding conditions between the AuNPs-aptamer and the target molecule H3. Histatin 3 was immobilized on the nitrocellulose membrane (No Probe-TL and Probe-CL), and conjugate pad blocking buffer and BB were tested as the sample buffer.

4.2.4.5 Detection of H3

Under the optimal conditions, 20 µg/ml H3, prepared in the conjugate pad blocking buffer, was tested with the developed aptamer-based test strip. For the control, the strip

137 was tested with the conjugate pad blocking buffer without H3. The control strip displayed the red bands at both the test and the control lines (Figure 4-42). Whereas, the sample with

H3 was able to diminish the test line intensity substantially while exhibiting a colored line at the control zone. These results indicate that the developed strip was successful in differentiating the samples with and without H3. The concentrations of H3 in this experiment was slightly higher than the salivary level of H3 in a healthy individual (1.5-

11.5 µg/ml). Although such higher concentrations might simulate stressed conditions, it is also important for the test strip to detect in the salivary range. In order to make the strip respond to lower concertation, other optimizations, such as blocking of the sample pad and nitrocellulose membrane, should be performed. The blocking process can reduce the nonspecific adsorption of H3 on these components, making more H3 available to interact with AuNPs-aptamer.

Figure 4-42: Response of lateral flow test strip for 0 (control) and 20 µg/ml H3 prepared in the conjugate pad blocking buffer at room temperature.

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One of the most important aspect of the test strip development is the reproducibility.

As can be seen in the images presented in this study, the width of the test and control lines had some variability. The variability was due to the manual deposition of the test and control line probes by the stamping method. Although various commercial machines are available for the fabrication of lateral flow test strips, they are more suitable for large-scale production and are expensive. Nevertheless, the stamping method in this project was able to demonstrate the proposed concept of the aptamer-based lateral flow test strip for POC detection.

4.3 Investigation of the use and challenges associated with saliva-based

measurements

This section presents the results and the discussions regarding the implementation of human saliva and artificial saliva with the developed assay formats such as ELASA and

AuNPs-based colorimetric assay.

4.3.1 Analysis of H3 doped artificial saliva samples with ELASA

To verify the usability of the aptamer for H3 detection, ELASA was performed by implementing artificial saliva samples spiked with external H3. Two different types of samples were used for this purpose. Group A was the commercial artificial saliva, Biotene

Dry Mouth Oral Rinse, which is used to get relief from dry mouth symptoms; whereas, group B was a lab-made fluid simulated to have a similar viscosity and electrolyte concentrations as human saliva. Both formulations were spiked with 15 µg/ml, 25 µg/ml, and 35 µg/ml of H3. The physiological level of salivary H3 in healthy adults is in the range of 1.7-11.8 µg/ml [263]. Slightly higher concentrations of H3 were used in this experiment

139 to mimic the physiological state in a stressed condition. As can be seen in Figure 4-43, the truncated aptamer sequence, LI-H3-APT-3, was able to discriminate all three concentrations of H3 externally supplemented in both saliva samples. The data clearly reveal that as the H3 concentration increased, the intensity of the signal also increased, depicting the increased binding between the immobilized H3 and aptamer. This trend was basically consistent in both types of saliva formulations. Compared to each other, group A samples exhibited a slightly higher signal than group B saliva. The diminished signal for group B can have two origins. First, some of the components of the group B saliva may have blocked the amine binding sites on the plates limiting the H3 coating. Second, the higher viscosity of group B saliva may have limited the access of H3 to the amine binding sites leading to reduced immobilization on the wells. Nevertheless, the ELASA assay was able to demonstrate aptamer-target binding and H3 quantification in complex biological samples. Overall, these results are the indication that the identified aptamer can be implemented on various assays and sensing platforms to develop more sensitive and advanced tests for the salivary H3.

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Figure 4-43: The binding performance of the aptamer LI-H3-APT-3 to the H3 doped in artificial saliva samples. Group A saliva was the commercially available product, Biotène Dry Mouth Oral Rinse, and Group B saliva was carboxymethyl cellulose-based formulation prepared in the lab. The blank samples did not contain any external H3 but were incubated with an equal amount of aptamer.

4.3.2 Implementation of human saliva with AuNPs-based colorimetric assay

format-2

In order to investigate the application of human saliva, the AuNPs-based assay format-2 was chosen. This assay can be accomplished in 12 minutes and has a linear range within the salivary level of H3.

4.3.2.1 Selectivity

Before testing saliva samples, a more detailed study to investigate the selectivity of the assay was performed. Saliva is a complex biological fluid containing proteins, enzymes, hormones, nucleic acids, cytokines, antibodies, peptides, polynucleotides, electrolytes, and other components. It is not possible to test each of the components; therefore, some of the

141 representative molecules, such as larger proteins (mucin and BSA), small peptide (H8), organic molecules (glucose and urea), and steroid (cortisol), were chosen for the selectivity test. Also, a blank sample was included for comparison. For each molecule, 5 µg/ml in BB was prepared and tested. As can be seen in Figure 4-44, the absorbance ratio for H3 was considerably larger than those of other molecules. The signals for the molecules other than

H3 were only approximately around the background, or even smaller. The relative values of signal increase or decrease compared to H3 were 2.6%, 2.1%, 7.9 %, 12.4%, 12.6%, and

8.6 % for H8, glucose, urea, BSA, mucin, and cortisol, respectively (relative signal% =

((signal due to the interfering molecule - signal due to blank sample)/(signal due to H3 - signal due to blank))x100). This excellent selectivity of the assay can be contributed to the higher specificity of the aptamer.

0.4

0.3

A /

0 0.2

6 A 0.1

0.0 e l k 3 8 s ea A in o an H H o r S c is l c U B u rt B lu M o G C

Figure 4-44: Selectivity of the AuNPs-based assay format-2 against various components of saliva. Absorbance ratio for 5 µg/ml of each molecule was obtained. For blank sample, BB was tested.

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4.3.2.2 Application with human saliva and artificial saliva

To investigate the feasibility of the assay format-2 for H3 detection in human saliva, the undiluted saliva sample doped with 10 µg/ml H3 was tested. As depicted in Figure 4-

45, the assay was unable to differentiate externally added H3 in human saliva. Following this finding, the artificial saliva was also tested with a similar technique. Surprisingly, similar results were obtained for artificial saliva as well (Figure 4-45). In both of these formulations, the doped H3 in the samples was unable to elicit any response.

0.5

0.4

0 2

5 0.3

0 4

6 0.2 A

0.1

0.0 k 3 k 3 k 3 n H n H n H la l la l la l -B /m -B /m -B /m a g a g r g iv µ iv  fe µ l 0 l 0 f 0 a 1 sa 1 u 1 S - l - b - a a a g r iv i iv n fe l ic l i f a if sa d u S t l in b r a B g A ci in fi d i n rt i A B

Figure 4-45: Absorbance ratios of the blank and H3 (10 µg/ml) doped human saliva, artificial saliva, and buffer samples.

In order to explain these results, an investigation was done to understand the interactions of the proteins and CMC with the AuNPs. The colloidal AuNPs are stabilized by a citrate anion layer, which is labile and offers ligand exchange. Whenever the AuNPs

143 come into contact with a complex biological fluid, the protein molecules in the sample adsorb onto the surface of AuNPs through electrostatic, hydrophobic, van der Waals, and dispersive forces [278]. The adsorbed molecules form a dense protein coating known as protein corona. This initial protein layer consists mainly of the more abundant proteins loosely bound to the AuNP surface, which slowly transforms into a more densely packed layer of higher affinity proteins [279]. Finally, the protein corona shields the AuNPs and alters their size and composition, providing the AuNPs with a new biological identity [278].

Similarly, it has been reported that the carboxylic groups (-COO-) of the CMC demonstrate a strong affinity to the AuNPs. Therefore, the interaction of CMC with AuNPs leads to the formation of a large polymer layer on the surface that protects the AuNPs against the salt-induced aggregation [280, 281]. The CMC layer, however, is labile and can be replaced with the thiol ligands [282].

Based on these explanations, it was anticipated that the mucin in saliva and CMC in artificial saliva have formed a shielding layer around the AuNPs, preventing their aggregation.

Another observation in Figure 4-45 was that the absorbance ratios of the blank

(saliva and artificial saliva) and H3 doped samples (saliva and artificial saliva) were slightly lower than the buffer-blank. As AuNPs exhibit distance-dependent optical properties, the lower absorbance ratios might have resulted from the increased inter- particle distance caused by the higher amount of repulsion between AuNPs after being modified with the proteins or CMC.

In order to verify that the mucin and CMC suppressed the assay signal, the samples of mucin (1.0 mg/ml) and CMC (10 mg/ml) spiked with the H3 (10 µg/ml) were examined.

144

The mean salivary concentration of mucin in the whole saliva is about 1.19 mg/ml [283].

Human saliva does not contain CMC; however, it is a crucial component of commercial saliva substitutes [284]. The concentration of CMC used to prepare artificial saliva for this study was 10 mg/ml. As expected, in both types of samples, doped H3 was unable to demonstrate the signal (Figure 4-46). These outcomes provided further support to our explanation that the mucin and CMC formed a capping layer on the surface of AuNPs, which prevented their aggregation.

0.20

0.15

A /

0 0.10

6 A 0.05

0.00 l) 3 l) 3 /m H /m H g l g l /m /m m g m g (1 µ 0 µ 0 (1 0 in 1 1 c )- C - u l l) m M M / C /m g g m m 1 0 ( 1 n ( ci u C M M C

Figure 4-46: Response to externally added H3 (10 µg/ml) in 1 mg/ml mucin and 10 mg/ml CMC.

To further elucidate the interaction of mucin and CMC with AuNPs, mucin (1 mg/ml) and CMC (10 mg/ml) samples (10 µl) were reacted with the citrate-stabilized

AuNPs (200 µl), and NaCl (40.32 mM) was added. The results were compared with the

145

AuNPs treated with 25 µM aptamer (10 µl) and 40.32 mM NaCl. None of these samples contained H3. For the blank sample, AuNPs were treated with NaCl only. As shown in

Figure 4-47, the presence of mucin, CMC, and aptamer prevented the NaCl-induced aggregation of AuNPs. Whereas the blank sample demonstrated the aggregation when challenged by the NaCl. These results further confirmed that the mucin and CMC adsorb to the surface and protect the AuNPs, acting similar to the aptamers.

1.5

0 1.0

4 6

A 0.5

0.0 l) l) ) k m m M n / / µ la g g 5 B m m 2 1 0 ( ( 1 r n ( e ci C m u ta M p M C A

Figure 4-47: Interaction of mucin, CMC, and aptamer with citrate-stabilized AuNPs.

After confirming the interference of mucin and CMC, additional experiments were performed to identify the interfering levels of these components. Various concentrations of mucin and CMC were prepared in BB and doped with 10 µg/ml H3. The samples were then examined with the assay. As can be seen in Figure 4-48, mucin and CMC exhibited a significant effect on the assay signal until the concentration of 0.05 mg/ml for mucin

(Figure 4-48 (A)) and 0.025 mg/ml for CMC(Figure 4-48 (B)).

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Figure 4-48: Response to 10 µg/ml H3 spiked in various concentration of mucin (A) and CMC (B) samples.

In the next experiment, the tests were performed to find out if the components of artificial saliva other than CMC interfere with the assay. In order to inspect this, artificial saliva without CMC was prepared and doped with 10 µg/ml H3. For comparison, BB-blank and BB containing 10 µg/ml H3 were also run. As displayed in Figure 4-49, the assay responded to the H3 doped artificial saliva sample, which indicated that the CMC was the only component in artificial saliva that hindered the assay. Also, it was observed that the absorbance ratios for both artificial saliva-blank and artificial saliva-H3 were slightly higher than their counter-part BB-blank and BB-H3 samples, respectively. Nevertheless, the net difference between the blank and the H3 sample in both cases was similar, representing an excellent recovery. As the assay signal depends on the salt concentration, the difference in the background may occur due to the different ionic compositions of the samples. This experiment also revealed the outstanding selectivity of the assay as none of the components of artificial saliva at such a higher concentration interfered.

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0.5

0.4

0 2

5 0.3

0 4

6 0.2 A

0.1

0.0 ) 3 k 3 C H n H M l la l C /m -B /m o g r g n µ fe u ( f 0 a 10 u 1 iv )- b - l g er sa C in ff l M d u ia C n b c i g fi o B n ti (n i r a d A v in li B sa l ia ic if rt A

Figure 4-49: Response to H3 (10 µg/ml) doped artificial saliva sample without CMC.

In the following experiment, the dilution needed for human saliva in order to suppress the matrix effect in the assay was investigated. The processed human saliva sample was diluted to 1:10, 1:20, 1:50, and 1:100 in BB, and the resulting samples were spiked with 10 µg/ml H3. The spiked samples were then tested with the assay. The data are presented in Figure 4-50. As can be seen, the H3 in 1:10 and 1:20 diluted samples did not elicit any response. At these dilutions, the concentration of the interfering molecules in the sample might be sufficient to inhibit the AuNPs aggregation. Beyond the 1:20 dilution, the assay was able to respond to the H3, and in fact, the signal recoveries in 1:50 and 1:100 diluted samples were 98.1% and 107.7%, respectively. The recoveries (%) were estimated

148 by comparing the obtained readings with the calculated value for 10 µg/ml H3 from the calibration curve.

0.5

0.4

0 2

5 0.3

0 4

6 0.2 A

0.1

0.0 a a a a 3 v 3 v 3 v 3 v li H li H li H li H a l a l a l a l S /m S /m S /m S /m 0 g 0 g 0 g 0 g :1 µ :2 µ :5 µ 0 µ 1 0 1 0 1 0 :1 0 -1 -1 -1 1 -1 a a a a v v v v li li li li a a a a S S S S 0 0 0 0 1 2 5 0 : : : :1 1 1 1 1

Figure 4-50: Matrix effect of human saliva in the assay. The processed saliva was diluted to various ratios (1:10, 1:20, 1:50, and 1:100) with BB, and the resulting samples were spiked with 10 µg/ml H3.

To further demonstrate the feasibility of this assay, experiments were performed by standard addition method in diluted human saliva. Various concentrations of H3 (2.5, 5,

10, and 20 µg/ml) were spiked in 1:50 diluted saliva sample and tested. The results and calculations are summarized in Table 4.4. The recoveries of the samples were in the range of 89.23% to 104%, which were satisfactory results. Even though the assay was able to demonstrate the sensing in 50-fold diluted saliva, it was not sensitive enough to detect the residual H3 in such higher dilution. If the sensitivity of the assay can be enhanced approximately 50 times, it would be possible to quantify H3 in human saliva. The

149 sensitivity of the assay can be enhanced to some extent by implementing some of the techniques, such as reducing the H3 adsorption on the tubes, decreasing the amount of

AuNPs, decreasing the corresponding amount of aptamer, and increasing the volume of the testing sample. Therefore, further investigation is required to design an appropriate procedure to increase the sensitivity for practical application. Moreover, by just changing the aptamers, this type of colorimetric assays may find extensive application in food or the environment monitoring where the samples do not present significant matrix effect.

Table 4.4: Analytical results for H3 in human saliva samples. The processed saliva was diluted 50 times with BB and adulterated with various concentration of H3.

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Chapter 5

5 Conclusions and Recommendations

5.1 Conclusions of this research project

This research project attempted to establish a widely applicable aptamer selection method and different assay platforms based on the identified aptamers for H3 detection in human saliva. For the purpose of establishment, first known aptamers of H3, capable of changing conformation while binding to the target, were successfully identified by the library immobilization version of the SELEX method. During the SELEX process, the selection conditions were tuned to obtain the aptamers with specificity and higher affinity.

In order to remove the sequences that can nonspecifically bind to the closely related molecule H8, counter-selection steps were incorporated within the SELEX method. After multiple rounds of selection and counter-selection, 4 unique aptamers sharing a consensus region were obtained. The sequences were first characterized by utilizing computational tools to identify secondary structures and stability. Based on the computational studies, the most promising aptamer candidate was taken to develop an MB-based affinity capture assay combined with SDS-PAGE for visual conformation of binding and specificity. The aptamer displayed binding affinity and specificity to H3, indicating the successful aptamer selection process.

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To carry out a more detailed study on affinity and specificity of the aptamers, a direct format ELASA was established. Various sequences were examined, and the highest affinity sequence was identified. Also, the effect of the primer binding region on the affinity of the aptamer was assessed. All of the aptamers, including the truncated sequence, displayed affinity and specificity towards H3, thereby supporting the results of the MB- based capture assay and further validating the success of the SELEX process. The truncated aptamer sequence displayed the highest affinity, and thus was chosen for the development of various assays.

In a further experiment, two different formats of AuNPs-based colorimetric assays were developed by utilizing the signaling feature of the aptamer to detect H3 quantitatively.

For assay format-1, the strategy relied on the aggregation of AuNPs based on the specific interactions between the H3 aptamer adsorbed on the surface of AuNPs and the target molecule H3. Upon contacting the target molecule, the adsorbed aptamer changed the structural conformation and lost the ability to protect AuNP against salt-induced aggregation. By optimizing the key parameters, such as the amount of salt, the amount of aptamer, and the incubation times, the established assay showed good linearity between

1.5 to 20 μg/ml, with a detection limit of 0.199 μg/ml. The assay format-2 also utilized the same technique; however, it had a different order of interaction between the assay components. The aptamers and target molecules were reacted first, and then AuNPs were added. Through this format, we were able to reduce assay time from 4.5 hours for format-

1 to 12 minutes in format-2 in exchange for slightly reduced sensitivity. The assay format-

2 had a similar linear range as format-1 and a limit of detection of 0.505 µg/ml. Both of

152 these formats utilized unmodified aptamers and did not require any modifications or the washing and processing steps.

In addition, in an effort to develop a highly sensitive assay, a SERS-based magnetic aptasensor in a competitive format was proposed. The AuNPs were modified with an optimal amount of Raman reporter molecule, 4-NTP, and thiol modified H3 aptamer to fabricate the signal probe. Capture probes were fabricated by conjugating MBs with the cDNA sequence. Upon increasing the concentration of H3, the aptamers on the signal probe were occupied with the target molecules, decreasing the number of signal probes that could bind to the capturing substrate. Hence, at higher concentrations of H3, the SERS signal was decreased. This method was able to differentiate various concentrations of H3

(0, 1, and 5, µg/ml) in buffer media, demonstrating the promising quantitative nature.

However, due to the highly sensitive nature of the SERS-based assays, the assay was expected to have the sensitivity for slightly lower concentrations. Such a response would allow dilution of saliva samples to eliminate the matrix effect associated with the viscous behavior of the saliva. Therefore, with further investigation and improvement, it could be a promising platform for the detection of deficient biomarkers in saliva samples.

Moreover, an aptamer-based lateral flow test strip based on the competitive format was successfully demonstrated for the rapid on-site detection of H3. The competition occurred between the migrating H3 in a sample and the Probe-TL-2 immobilized on the test zone of the strip for capturing AuNPs-aptamer conjugate. After optimizing some key parameters, the developed test strip was able to differentiate samples with and without H3.

The entire detection process could be accomplished within 5 minutes. With some additional testing and optimizations, the test strip could overcome the limitations of saliva as a

153 diagnostic fluid for point-of-care testing by replacing the laboratory-based infrastructure required for the conditioning of saliva. Also, it might offer inexpensive, quick, reliable, and easy testing of biomarkers in resource-limited areas, such as militaries, developing countries, and the underserved population in developed nations.

Ultimately, to investigate the challenges and possible use of saliva with the developed assays, artificial and human saliva samples were evaluated with the ELASA and

AuNPs-based colorimetric assay format-2. In ELASA, the aptamer was able to differentiate between various concentrations of externally supplemented H3 in both formulations of artificial saliva samples. For the AuNPs-based assay, the molecules interfering with the assay in saliva were identified, and the possible mechanism of interference was discussed.

The dilution of the sample mitigated the interference associated with saliva, and the assay demonstrated an excellent recovery for 1:50 diluted human saliva samples spiked with a known amount of H3. These results suggested that, if the assay is sufficiently sensitive for lower concentrations of H3, it has the potential to analyze the H3 in saliva. Therefore, it is essential to enhance the sensitivity of the assay for the deficient levels of analytes.

Nevertheless, this type of method might be useful for the detection of other molecules in less complex samples by using the corresponding aptamer sequence.

In summary, this dissertation has established the key foundation work for the identification of signaling aptamers and the development of aptamer-based simple, sensitive, and rapid assays for the detection and quantification of novel biomarkers. While the study has been established for H3, it is certainly possible to adapt to other biomarkers, which currently lack more accessible detection technologies.

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5.2 Future research recommendations

5.2.1 Aptamer selection and modifications to improve affinity

In the present study, aptamer selection was performed for 14 rounds, and within the

SELEX method, counter-selection steps were established with H8. Based on the performed studies, no cross-reaction with the tested salivary components were observed while comparing the responses to the equivalent concentration of H3. However, saliva contains several other histatins, which were not considered in this study for testing cross-reactivity of the aptamers. It might not be possible to test these aptamers against all histatins; however, by selecting some of the variants that closely match H3 should be investigated.

In the case of nonspecific binding, the SELEX process should be revisited by establishing counter-selection with the interfering molecules. Also, the future selection work should be accomplished by using partially randomized (doped) aptamer library. Since the currently selected aptamers have already shown some affinity and specificity to the target molecule, by finding their critical motifs responsible for target binding, a doped aptamer library can be fabricated to reselect optimal aptamers: this can usually improve their affinity to the target [285].

In addition, there are several ways to enhance the affinity of the aptamer through post-SELEX modifications. The results presented in section 4.2.1.1 demonstrated that the primer binding sites hindered the binding affinity of the aptamer to the target molecule H3.

The truncation study should further be expanded to map the effective binding regions within the aptamers that are critical for the target binding. Truncation of the aptamers might result in high-affinity sequences [286]. Further investigations might be performed to see if these aptamers have an affinity to another epitope of H3. If the aptamers that recognize

155 different epitopes of the target molecule are found, the affinity to the target can be increased significantly by joining these aptamers [287]. Also, such a pair can be applicable in various diagnostic platforms based on a sandwich format of the assay. Besides, the binding affinities of aptamers are limited by the low hydrophobicity of nucleic acids. Hence, the addition of hydrophobic moieties into aptamers expands the diversity of interactions and enhance the binding affinity [287]. By implementing above mentioned techniques, the affinity of the aptamer can be improved.

5.2.2 Gold nanoparticles-based colorimetric assay

In this study, two different formats of the colorimetric assay utilizing the structural feature of the identified aptamer were developed. Various aspects of the assay were studied to obtain the optimal response. As the assay format-2 can be performed in a short time period and offer visual detection, it might be a promising method for the resource-limited areas for preliminary screening. However, further studies should be focused on developing a more sensitive method to detect lower concentration samples. Additionally, the development of a cellphone-based app for color analysis can aid the implementation of the assay in the POC environment [190].

5.2.3 Improvement on SERS aptasensor

As explained in section 4.2.3.5, the SERS aptasensor was unable to respond to the lower concentrations of the target molecules. The future work should be focused on making it respond to the lower concentrations. First, the nonspecific adsorption of the peptide on the surface of the tubes should be addressed. The techniques mentioned in section 4.2.3.5, such as the use of surfactants and the blocking of the tubes can be performed. Another way to improve the sensitivity of the SERS assay is to develop the SERS substrate with high

156

SERS intensity. Raman reporter embedded core-shell nanoparticles such as gold-coated silver (Ag@Au) or silver-coated gold (Au@Ag) are the most promising SERS-active probes due to their enhanced EM field between two close metal layers [268, 288, 289].

The SERS activity of the core-shell assemblies can be accurately tuned by altering the core/shell shape and size and shell thickness [290]. In these structures, the outer layer prevents the dissociation of Raman reporter molecule from the surface, thus increasing the chemical stability of the probe molecules, and also provide the platform for the conjugation of affinity molecules [268, 269]. Additionally, further signal enhancement can be obtained by encapsulating the capture substrates (magnetic particles) with a metallic layer of gold or the silver. The metallic layer provides the surface for the immobilization of affinity molecules while retaining the magnetic properties of the embedded magnetic nanoparticles. When combining capture probes with the core-shell nanoparticles (Figure

5-1), the formation of hot spots between the capture and signal probes takes places, enhancing the SERS signal to exceptionally high levels [270]. According to the assay mechanism, the presence of the target molecules will result in the depletion of the SERS signal corresponding to the increased target concentration.

157

Figure 5-1: Proposed improvement to the SERS based magnetic assay.

5.2.4 Lateral flow test strip

The first aspect that needs to be addressed is the reproducibility of the test and control lines. As mentioned in section 4.2.4.5, by using commercially available devices for assembling the test strip and dispensing of test and control lines, highly reproducible test strips can be achieved. Following this, the components of the test strips such as the sample pad and the nitrocellulose membrane should be blocked to enhance the sensitivity to the lower concentrations of H3.

The ultimate purpose of the developed test strip is the detection of H3 in human saliva. For implementation with saliva, some of the parameters of the test strip detection method should be optimized. One of the known hurdles for application of saliva is the viscosity. In general, saliva requires laboratory-based processing methods to eliminate

158 mucin and food particles. The developed test strip is expected to mitigate the effect of viscosity as its components are anticipated to filter the mucin and food particles to some extent. To obtain the desired performance, the length of the various components of the test strip should be optimized so that the test strip can filter out the mucin and other particulate matters effectively. Also, the materials with different physical properties such as smaller pore size as can be tested. Other aspects that need to be evaluated are the impact of pH, ionic concentrations, and temperature on the signal intensity. The stability of the test strip should also be determined. In order to eliminate the variability relating to the sample volume, a sampling device can be added on the top of the sample pad. Following these optimizations, the tests can be made quantitative by determining the color intensity at the test line through the image processing apps that can be stalled on the cell phones. Such technologies will make the lateral flow test strip applicable to the resource-limited areas to preliminary screen the disease and or condition of an individual.

5.2.5 Continuous, real-time measurements of salivary biomarkers

The assay formats developed in this study generally perform a single time-point measurement, which limits their use for the continuous real-time monitoring of the biomolecules. A sensing system capable of continuously measuring salivary biomarkers in real-time would be highly beneficial to track the circadian variations and the real-time responses to the therapeutic interventions. The possible solution to this would be the incorporation of electronic aptamer-based (E-AB) biosensor within the mouthguard along with electronics and microfluidic sampling system. The E-AB biosensors have already been implemented for monitoring circulating drugs, such as doxorubicin and kanamycin, in live rats and in human whole blood for several hours without requiring exogenous

159 reagents. Interaction with a target leads to the conformational rearrangement of the aptamer, modified with a redox moiety at one end, generating the electrochemical signal.

The conformational change is reversible, which enables continuous real-time measurements. Moreover, the sensors have shown stability, selectivity, and sensitivity in complex sample matrices [291]. However, no such aptamer-based systems have been developed for testing saliva. Also, miniaturization of electronics and power requirements may hinder the application of E-AB biosensor in the oral cavity. The other idea could be the implementation of a passive radio-frequency identification (RFID) technology-enabled biosensors. Recently, the integration of passive RFID technology with biosensors has shown a promising possibility for the development of miniaturized, wireless, battery-free, and cost-effective biosensing systems. In RFID enabled biosensors, the receiver placed nearby transmits the power to the RFID tag, which is connected with the sensing element.

The binding of analytes on the sensing component can influence the electrical properties, such as permittivity, impedance, and resonance frequency, of the system, which is then transmitted back to the reader device [292]. By functionalizing the sensing element with the specific recognition elements, the selectivity can be obtained. Based on a similar idea,

Mannoor et al. have reported a novel graphene-based dental tattoo platform capable of continuously monitoring bacteria and transmitting data wirelessly. To obtain the selectivity, the sensing element was functionalized with antimicrobial peptides. The binding with bacteria changed the electrical conductivity of the sensing element, which was then read by the RF reader. The testing was performed in pooled saliva [293]. Similar technologies can be employed for the development of RFID enabled aptasensors.

Implementation of biosensors in the oral cavity, however, might need the microfluidic

160 sampling device capable of filtering mucin and particulate matters and continuously supplying fresh saliva. As saliva is gaining momentum as a diagnostic fluid recently, the development of novel biosensing technologies with the functionality of continuous and real-time measurement is anticipated to increase in the near future.

161

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Appendix A: Experimental protocol for library immobilization SELEX

A.1 Library, primers, and capture probes design and synthesis

1. Synthesize the following sequences:

Library:5’-GGTGACTGCTACTGTGTTGG-N44-

CCACACATCCAAGCAGAACC-3’

Forward primer: 5’-GGTGACTGCTACTGTGTTGG-3’

Reverse primer: 5’-GGTTCTGCTTGGATGTGTGG-3’

2. Design the capture probes of 7 bases and 9 bases complementary to the 5’ region

of the library sequences.

7-mer capture probe: 5’-GTCACCC/3Bio/-3'

9-mer capture probe: 5'-CAGTCACCC/3Bio/-3'

3. Reconstitute these oligonucleotides in Tris-EDTA (10 mM Tris and 0.1 mM

EDTA (pH 7.5)) buffer at 100 µM concentration and store at -20ºC.

A.2 Immobilization of biotinylated capture probes on streptavidin- functionalized MBs

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1. Transfer 100 μL of streptavidin-coated MBs in a 1 ml microcentrifuge tube and

wash three times with 500 μL BB.

 BB: 50 mM Tris, 137 mM NaCl, 5 mM MgCl2, pH 7.4

 For washing MBs, gently agitate the sample for 5 minutes, put the sample

tube on a strong magnet for two minutes, remove most supernatant using a

200 µl pipette, and then remove the rest of supernatant using a 10 µl pipette

carefully.

2. Following the washing, resuspend the MBs in 50 µl of BB.

3. Add 50 µl of 100 µM 7-mer biotinylated complementary capture probe to the

washed beads, mix well, and incubate for 30 minutes at room temperature with

gentle shaking.

 Use 7-mer probe for round 1-9 and 9-mer probe for 10-14 (Table A.2).

4. Wash beads four times in 500 μL of BB and resuspend in 100 µl of same buffer.

5. Store at 4 °C for further use.

A.3 Immobilization of library sequences on capture probe-immobilized

MBs

1. Transfer 20 µl of capture probe-coupled MBs to a 0.2 ml vial and add 2.5 µl of 100

µM ssDNA library pool (previously heated for 5 minutes at 95 ˚C, snap cooled on

ice for five minutes, and equilibrated at room temperature for 10 minutes), mix

well, and incubate for 30 minutes at room temperature with gentle shaking.

2. After incubation, wash the library-hybridized MBs three times with 200 µl BB.

3. Resuspend the washed beads in 20 µl of BB.

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A.4 Binding and amplification

1. For first round of SELEX, add 40 µl of 100 µM H3 diluted in BB to the 20 µl of

above prepared library immobilized MBs and incubate for 30 minutes using gentle

rotation.

 The concentration of H3 for each round is listed in Table 4.1.

2. Following the incubation period, retain the supernatant by using a strong magnet to

pull down the MBs.

3. Dialyze the retained supernatant using 10,000 Da cutoff dialysis cassettes

according to the manufacturer’s instructions.

 Briefly, insert sample into the dialysis cassettes and incubate for 2 hours in

binding buffer, change the buffer, and incubate the cassette for another 2

hours, followed by a final buffer change, incubate at 4 °C overnight.

4. Remove the dialyzed supernatant and perform PCR (Exponential amplification

phase). For PCR, take a PCR tube and add 12.5 µl of SYBR Green I, 9.5 µl of

dialyzed supernatant, 1.5 µl of 10 µM each forward and reverse primer. Setup the

following parameters and place the PCR tube in sample holder of iQ5 system.

Polymerase chain reaction settings.

 Initial denaturation: 94 °C for 2 minutes-1 cycle

 Amplification cycles-16 cycles

o Denaturation: 94 °C for 30 seconds

o Annealing: 60 °C 1 minutes

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o Extension: 72 °C for 1 minutes

 Final extension: 72 °C for 7 minute-1cylce

 Chill: 4 °C

 During setup in the iQ5 software, make sure the volume, tube, and seal types

are matching with the actual settings.

 Place some water containing PCR tubes around the sample in the iQ5

sample holder. This helps an equally heat distribution of heat and prevents

the overheating of sample.

5. After the exponential PCR phase, convert the dsDNA sequences to ssDNA by

running asymmetric PCR. For which, take out the PCR tube and add 12.5 µl of

SYBR Green I, 2.5 µl of 10 µM forward primer, and 10 µl of ultrapure water, and

run PCR with same settings.

 Use forward primer only for asymmetric PCR.

 Total volume of the sample will be 50 µl and use same temperature cycles

as shown in Table A.1.

6. Use this asymmetric PCR product for next round of SELEX.

 Perform the UV-Vis absorption measurement and determine the

concentration of the PCR product by using the absorption value at 260 nm.

Use same amount of library as first round for other rounds of SELEX

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A.5 Counter-selection 1. Start negative-selection at round six. Following the immobilization of library

sequences on the MBs, add 40 µl of H8 in BB and shake gently for 30 minutes.

Using the magnet, discard supernatant and retain the MBs.

 The concentration of H8 for each round, wherever applicable, is given in

Table 4.1.

 The sequences having affinity to H8 gets released from the MBs and are

discarded. The MBs with remaining library sequences are retained for

further SELEX.

2. Wash retained MBs three times with 100 µl of BB and resuspend in 20 µl of same.

3. Using the washed MBs, resume the selection cycle for H3.

4. Repeat the SELEX process for 14 rounds.

5. For the last SELEX round, after selection, perform exponential PCR and follow the

product verification, transformation, cloning, and sequencing steps.

A.6 Luria-Bertani (LB) medium plates preparation for culturing cells

1 Take 950 ml of ultrapure water in a clean beaker and add 10 g tryptone, 5 g yeast

extract, and 10 g NaCl.

2 Adjust pH to 7.0 and bring the total volume to 1 liter.

3 Add 15 g agar.

4 Autoclave on liquid cycle for 20 minutes at 15 psi.

5 Cool the media to 55 ºC, add kanamycin (final concentration 50 µg/ml), and swirl

to mix.

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6 Pour approximately 20 ml to the petridish.

7 Let the media harden for 10 minutes, seal, invert, and then store at 4 ºC in dark.

A.7 Cloning and Sequencing

1 Before starting the process, incubate the plates at 37 ºC for at least 1 hour.

2 For ligation, add 1 µl fresh PCR product, 1 µl salt solution form TOPO kit, 1 µl

ultrapure water, and 1 µl TOPO vector.

3 Mix with pipette and incubate for 30 minutes at room temperature.

4 After 30 minutes, add 18 µl of ultrapure water to the ligation reaction to dilute the

high salt concentration.

5 Place the tube on ice.

6 Add 2 µl of the diluted TOPO cloning reaction to a vial of One Shot E. coli and

mix gently.

 For mixing, only swirl gently, do not vortex. The One Shot E. Coli cells are

very fragile.

7 Transfer the reaction mixture to an electroporation vial.

8 Electroporate the cells at 1.5 kV.

9 Transfer the cells to a dilution vial containing 1 ml of room temperature equilibrated

S.O.C. medium.

10 Close the lid and place the dilution vial in a 37 ºC water bath for 1 hour while

shaking at 150 RPM.

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11 During this incubation time, spread 50 µl of 50 mg/ml X-gal on the LB plates with

the help of the sterilized glass beads and warm the plates to 37 ºC.

12 Spread 10 µl of the sample on pre-warmed LB plates containing antibiotic and X-

gal. For the control, spread the sample on the LB plates without antibiotic.

13 Incubate plates overnight at 37 ºC.

14 Pick the white colonies and transfer them to fresh plates containing kanamycin.

15 Label the colonies and ship plates on ice for sequencing or can be stored at 4 ºC for

about a week.

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Appendix B: Agarose gel electrophoresis protocol for

PCR end product verification

After a SELEX round, it is preferred to verify the PCR end products, dsDNA and ssDNA, through agarose gel electrophoresis. In addition to real-time PCR, agarose gel electrophoresis is an important tool to verify and troubleshoot the selection process. To run the agarose gel electrophoresis, Tris-borate- ethylenediamine tetraacetic acid (TBE) buffer was used to prepare and run the agarose gel. The method for preparation of TBE buffer is as follows:

1) Ethylenediamine tetraacetic acid does not get dissolved completely into solution until the

pH is adjusted to about 8.0. So, a stock solution of EDTA (0.5 M) is prepared separately

ahead of time. For a 500 mL stock solution of 0.5 M EDTA, add 93.05 g EDTA disodium

salt (FW = 372.2) to 400 mL ultrapure water, and adjust the pH 8.0 with NaOH. Mix until

it gets dissolved and top up the solution to a final volume of 500 mL.

2) Prepare 5x stock solution of TBE buffer. To make 1 liter of 5x TBE solution, add 54 g Tris

base (FW 121.14) and 27.5 g boric acid (FW 61.83) to 900 ml of ultrapure water. Add 20

ml of 0.5 M EDTA (pH 8.0) and dissolve all the components by mixing. Afterward, adjust

final volume to 1 liter.

3) Store at room temperature.

 Discard if precipitate forms.

4) The working solution of TBE is 0.5x. Dilute the 5x concentrated stock solution 10 times

when needed.

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5) Prepare the 2% agarose gel as follows:

 Take 100 ml of 0.5x TBE buffer in a 250 ml glass bottle and add 2 g of agarose.

 Microwave the mixture for 1 minutes and 20 seconds with a loose cap (continue heating

until agarose is completely dissolved)

 Remove bottle from microwave and let it cool around 50°C (until you can comfortably

hold bottle in hand)

 Add 10 µl of 10 mg/ml Ethidium bromide solution and gently swirl to mix.

i) Dispose the pipette tip in a 50 ml centrifuge tube for disposal with hazardous

chemicals

6) Assembly of casting basin and poring of gel

 Place an appropriate casting tray in a casting basin and clamp tightly.

 Place the desired combs.

 Slowly pour cooled agarose into the casting tray.

 Remove any bubbles within the gel with the help of pipette tip.

 Allow it to solidify (usually approximately 30 minutes) and gently remove the combs

by pulling them straight up.

7) Assembly of running basin

 Remove casting tray from casting basin and place in a running basin

i) Make sure that the gel is in correct configuration so that the samples run from

BLACK to RED.

 Completely submerge gel in 0.5x TBE (make sure not to overfill the basin)

8) Sample preparation

 In a small tube combine 3 µl of 6x agarose loading dye with 12 µl of PCR product

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 Similarly, mix 1 µl of molecular weight marker with 1 µl of 6x agarose loading and

add 4 µl of ultrapure water.

9) Loading of samples

 Using a clean pipette tip, carefully load samples into wells of the gel and allow to settle

into bottom.

 Be careful not to puncture the bottom of the well and make sure not to spell the samples

outside the wells.

 Load molecular weight marker into the first and last well.

10) Running gel

 Following the sample loading, place the cover of the running basin and make sure the

electrodes are connected.

 Plug into power supply (Black wire to the black port and red wire to the red port)

 Turn on main power supply of the control box.

 Run at 100 V and 25 mA

i) Visually confirm function by identifying “bubble curtain” which will appear within

1 minute on the negative electrode wire at the top of the gel.

 Run for 2 hours

i) Larger gels can be run longer. You can hit START again without resetting any

parameters if you want more distance in your gel.

11) Viewing gels

 Remove the gel tray and place the gel on the saran wrap.

 Transfer the gel with saran wrap on the UV light board, and place UV protective

covering.

 Turn on UV light and turn off room lighting.

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Appendix C: Running SDS-PAGE

Use Mini-PROTEAN TGX Precast Gels, 12% (BIO-RAD Catalog No. 456-1043)

Running buffer Stock (10X SDS-PAGE Buffer)

Prepare 10X SDS-PAGE rurring buffer: 250 mM Tris, 1.92 M glycine (SIGMA; G8790),

1% SDS (SIGMA; L4509), pH 8.3

 Add 30.3 grams of Tris base (Fisher Scientific; BP152-1),144.1 grams of glycine,

10 grams of SDS to 800 ml of DI water

 Make final volume 1 L

 No need to adjust pH (pH is approximately 8.3)

To run the gel use 1X SDS-PAGE buffer

 Take 100 ml 10X SDS-PAGE buffer and dilute with 900 ml of DI water.

Sample Buffer

Use 2x Laemmli sample buffer (BIO-RAD, Catalog #161-0737).

Add β-mercaptoethanol (50 µl to 950 µl sample buffer) before use.

Sample Preparation:

 Sample: 5 µl

 Laemmli buffer: 4.75 µl

 β-mercaptoethanol: 0.25 µl

Heat samples at 90-100 °C for 5 min (or at 70 °C for 10 min).

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Protein Ladder/marker

Take 10 µl of Precision Plus Protein All Blue Prestained Protein Standards (BIO-RAD;

Catalog # 161-0373) and load to the gel.

Run the gels with following conditions:

 Voltage: 100 V

 Current: 25 mA

 Time: 85-95 minutes

Gel Staining with Bio-Safe Coomassie Stain (BIO-RAD, Catalog #)

 Wash gel with three times for 5 min each with 200 µl of DI water to remove SDS,

which will interfere with staining.

 Remove all the ultrapure water from staining container and add 50 ml Bio-Safe

Coomassie stain

 Gently shake for an hour. Protein bands will start to appear after 20 minutes and

reach maximum intensity within 1 hour. Longer incubation time will not increase

background.

 Rinse the gel in 200 ml of ultrapure water for at least 30 min. Rinsing the gel

extensively after staining will further reduce background. Stained gels can be

stored in water.

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