Development of Carbon Dots As Fluorescent Probes for Fluorescence in Situ Hybridisation (FISH) Application

Development of Carbon Dots as Fluorescent Probes for Fluorescence In Situ Hybridisation (FISH) Application

Phyllis Jacqueline Nishi

A thesis submitted to the Faculty of Engineering, Computing and Science

Swinburne University of Technology Sarawak

In fulfilment of the requirements for the degree of

Master of Science by Research

2019

Abstract Fluorescence in situ hybridisation (FISH) is an important bioimaging technique in molecular cytogenetics that utilises fluorescent probes that can bind specifically to a target nucleic acid sequence of a DNA or RNA. It is important that the fluorophores used to fluorescently tag probes are bright, small-sized, non-toxic and come in various colours. Thus, there is a need for new and alternative fluorescent labels to be developed to improve the performance of FISH. This thesis explores on the synthesis and application of carbon dots (CDs) as a class of versatile fluorescent label in FISH. The synthetic method for the production of CDs for use as fluorescent probes in FISH was described. CDs were synthesised by hydrothermal treatment of a carbon source in concentrated phosphoric acid solution. In this work, carboxymethylcellulose (CMC) was selected as the starting precursor and then converted into CDs with carboxylic functional groups. The optical properties and characterisation of the synthesised CDs were carried out. An amine-functionalised oligonucleotide probe was designed to detect and localise glyceraldehyde 3-phosphate dehydrogenase (GAPDH) mRNA in human adult low calcium high temperature keratinocytes (HaCaT) cell line. CDs were neutralised, isolated and dried before conjugated with the GAPDH oligonucleotide probe via carbodiimide crosslinker chemistry. The conjugated CD-GAPDH probe were applied for in situ hybridisation in fixed HaCaT cell line. The performance of the probe was evaluated via fluorescence microscopy and discussed in details in this study.

Acknowledgements First, I would like to express my utmost gratitude to my mentor and coordinating supervisor, Associate Professor Ng Sing Muk for all the support, opportunities and invaluable guidance he has provided for me in my postgraduate journey. Thank you for your constructive advices and for being very patient with me. You have set a very good example of a leader and I am very proud to be under your wings.

I would also like to thank Dr. Chua Hong Siang for helping out in my research work with his expertise in electronics and Dr. Hwang Siaw San for allowing me to carry out and learn cell culturing while using her utilities. My seniors, Dr. Jessica Fong and Ng Yann Huey for sharing their experiences and knowledge to me.

I also dedicate my thanks to Dr. Paul Neilsen and the lecturers of Swinburne University of Technology Sarawak, Dr. Irine Ginjom, Associate Professor Peter Morin Nissom, Associate Professor Moritz Mueller and Dr. Daniel Tan for the advice and suggestions they have provided for my project.

My time here in Swinburne would not be possible without the Research and Consultancy Office of Swinburne University of Technology Sarawak and the Tuition Fee Waiver Award granted to me, which I will forever be grateful for.

Special thanks to Professor Dr. Zainab binti Ngaini and her students, Arif and Eswaran, from Universiti Malaysia Sarawak (UNIMAS) for providing us with the SEM and FTIR analysis needed for this study and for guiding me throughout the whole process.

Likewise, I would like to express my thanks to Sarawak Biodiversity Centre (SBC) for allowing me to use their fluorescence imaging facilities and a very kind thank you to Dr. Kon Nyuk Fong for helping me with using the instrument.

Thank you to the science laboratory technicians; Chua Jia Ni, Marciana Jane Anak Richard, Cinderella Anak Sio and Nurul-Arina binti Salleh. They have taken good care of the researchers by making sure all the laboratory safety and technicalities are handled with properly. They also helped me with any problems I faced when dealing with lab equipment and facilities.

My thanks also go to my friends and officemates; Andrew Lim, Lee Boon Kiat, Cindy Wee, Diana Choo, Diyana Musa, Edwin Sia, Kong Ee Ling, Fay Fay, Fiona Chung,

Jasper Sing, Tay Jia Jun, Mertensia Kho, Reagan Entigu, Jong Siu Mei, Vivian Lee and Chan Wen Loong who have been alongside me during my candidature. Many of them provided me with help and advice for my lab work and thesis. I have never felt more at home in the university thanks to you all. Also a very big thank you to Johnny Tang, for being with me and supporting me throughout my candidature.

Finally, I would like to thank my parents for the support and care that they have provided for me throughout my life. Without them, getting this far is likely impossible. Last but not least, Phoebe Nishi, for being the sister and best friend that I know I could always lean on. I love you all.

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Declarations I, Phyllis Jacqueline Nishi, hereby declare that this thesis was composed by me and the work presented in it is of my own. Where work was carried out either jointly or wholly by others, due acknowledgement was given to the relative contributors, workers and authors. Any mentions of work and material previously published and written by others were quoted and referenced accordingly. No part of this thesis was previously submitted for any other degree or professional qualification.

Signed: Date: 7th June 2019

Table of Contents Abstract ...... i

Acknowledgements ...... ii

Declarations ...... iv

Table of Contents ...... v

List of Figures ...... ix

List of Tables ...... xv

1 Introduction...... 1

1.1 Brief Overview ...... 1

1.2 Problem Statement ...... 1

1.3 Thesis Aim ...... 2

1.4 Objectives ...... 2

1.5 Thesis Outline ...... 2

1.6 Scope and Delimitation ...... 4

2 Literature Review ...... 5

2.1 Fluorescence Bioimaging ...... 5

2.1.1 Fluorescence and Fluorophores ...... 7

2.1.1.1 Intrinsic Fluorophores ...... 10

2.1.1.2 Extrinsic Fluorophores ...... 12

2.1.1.3 Quantum Dots ...... 15

2.2 Carbon Dots ...... 18

2.2.1 Introduction to Carbon Dots ...... 18

2.2.2 Physicochemical Properties ...... 18

2.2.3 Top-down Approaches ...... 21

2.2.3.1 Arc Discharge Method ...... 21

2.2.3.2 Laser Ablation ...... 21

2.2.3.3 Electrochemical Methods ...... 22

2.2.4 Bottom-up Approaches ...... 22

2.2.4.1 Thermal Routes ...... 22

2.2.4.2 Microwave-assisted Methods ...... 22

2.2.4.3 Hydrothermal/Solvothermal Routes ...... 23

2.2.5 Surface Functionalisation ...... 24

2.2.6 Applications ...... 25

2.2.6.1 Biosensing ...... 25

2.2.6.2 Chemical Sensing ...... 27

2.2.6.3 Imaging ...... 28

2.2.6.4 Catalysis ...... 30

2.3 Fluorescence In Situ Hybridisation ...... 32

2.3.1 Mechanism of FISH ...... 33

2.3.2 FISH Applications ...... 36

2.3.2.1 Gene Positioning ...... 36

2.3.2.2 Multicolour-FISH ...... 37

2.3.2.3 Diagnosing Chromosomal Abnormalities...... 38

2.3.2.4 Diagnosing Diseases ...... 39

3 Synthesis and Characterisation of Carbon Dots ...... 42

3.1 Materials and Instruments ...... 42

3.1.1 Materials...... 42

3.1.2 Instruments ...... 42

3.2 Hydrothermal CD Synthesis...... 42

3.3 Optical Measurement ...... 44

3.4 Optimisation of Parameters for CDs Synthesis ...... 46

3.4.1 Temperature ...... 46

3.4.2 Duration of Synthesis ...... 46

3.4.3 Effect of Acid Ratio ...... 46

3.4.4 CMC Concentration ...... 47

3.5 External Effects ...... 47

3.5.1 Effect of Metal Ions ...... 47

3.5.2 pH ...... 48

3.6 Results and Discussion ...... 49

3.6.1 Optical Properties ...... 49

3.6.1.1 Neutralisation and Isolation of CDs ...... 49

3.6.1.2 SEM Analysis ...... 50

3.6.1.3 FTIR Analysis ...... 51

3.6.2 Optimisations ...... 53

3.6.2.1 Temperature ...... 53

3.6.2.2 Duration of Synthesis ...... 56

3.6.2.3 Effect of Acid Ratio ...... 59

3.6.2.4 CMC Concentration ...... 61

3.6.3 Effect of External Surroundings ...... 63

3.6.3.1 Effect of Metal Ions ...... 63

3.6.3.2 pH ...... 68

3.7 Conclusions ...... 69

4 Conjugating Carbon Dots with GAPDH Probe ...... 70

4.1 GAPDH Probe Design ...... 70

4.2 CD-aniline EDC-NHS Conjugation ...... 71

4.3 CD-oligonucleotide EDC-NHS Conjugation ...... 74

4.4 Conclusions ...... 75

5 CD-FISH ...... 76

5.1 Preliminary Testing for CDs Imaging ...... 76

5.1.1 Webcam and Smartphone Camera ...... 76

5.1.2 Microscope Camera ...... 78

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5.2 HaCaT Cell Culture ...... 79

5.2.1 Reviving Cryo-preserved HaCaT Cells ...... 79

5.2.2 Sub-culturing HaCaT Cells ...... 80

5.3 Fluorescence In Situ Hybridisation ...... 80

5.3.1 Fixation of Cells ...... 81

5.3.2 Hybridisation ...... 81

5.4 Results and Discussion ...... 82

5.4.1 Preliminary Testing for CD Imaging ...... 82

5.4.2 Fluorescence In Situ Hybridisation...... 83

5.5 Conclusions ...... 87

6 Conclusion ...... 88

6.1 Summary ...... 88

6.2 Future Directions ...... 89

References ...... 90

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List of Figures Figure 2.1 Schematic representation of a double focussing stage scanning microscope. Light source A shines through pinhole B. The light is subsequently focused via objective lens C to a diffraction limited at spot D in the specimen. Then, light from spot D is collimated by a second objective lens E which focuses the light onto the exit pinhole F. Phototube G detects the light coming out of this pinhole. [24] ...... 7 Figure 2.2 The principle of fluorescence. Fluorescence phenomenon represented schematically in the classical Bohr model. From the ground state, absorption of the light quantum (blue) makes an electron move to a higher energy orbit. After residing in this “excited state” for a particular time, an occurrence termed the “fluorescence lifetime”, the electron falls back into its original orbit and the excess energy is dissipated by the fluorochrome through emission of a photon. [1] ...... 7 Figure 2.3 Single photon excitation. (A) Epifluorescence microscopy: The sample is irradiated with an excitation light under a fluorescence microscope and the fluorescence signal from sample is detected through the same objective lens. (B) Jablonski diagram: A quantum of light (single photon) excites an electron from its ground state GS0 (electronic singlet) to a higher state and then loses energy as the electron quickly relaxes to a lower vibrational excited state (orange line). Fluorescence is emmitted when the electron dissipates the remaining energy by emitting a longer wavelength photon as it returns to the ground state. The electrons’ spins in the singlet states (paired or unpaired anti-parallel spins) compared to the triplet state (unpaired, parallel spin) are illustrated. Notice that spin conversion is required for intersystem crossing from ES1→ET1. [1]...... 10 Figure 2.4 Intrinsic biochemical fluorophores. R represents a hydrogen in NADH and a phosphate group in NADPH. [27] ...... 11 Figure 2.5 The function of sodium fluorescein concentration is interpreted by calculating the illumination absorptance, fluorescence efficiency, and intensity of fluorescence. Intensity is calculated as the product of absorptance and efficiency (k=1) in the equation I(c)=kA(c)E(c). [36] ...... 14

Figure 2.6 Absorbance and fluorescence emission of fluorescein. Stokes shift is the difference, in nanometres, between the excitation maxima and emission maxima...... 14 Figure 2.7 Comparison of photostability of antibodies labelled with 5 types of dyes in cells on a fixed slide. A fluorescence microscope was used to measure the intensities [27, 38]...... 15 Figure 2.8 A schematic representation of a core-shell configuration of a typical quantum dot, with CdSe or CdS core and shell typically composed of ZnS, CdS or ZnSe...... 16 Figure 2.9 Scheme of bandgap changing of CDs, m-Cds and r-CDs. [67] ...... 19 Figure 2.10 Schematic diagram of Pi detection based on the off-on fluorescence probe of CDs adjusted by Eu3+. [98] ...... 25 Figure 2.11 A schematic illustration of the CD-based fluorescent nucleic acid detection. [99] ...... 26 Figure 2.12 Fluorescent images of HeLa cells clamped at pH 6.0, 6.5, 6.8, 7.2, 7.5, and 8.0, respectively. The images of the first row (FITC channel) and second row (RBITC channel) were collected in the ranges of 510–550 nm and 570– 610 nm, respectively. The third row shows the corresponding differential interference contrast images. Olympus software (FV10-ASW) was used to generate the images of the fourth row (the ratio channel). The bottom colour strip represents the pseudocolour change with pH. Scale bar, 20 mm. [104] ...... 27 Figure 2.13 Top: Images from histopathological analyses of liver, spleen, and kidneys. Bottom: Fluorescence images (two-photon excitation at 800 nm) of sliced liver and spleen harvested from mice 6 h after intravenous exposure to C- dots. [58] ...... 29

Figure 2.14 Possible catalytic mechanism of the TiO2–CD nanocomposites under visible light. [75] ...... 31 Figure 2.15 Base pairing in DNA. T and A are connected by two hydrogen bonds whereas G and C are connected by three hydrogen bonds. The sugar- phosphate backbones (grey) run anti-parallel to each other, so that the 3’ and 5’ ends of the two strands are aligned. [131] ...... 34

Figure 2.16 Principles of fluorescence in situ hybridisation (FISH). (a) A DNA probe and a target sequence make up for the basic elements of FISH. (b) Prior to hybridisation, the DNA probe can be labelled through different methods such as nick translation, random primed labelling and PCR. The most common labelling strategies used are indirect labelling (left panel) and direct labelling (right panel). Indirect labelling uses probes that are labelled with modified nucleotides that contain a hapten, while direct labelling utilises nucleotides that are already have a fluorophore attached. (c) Denaturation of labelled probe and target DNA. (d) When combined together, the denatured probe anneals to complementary target DNA sequences. (e) For the probe that is indirectly labelled, the nonfluorescent hapten is visualised by an additional step that uses an enzymatic or immunological detection system. [132] ...... 35 Figure 2.17 (a) Fluorescent signals are seen at cytogenetic bands (grey) where fragments of a sequence-tagged BAC hybridise (red) via FISH. (b) The breakpoint of a translocation involving chromosomes 11 and 19 in a patient with multiple congenital malformations and mental retardation (DGAP012) were mapped out using clones selected based on band location. Clone CTD- 3193o13 spans the breakpoint on chromosome 19 where the red signal is split between the derivative chromosome 11 and derivative 19 chromosomes. The red signal is also present on the normal chromosome 19. The GTG-banded karyotype for this patient is 46,XY,t(11;19)(p11.2;p13.3). [133] ...... 36 Figure 2.18 Spectral karyotyping and multicolour-FISH paint each human chromosome in one of 24 colours. Cytogenetic localization of DNA sequences via FISH. [136] ...... 38 Figure 2.19 Interphase FISH for uncultured amniocytes (right) using a fluorescent probe for chromosome 21 (orange), and chromosome 13 (green) showing 3 signals of chromosome 21 giving the diagnosis of trisomy 21 or Down syndrome. The karyogram (left) done on cultured amniotic fluid cells confirms the FISH diagnosis. [137] ...... 39 Figure 2.20 Detection of EBER by QD-FISH. NPC tissue sections were subjected to QD-FISH and observed under blue light excitation in a fluorescence

microscope. (A–C) NPC specimens showed positive signals for EBER. (D) In the hybridization mixture containing EBER oligonucleotides without biotin, negative signals were detected. Original magnifications: A, D: 200×; B, C: 400×. [138] ...... 40 Figure 2.21 Fluorescence observed in the PK-FISH assay with the P. knowlesi-specific probe (green) and the Plasmodium genus-specific probe (orange) on monkey blood smears containing P. knowlesi. Dual colour fluorescence seen in a single microscope field at ×1000 magnification using the two different filters is shown in each set of paired photographs. The four sets of paired photographs are from four different fields (i) - (iv). Scale-bars: 5 μm. [139] ...... 41 Figure 3.1 Chemical structure of sodium carboxymethylcellulose...... 43 Figure 3.2 Salt removal from CDs solution using salting out extraction. (a) Two layers formed after mixing acetone with the neutralised CD solution. The top layer consists of acetone mixed with CDs while the bottom layer is salt. (b) Salt precipitates filtered out from the CDs solution. (c) Isolated CDs layer mixed in water and acetone. (d) Solid CDs after drying out in oven at 80°C overnight...... 45 Figure 3.3 Emission profile of neutralised CDs (pH 7) in acetone solution after salt precipitates were filtered out...... 49 Figure 3.4 Emission profile of CDs reconstituted in water (30.0 mg/mL) after neutralisation, isolation and drying process...... 50 Figure 3.5 (a) SEM image and (b) size distribution of CDs after neutralisation and isolation. Mean particle size= 0.194 µm. Standard deviation= 0.187 µm. The size distribution represents 95% of measured particles...... 51 Figure 3.6 (a) SEM image and (b) size distribution graph of CDs after neutralisation and isolation prepared on a different day using the same parameters but with an extra filtration step using 0.22 μm syringe filter after neutralisation. Mean particle size= 0.134 µm. Standard deviation= 0.055 µm. The size distribution represents 100% of measured particles...... 51 Figure 3.7 FTIR spectra of the neutralised and isolated CDs...... 52 Figure 3.8 Filtered raw CDs solutions synthesised at different temperatures of 100, 150, 200, 250 and 300°C under (a) normal light and (b) UV light...... 54

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Figure 3.9 Fluorescent emission spectra of CDs synthesised at (a) 150°C; (b) 200°C; (c) 250°C and (d) 300°C at their optimum emission peaks...... 55

Figure 3.10 The effect of temperature on the emission of CD at at λex = 400 nm, λem = 595 nm...... 56 Figure 3.11 Filtered raw CDs solutions synthesised with heating durations of 1.0, 1.5, 2.0, 2.5 and 3.0 h for time optimisation under (a) normal light and (b) UV light...... 57 Figure 3.12 Fluorescent emission spectra of CDs synthesised at (a) 1.0 h; (b) 1.5 h; (c) 2.0 h; (d) 2.5 h and (e) 3.0 h, at their optimum emission peaks...... 58

Figure 3.13 The effect of heating time on the emission of CD at λex = 400 nm, λem = 595 nm...... 59 Figure 3.14 Fluorescent emission spectra of CDs synthesised with phosphoric acid concentrations of (a) 8.5 %; (b) 17.0 %; (c) 25.5 %; (d) 34.0 % and (e) 42.5%, at their optimum emission peaks. Cary Eclipse emission and excitation slit settings used for (a), (c), (d) and (e) is 10 nm while for (b) the slit size used is 5 nm...... 60 Figure 3.15 The effect of phosphoric acid concentration on the emission of CD at λex = 400 nm, λem = 595 nm...... 61 Figure 3.16 Fluorescent emission spectra of CDs synthesised from CMC of increasing concentrations (a) 2.5 mg/mL; (b) 5.0 mg/mL; (c) 7.5 mg/mL; (d) 10.0 mg/mL and (e) 12.5 mg/mL, at their optimum emission peaks...... 62 Figure 3.17 The effect of concentration of CMC on the emission of CDs at λex = 400 nm, λem = 595 nm...... 63 Figure 3.18 Comparisons of CD samples added with 50 µL water and various metal ions solution under (a) 450 nm, (b) 490 nm, and (c) 595 nm emission wavelengths...... 64

Figure 3.19 Normalised intensity of CD at λex=390 nm, λem=490 nm against

concentration of Al(NO3)3...... 65

Figure 3.20 Normalised intensity of CD at (a) λex=350 nm, λem=450 nm and (b) λex=390

nm, λem=595 nm against concentration of Fe(NO3)3...... 66

Figure 3.21 Normalised intensity of CD at (a) λex=350 nm, λem=450 nm and (b) λex=390

nm, λem=450 nm against concentration of SnCl2...... 67 Figure 3.22 Graph of pH curve of CDs at λex = 400 nm, λem = 595 nm...... 68

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Figure 4.1 Chemical structure of aniline...... 71 Figure 4.2 Schematic of Sulfo-NHS plus EDC (carbodiimide) crosslinking reaction [168]...... 73 Figure 4.3 FTIR spectra of CD-aniline, aniline and CD...... 74 Figure 5.1 (a) Nikon Eclipse Ti inverted microscope. (b) Placement of 390 nm UV torchlight above condenser unit...... 77 Figure 5.2 Images of onion cell epidermis after incubation with CDs solution. Top: Onion cells image under white light (left) and UV light (right) taken with Logitech webcam. Bottom: Onion cells image under white light (left) and UV light (right) taken with iPhone SE camera...... 78 Figure 5.3 Nikon Eclipse Ti inverted microscope with Nikon SMZ 745T microscope camera installed...... 79 Figure 5.4 Fluorescence images of onion epidermis incubated in (a) distilled water, (b) highlighter dye, and (c) 20.0 mg/mL CDs solution illuminated under UV light. Photos were taken with Nikon SMZ 745T microscope camera...... 82 Figure 5.5 A comparison of fluorescence images of fixed HaCaT cells incubated in (a) hybridisation medium without probe and (b) 15.0 mg/mL CD-GAPDH probe viewed in mounting oil without p-phenylenediamine and with p- phenylenediamine. All images were taken under the same exposure settings...... 83 Figure 5.6 Fluorescence images of fixed HaCaT cells incubated with different samples: (a) hybridisation medium without probe, (b) 15.0 mg/mL CDs, and (c) 15.0 mg/mL CD-GAPDH probe. All images were taken under the same exposure settings...... 85 Figure 5.7 A zoomed in image comparison between a) CDs and b) CD-GAPDH probe in HaCaT cells. Arrows are pointed to clusters of bright spots likely to be GAPDH mRNA. Same contrast adjustments were applied on both images to enhance fluorescence signal...... 86 Figure 5.8 Localisation of GAPDH in U-251 MG cell line (left) and A549 cell line (right). The nucleus is stained in blue, microtubules in red and GAPDH in green. [176] ...... 86

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List of Tables Table 3.1 Preparation of different phosphoric acid concentrations...... 46 Table 3.2 Preparation of different CMC concentrations...... 47 Table 3.3 Preparation of CDs in different pH conditions...... 48

1 Introduction

1.1 Brief Overview

Fluorescence in situ hybridisation (FISH) is a powerful technique used for detecting and localising specific DNA or RNA cells or tissues. This method can also reveal the location of a specific nucleic acid on a chromosome. In the field of medicine, molecular cytogeneticists use FISH as a diagnostic tool for detecting gene abnormalities, cancer and for gene mapping. FISH is a form of hybridisation that utilises probe, in which fluorescent tag is labelled to a single-stranded fragment of DNA or RNA that targets to a specific nucleotide sequence. The probes can be tagged with various kind of fluorescent molecules such as organic dyes, quantum dots, phycobiliprotein derivatives and lanthanide chelates [2]. The characteristics of these dyes are different and each has its own advantages and disadvantages depending on the nature of application. For example, quantum dot probes are excellent for their long-term photostability, narrow emission spectra and brilliant colours. However, quantum dots were reported to exhibit toxicity in cells such as causing oxidative stress in plants [3], mitochondria impairment [4] and inducing endothelial toxicity [5]. Thus, there is a need to expand the choice of fluorophores in hybridisation probes to suit research needs.

A new type of carbon nanoparticle called carbon dots (CDs) has recently caught the attention of researchers for being a good alternative to current commercially available fluorophores. CDs are reported to have optical properties similar to that of QDs and are generally non-toxic [6]. Although many studies on bioimaging of cells incubated with CDs have been done [7-9], so far their utilisation as hybridisation probes in FISH is still very limited [10].

1.2 Problem Statement

The increasing demand and use of biomarkers and fluorescent tags has called for the need of developing suitable fluorophores to suit an intended application. There is already a wide range of fluorescent probes available commercially. However, most of these probes are either too expensive, limited in application, or toxic to health or environment. In bioimaging, it is important to have non-toxic, biocompatible fluorescent tags that are low-cost and easily available for in vivo and in vitro 1 applications. CDs are a versatile class of fluorescent nanoparticles with these properties and show great potential as a fluorescent label for a probe. So far, studies on CDs in in situ hybridisation techniques are very limited and need more attention to ensure that CDs can be fully utilised for monitoring purposes. Thus, there is a real need to explore the use of CDs in FISH applications.

1.3 Thesis Aim

The main aim of this study is to synthesise CDs that are suitable for use in FISH application on cells.

1.4 Objectives

CDs were utilised as fluorescent probes by conjugating to an oligonucleotide probe for specific gene detection in cells. This study explores on the use of the versatile CDs in bioimaging and FISH techniques by carrying out the following objectives:

1. To synthesise CDs with carboxyl surface groups and red-shifted emission. Additionally, the CDs will be isolated, and characterised via fluorescence spectroscopy, SEM and FTIR analyses. 2. To design an oligonucleotide probe conjugated with isolated CDs. 3. To assess the potential of the designed CD-oligonucleotide probe in fixed cultured cells for FISH application by using fluorescence microscopy.

1.5 Thesis Outline

This thesis is divided into 6 chapters. The experimental section, results, and discussions of this study are divided into 3 main stages namely: synthesis and characterisation of carbon dots, conjugating carbon dots with GAPDH probes, and CD-FISH, which are covered in chapter 3, 4, and 5 respectively.

Chapter 1

Introduction of the motivation and objectives of this study.

Chapter 2

A comprehensive literature review on subjects detailing the general background and principle of fluorescence, fluorophores and their applications. This chapter emphasises on the physicochemical properties, synthetic methods, surface functionalisation and applications of CDs. Additionally, principles of FISH and its applications are reviewed.

Chapter 3

This chapter presents the experimental methods for the synthesis of CDs including the optimisation of synthesis parameters and isolation of CDs. Subsequently, characterisation studies of the CDs including effects of metal ions, pH test, SEM analysis, and FTIR analysis are described. This is followed by the results and discussion of each experimental study.

Chapter 4

This chapter describes the design of GAPDH oligonucleotide probe followed by conjugation with isolated CDs using carbodiimide chemistry. Trial conjugation w done using aniline as a substitute for the GAPDH probe. The confirmation of successful conjugation between CDs and aniline using FTIR analyses are presented and discussed.

Chapter 5

This chapter includes the preliminary set-up for testing the imaging of CDs in plant cells and experimental methods for HaCaT cell culture and FISH. HaCaT cell line was incubated with the CD-oligonucleotide probes and subsequently imaged for fluorescence signal. Fluorescence imaging results are presented and discussed at the end of this chapter.

Chapter 6

This final chapter concludes the thesis with a summary of the experimental results obtained and contribution of this study to the biosensing application of CDs. This is followed by recommendations of potential further works for this research project.

1.6 Scope and Delimitation

This study focuses on the synthesis of CDs suitable for probe labelling and its application as a biosensor, particularly as a FISH probe. CDs were synthesised via hydrothermal method with the addition of acid to red-shift the resultant CDs emission to a colour that is preferable for bioimaging analyses. The carbon source used for synthesis was obtained from a sustainable and non-toxic origin that also contains carboxylic functional groups to create carboxyl-functionalised CDs.

CDs were neutralised and cleaned from impurities via salting out extraction using acetone. Then, CDs were conjugated to an oligonucleotide designed to detect GAPDH mRNA. This conjugation was performed using 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) and N-hydroxysuccinimide (NHS) crosslinking chemistry. The CD-GAPDH probe was subsequently used for FISH on HaCaT cell lines and imaged under a fluorescence microscope.

Certain general characterisations of CDs may not be available due to limitations of facilities. Equipment for fluorescence microscopy was also not available and limited appointments had to be made to borrow this equipment at another institution. Thus, an initial attempt for imaging of CDs using an available inverted microscope was performed. In addition to FISH studies, the scope of this project also includes the sensing potential of CDs for metal ions and pH change.

2 Literature Review

2.1 Fluorescence Bioimaging

Bioimaging refers to techniques or methods for visualisation of biological processes in real time with the aim of being as non-invasive (no cutting of the body or isolating cellular components) as possible [11]. Studies done using bioimaging can span from an entire cell and its subcellular structures up to tissues and a whole multicellular being. Some examples of imaging technologies applied for bioimaging are X-ray, computed tomography (CT), positron emission tomography (PET), magnetic resonance imaging (MRI), magnetoencephalography (MEG), light microscopy and fluorescence microscopy. From clinical diagnosis to the study of cellular biology, the importance of bioimaging in biomedical and biological sciences is unprecedented.

Recent advances in bioimaging has seen increasing popularity with the use of fluorescence imaging on account of its advantages such as providing a safe, highly sensitive and less invasive method of detection. This is performed using a wide range of readily available fluorescence imaging instruments. Sir Frederik William Herschel first reported the fluorescence optical phenomenon in 1845 in which he observed that the quinine solution that is usually colourless and transparent showed “vivid and beautiful celestial blue colour” when exposed under a certain way under the sunlight [12]. Decades later, limitations of light microscope were demonstrated [13] and this transpired the notions that better differentiated images could be obtained if the object of study are self-illuminating [14]. The first fluorescence microscopes came about in the 1910s, which were produced by two companies; Carl Zeiss and Carl Reichert [15-17]. One of the earliest approach to observe fluorescence in a large and opaque organism were done in 1929 by Ellinger and Hirt where they illuminated UV light on living organisms that had been treated with fluorescent substances and observed using an “intravital microscope” that they had devised [18]. Since then, labelling specimens has evolved to be more specific and efficient using fluorescent probes that utilise antibody labelling [19], in situ hybridisation (ISH) [20], cloning of the green fluorescent protein (GFP) gene [21] and development of different variants of GFP that genetically encodes into protein of interest to label them [22]. As various labelling techniques have been well established, fluorescence microscopy is the preferred method of choice for imaging

5 live specimens and it is currently the standard procedure to study normal and pathological cell biological processes.

In biology, clear visualisation of specific target in samples is very important and has been the reason for major scientific discoveries. This has driven researchers to enhance the quality and resolution of fluorescence microscopy images. Resolution or optical resolving power in fluorescence microscopy determines the amount of details that can be observed in a specimen [1]. The resolution itself is determined by the limitations of the instrument and other physical factors. It is important to note that resolution in fluorescence microscopy is not directly governed by magnification. Also, contrast and resolution are interrelated parameters that should not be viewed as two different entities. In fact, the contrast enhancement and selectivity in fluorescence microscopy are what make it a great and powerful tool for imaging. Selectivity is achieved through the labelling methods mentioned previously while the increase in contrast is provided by the microscope itself. To obtain a clear and sharp image, the objective of the microscope must capture as much of the emitted fluorescent light as possible while minimizing the capture of incident excitation light. The contrast of an image obtained heavily relies on the microscopes ability to direct fluorescent light to a detector while blocking excitation light.

When using a conventional microscope, the lens will collimate the light coming from the sample and focus it towards the eye. However, light from out of plane of the object enters the eyepiece and this out of plane light causes blurriness in the perceived image. In the 1950s, Marvin Minsky invented the confocal microscope (patented as ‘double focusing stage scanning microscope’) that would eliminate the blurred effect and improve resolution of fluorescence images (Figure 2.1) [23]. In his invention, a pinhole apertures was introduced into the optical path of a microscope set-up. The pinhole was placed in front of the light source and another one in front of the detector. The modern microscopes today are based on this “double focusing system” created by Minsky.

Figure 2.1 Schematic representation of a double focussing stage scanning microscope. Light source A shines through pinhole B. The light is subsequently focused via objective lens C to a diffraction limited at spot D in the specimen. Then, light from spot D is collimated by a second objective lens E which focuses the light onto the exit pinhole F. Phototube G detects the light coming out of this pinhole. [24]

2.1.1 Fluorescence and Fluorophores The fluorescence phenomenon is a form of luminescence in which a substance reemits light from its excited state after absorbing light or radiation of a shorter wavelength [1]. Substances or compounds that display fluorescence are generally referred to as fluorescent probes or dyes and the more appropriate term is ‘fluorochrome’. Although the terms ‘fluorochrome’ and ‘fluorophore’ are often used interchangeably, when taken strictly the term ‘fluorophores’ refers to fluorochromes that are conjugated to biological macromolecules.

Figure 2.2 The principle of fluorescence. Fluorescence phenomenon represented schematically in the classical Bohr model. From the ground state, absorption of the light quantum (blue) makes an electron move to a higher energy orbit. After residing in this “excited state” for a particular time, an occurrence termed the “fluorescence lifetime”, the electron falls back into its original orbit and the excess energy is dissipated by the fluorochrome through emission of a photon. [1]

Fluorescence is a process that results in the emittance of a photon with a longer wavelength. When a particular wavelength of light hits a fluorescent compound, the atom, ions or molecules absorb a specific quantum of light (Figure 2.2). This causes a valence electron to be excited from its ground state GS0—the initial state in which all the electrons have opposite spin and the net spin is 0—into a higher energy level at its excited state ESn. This fast process happens in the matter of femtoseconds and requires the energy of at least ΔE = EESn − EGS0 to bridge the gap between the excited and ground states to induce the excitation. Planck’s Law describes the energy of photons involved in fluorescence and a quantum of light can be expressed via the following equation [25]:

= . = . � � ℎ � ℎ where E is the quantum’s energy (J), h is Planck’s �constant (J.s), v is the frequency (s-1), λ is the wavelength of the photon (m) and c is the speed of light (m.s-1). From Planck’s Law, energy of an absorbed photon is directly proportional to the frequency and inversely proportional to the wavelength. In other words, shorter incident wavelengths possess greater quantum energy. Depending on the fluorescent species, the excited state sublevels or vibrational levels reached will be different.

When under UV or visible light, common fluorophores generally get excited to higher vibrational levels of the first (ES1) or second (ES2) singlet energy state. Referring to Figure 2.3B, an absorption (left-hand blue arrow) occurs from the lowest vibrational energy level of the ground state to a higher vibrational level in the second excited state

(transition is denoted as GS0=0 to ES2= 3). The second absorption (right-hand blue arrow) transition starts from the second vibrational level of the ground state to the highest vibrational level in the first excited state (denoted as GS0=1 to ES1= 5). A typical fluorophore irradiated with a wide spectrum of wavelengths will generate an entire range of allowed transitions that populate the various vibrational energy level of the excited states. Some of these transitions will have a much higher chance of occurrence than others, and when combined, will constitute the absorption spectrum of the fluorophore. For most fluorophores, the absorption and excitation spectra are distinct. However, they often overlap with each other and become indistinguishable. In some cases, fluorescent dyes such as fluorescein, has an absorption spectra that is clearly separated from the excitation spectra.

Instantly after absorption of a photon, multiple processes will occur with varying probabilities but the relaxation to the lowest vibrational energy level of the first excited state (ES1= 0 ; Figure 2.3B) will most likely happen. This process generally occurs in a picosecond or less and is known as internal conversion of vibrational relaxation (loss of energy in the absence of light emission). Due to the significant number of vibration cycles that transpire during the lifetime of excited states, the molecule virtually undergo constant complete vibrational relaxation during their excited lifetimes. The excess vibrational energy is converted into heat which is absorbed by neighbouring solvent molecules.

An excited molecule exists in the lowest excited singlet state (ES1) for periods on the order of nanoseconds (the longest time period in the fluorescence process by several orders of magnitude) before finally relaxing to the ground state. If a photon emission accompanies the relaxation from this long-lived state, this process is formally known as fluorescence. Fluorescence is normally observed as emission intensity over a band of wavelengths rather that a sharp line or peak. This is because the closely spaced vibrational energy levels of the ground state, when coupled with normal thermal motion, produce a wide range of photon energies during emission. Most fluorophores can repeatedly undergo the excitation and emission cycle for hundreds to thousands of time before the highly reactive excited state molecule becomes photobleached, where fluorescence is extinguished. Fluorescein isothiocyanate (FITC) for example, can undergo excitation and relaxation for approximately 30,000 cycles before the molecule no longer fluoresces when irradiated with incident illumination [26].

There are other relaxation pathways that compete with the fluorescence emission process via different probabilities. The energy of the molecule at its excited state can be dissipated non-radiatively as heat or it can collide with another molecule to transfer energy in the second form of non-radiative process known as quenching (represented by the darker blue wavy arrow in Figure 2.3B). Other than the non-radiative processes, the intersystem crossing to the lowest excited triplet state phenomenon (short red arrows in Figure 2.3B) can occur as well. Intersystem crossing happens relatively rarely but ultimately results either in emission of a photon through phosphorescence or a transition back to the excited singlet state that yields delayed fluorescence. According to quantum theory, it is forbidden for the molecule to transition from the triplet excited state to the

9 singlet ground state and thus, the rate constants for triplet emission are several orders of magnitude lower than those for fluorescence.

Figure 2.3 Single photon excitation. (A) Epifluorescence microscopy: The sample is irradiated with an excitation light under a fluorescence microscope and the fluorescence signal from sample is detected through the same objective lens. (B) Jablonski diagram: A quantum of light (single photon) excites an electron from its ground state GS0 (electronic singlet) to a higher state and then loses energy as the electron quickly relaxes to a lower vibrational excited state (orange line). Fluorescence is emmitted when the electron dissipates the remaining energy by emitting a longer wavelength photon as it returns to the ground state. The electrons’ spins in the singlet states (paired or unpaired anti-parallel spins) compared to the triplet state (unpaired, parallel spin) are illustrated. Notice that spin conversion is required for intersystem crossing from ES1→ET1. [1]

2.1.1.1 Intrinsic Fluorophores A very crucial component of fluorescence imaging is the use of fluorescent molecules or fluorophores that are modifiable to create biomolecular probes for specific detection of DNA, mRNA, proteins or molecular structures. Fluorophores are also applicable as biological stains, indicators for pH and small molecule sensors. The potential of fluorophores in bioimaging is boundless, and thus there are countless of research and designing done to create nanosensors and bioprobes. In general, fluorophores are divided into two classes: intrinsic and extrinsic fluorophores [27].

Intrinsic fluorophores are naturally occurring fluorophores with fluorescence that originates from aromatic amino acids tryptophan (trp), tyrosine (tyr), and phenylalanine 10

(phe) which are also known as organic dyes [28]. Example of intrinsic biochemical fluorophores are shown in Figure 2.4. The indole group in tryptophan is responsible for the UV absorption and emission in proteins. Tyrosine and tryptophan has similar quantum yields but what differs them is that the tyrosine’s emission spectrum wavelength is more narrowly distributed. This gives the impression of a higher quantum yield.

Figure 2.4 Intrinsic biochemical fluorophores. R represents a hydrogen in NADH and a phosphate group in NADPH. [27]

Enzyme cofactors are usually fluorescent and are essential for assisting enzymes during catalysis of reactions. The cofactor nicotinamide adenine dinucleotide in its reduced form (NADH), also known as flavin reductase, is an intrinsic fluorophore with absorption and emission peaks at 340 and 460 nm respectively. The group responsible for its fluorescence is the reduced nicotinamide ring. In its oxidised form, NAD+ is non-fluorescent. When NADH binds to proteins, the quantum yield can increase up to fourfold [29] and the lifetime increases to approximately 1.2 ns. This is not always the case, however, as the NADH fluorescence may increase or decrease depending on the type of protein it binds to.

Another fluorescent cofactor is the pyridoxyl phosphate. Pyridoxyl groups are often coupled to lysine residues by the aldehyde groups and depending on its chemical structure in the protein, the absorption and emission spectra of this fluorophore vary [30, 31]. For example, emission spectrum of pyridoxamine is at shorter wavelengths than that of pyridoxyl phosphate. The emission wavelength of pyridoxamine is pH dependent while the emission of pyridoxyl group depends on its interaction with proteins [32]. The fluorescent properties of pyridoxyl groups are complex as this cofactor can exist in many forms.

Riboflavin, flavin mononucleotide (FMN), and flavin adenine dinucleotide (FAD) are intrinsic fluorophores that absorb light in the visible range ( 450 nm) and emit at approximately 525 nm. Opposite to NADH, flavins are fluorescent≅ in their oxidised forms instead of their reduced forms. The lifetimes for FMN and FAD are typically 4.7 ns and 2.3 ns respectively. In NADH, its flavin fluorescence is quenched by the adenine due to the complex formation between the flavin and adenosine [29]. This process is also known as static quenching. There may be other dynamic component that affects the quenching due to collisions between adenine and the reduced nicotinamide moiety. Unlike NADH that gets highly fluorescent upon protein binding, flavoproteins are nonfluorescent or weakly fluorescent although there are certain exceptions.

In general, nucleotides and nucleic acids are nonfluorescent but there are some exceptions. The yeast tRNAPHE contains the Y-base which is a highly fluorescent base with an emission maximum near 470 nm and a lifetime of approximately 6 ns. The molecules that have been described above represent the dominant fluorophores that exist in animal tissues. Other intrinsic fluorophores that are discovered have been summarised [33].

2.1.1.2 Extrinsic Fluorophores Extrinsic fluorophores are synthesised dyes or modified biochemical added to samples to introduce fluorescence with specific emission properties. Often times, molecules of interest are nonfluorescent or have no intrinsic fluorescence such as DNA and lipids. To overcome this, the sample of interest is labelled with extrinsic probes to make them fluoresce. For proteins, it is more desirable instead to label them with chromophores with excitation and emission wavelengths that are longer than the aromatic amino acids. Example of extrinsic fluorophores includes fluoresceins, rhodamines, Alexa fluor dyes, 12 and boron-dipyrromethene (BODIPY). These fluorophores can be labelled to proteins through covalent or non-covalent bonding. Covalent probes have variety of reactive groups to allow coupling with amines and sulfhydryl or histidine side chains in proteins.

Fluoresceins and rhodamines dyes have favourable absorption maxima near 480 and 600 nm and emission wavelengths from 510 to 615 nm respectively [27]. They are not sensitive to solvent polarity and has high molar extinction coefficients near 80 000 M- 1cm-1, thus their use as fluorophores is widespread. Fluorescein and rhodamine are commonly used as label for antibodies as they are already a wide variety of fluorescein and rodhamine-labelled immunoglobulins that are sold commercially for use in fluorescence microscopy and immunoassays. Their high quantum yield and long absorption and emission wavelengths help minimise the problems of background fluorescence from biological samples.

BODIPY dyes are based on an unusual boron-containing fluorophore and are introduced as replacements for fluorescein and rhodamines. A wide range of emission wavelengths, from 510 to 675 nm, can be obtained depending on the BODIPY dye’s structure. BODIPY dyes also displays high quantum yield and are insensitive to solvent polarity and pH. When compared to fluorescein and rhodamine, the BODIPY has narrower emission spectra, so more light is emitted at the peak wavelength, possibly allowing the resolving of more individual dyes. These dyes however, has very small Stokes shift and the spectral overlap results in a Förster distance of about 57 A [34]. Small stokes shift are associated with self-quenching of fluorophores.

A disadvantage of fluoresceins, rhodhamines, and BODIPYs is their tendency to self- quench. Fluorescein-labelled proteins’ brightness is known to decrease in intensity as the extent of labelling increases (Figure 2.5). The effect of fluorescein concentration, c, on fluorescent intensity, I(c) can be explained by the equation:

( ) = ( ) ( ) where constant k is independent of� concentration.� �� When� absorptance, A(c), increases to a maximum of unity at high concentrations, the efficiency, E(c), decreases to low levels at high concentrations as a result of self-quenching phenomenon [35].

Figure 2.5 The function of sodium fluorescein concentration is interpreted by calculating the illumination absorptance, fluorescence efficiency, and intensity of fluorescence. Intensity is calculated as the product of absorptance and efficiency (k=1) in the equation I(c)=kA(c)E(c). [36]

This phenomenon can also be understood by small Stokes shift displayed by the examination of the fluorescein excitation and emission spectra (Figure 2.6). When there are multiple fluorescein groups attached to a protein, energy transfer could happen between them. Two fluorescein groups that are attached to the same protein are likely to be with the Förster distance that allows energy transfer, which is within 40 Å of each other. In other words, a high local fluorescein concentration is resulted from having multiple fluorescein groups bound to a protein.

Figure 2.6 Absorbance and fluorescence emission of fluorescein. Stokes shift is the difference, in nanometres, between the excitation maxima and emission maxima.

Most fluorophores experience photobleaching when exposed to continuous illumination such as in fluorescence microscopy where high intensity of light is subjected on them. One of the most useful and important properties of a probe is photostability. Fluorescein makes one of the least photostable dyes (Figure 2.7). Alexa Fluor dyes are a series of dyes created by a biotechnology company named Molecular Probes [37]. They come in a large range of emission maxima of 442-814 nm and were developed to be more photostable than conventional fluorescent dyes. However, the photostability of these dyes can be affected by their local environment.

Figure 2.7 Comparison of photostability of antibodies labelled with 5 types of dyes in cells on a fixed slide. A fluorescence microscope was used to measure the intensities [27, 38].

2.1.1.3 Quantum Dots Luminescent semiconductor nanocrystals or quantum dots (QDs) are nanosized semiconductor particles made up of elements from the periodic group of II-VI or III-V and have attractive optical properties that are controlled by the particle size, constituent material, size distribution and surface chemistry [39, 40]. QDs are sized at diameters of 1-6 nm [41]. The most common constituents of QDs used in life science applications are cadmium selenide (CdSe) and cadmium telluride (CdTe). Other possible alternatives include indium phosphide (InP) and indium gallium phosphide (InGaP) from the III/V group which do not contain cytotoxic cadmium ions. The quantum yields, decay kinetics, and stability of the QDs are determined by the number of dangling bonds on 15 the core particle surface. To optimise these features, inorganic passivation layers or organic capping ligands or both are attached to the particle surface [42]. Thus, QDs are typically a semiconductor core (e.g., Cd and Se) surrounded by a semiconductor shell such as zinc sulfide (ZnS) or just a core functionalised with different coatings such as visualised in Figure 2.8. The method of particle synthesis and surface modification plays a huge part in changing the properties of the QDs core-shell. Passivation shell on QDs often causes a slight red-shift in the absorption and emission due to the tunnelling of charge carriers into the shell.

Figure 2.8 A schematic representation of a core-shell configuration of a typical quantum dot, with CdSe or CdS core and shell typically composed of ZnS, CdS or ZnSe.

A special property of QDs is their broad excitation wavelength and narrow symmetric emission spectra. This meant that different QDs can be excited by one wavelength shorter than their emission wavelength. Hence, QDs are unlike conventional organic fluorophores, of which the narrow excitation and broad emission are likely to result in spectrum overlap or red tailing [42]. Their inorganic composition makes them more resistant to photobleaching [43] and also have longer fluorescence half-life than typical organic dyes [44].

The application of QDs in in vitro experiments have been reported, for example, in the detection of the cancer cell marker for human epidermal growth factor receptor 2 (Her2) on the surface of fixed and live cancer cells using immunofluorescent labelling with CdSe/ZnS core-shell QDs [45]. QDs also have been used a cellular markers are they can be internalised by cells through the means of a receptor [46] or by non-specific endocytosis [47].

In addition, QDs have been exploited for in vivo studies such as the study of the behaviour of specific cells during the early stage embryogenesis in Xenopus [48] and Zebrafish embryos by microinjection of micelle-encapsulated QDs [49]. Other than that, antibody-conjugated QDs were developed for in vivo human prostate cancer targeting and imaging [50].

As the use of QDs became more widespread in biological studies, more researchers started to explore the possibilities of using QDs fluorophores for specific detection of genes in FISH techniques [51]. In order to achieve this, the QDs were first capped before labelled with a biomolecule using either a direct or indirect method. In 2005, Chan and his team developed a method for direct QD labelling oligonucleotide probes via streptavidin and biotin interactions to target specific mRNAs in mouse brainstem sections [52]. The oligonucleotide probes were labelled with biotin and then conjugated with QD-streptavidin in the presence of biocytin to block excess streptavidin sites that could cause cross-linking of oligonucleotides.

Despite their great properties as fluorophores, QDs have been reported to show toxic and cytotoxic effects. The metalloid core of QD is toxic when presented as composite core (e.g., CdTe) or upon dissolution of the QD core to its constituent metals (e.g. Cd). Furthermore, some of the coating materials on the QDs such as mercaptoacetic acid were discovered to be cytotoxic. When in vivo, the degradation of the QD coating may also result in unwanted reaction of the QD. In short, the toxicity of QD depends on multiple factors derived from the physicochemical properties of the QD and also its environmental conditions [53]. Another limitation of the QD is its blinking mechanism, which is a fluctuation of photoluminescence between bright (on) and dark (off) states [54] [55]. During its ‘off’ state, the excited electrons of the QD relax in a way that produces heat rather than light. The turning on and off of the QD light emission which seemingly happens at random adversely affects its use in bioimaging applications.

2.2 Carbon Dots

2.2.1 Introduction to Carbon Dots The discovery of carbon nanoparticles such as graphene, carbon dots, carbon nanotubes, and their unique properties have driven an extensive research on their characteristics and potential applications. A newly discovered class of carbon nanoparticles is the carbon dots (C-dots or CDs) which are nano-sized carbonaceous particles, usually in sizes below 10nm [56]. These were first discovered by accident during the purification of single-walled carbon nanotubes carried out by Li and co-workers [57]. The interesting optical properties, fluorescent properties, physicochemical characteristics, and potential applications of CDs have yet to be fully untapped and are still being widely explored since their discovery.

CDs are often compared with another similar type of nanomaterial, QDs. The bright and tunable fluorescence properties of QDs make them widely applied in biosensing and bioimaging applications and are favoured over conventional organic dyes. However, QDs have some limitations such as being toxic to cells due to the use of heavy metals in the core shell which renders them unsuitable for certain biological applications [53] and they also have the tendency to blink. Thus, the discovery of CDs which are similar in its fluorescent properties with QDs without the apparent toxicity has driven the exploitation of CDs as an alternative fluorophore in bioimaging studies [58]. CDs are also reported to be stable against photobleaching and poses no blinking effect [59].

2.2.2 Physicochemical Properties CDs usually have an optical absorption in the UV region, with a tail extending out into the visible region and they commonly have a band centered on 250-300 nm that is known as the π-π* transition peak. Interestingly, CDs synthesised from different methods will have different optical properties. For example, glucose based CDs prepared via hydrothermal, microwave thermal, and ultrasonic methods has various absorption ranging from 250-300 nm [60-63]. CDs prepared through top-down methods usually exhibit size-dependent absorption ranging from 6.20 to 4.92 eV (200-252 nm) as the particle size is increased from 12 to 22 nm [64] [65]. Other than that, the surface functional groups of CDs may also determine the absorption wavelength. Such is the

18 case in a work demonstrated by Tetsuka and co-workers where a red-shift of the CDs was observed after functionalisation with amino groups [66].

Despite the numerous studies carried out to understand the physicochemical properties of CDs, the mechanism behind the photoluminescence (PL) is still being discussed to date. There are two widely accepted theories to explain the origin of PL of CDs: defect state emission (surface energy traps) and intrinsic state emission (electron-hole recombination, quantum size effect/zig-zag sites) [67]. As shown in Figure 2.9, the blue emission (shorter wavelength) is from the intrinsic state emission and the green emission (longer wavelength) originates from the defect states, confirmed by the emission colours of CDs grafted with amino functional groups (m-CDs) and CDs reduced by sodium borohydride (NaBH4) (r-CDs). The r-CDs emit at the UV region due to the elimination of the carboxylic functional groups. This resulted in the decrease of their surface state and the dominant PL now originated from the intrinsic state, which was further amplified by the PL of pristine CDs that has nearly no surface oxygen groups (defects) [68]. It has also been implied that PL from intrinsic state decayed (t < 5 ns) faster than that of the defect states (10 ns > t > 5 ns) [67, 68].

Figure 2.9 Scheme of bandgap changing of CDs, m-Cds and r-CDs. [67]

Governing the PL properties of CDs by controlling the surface functional groups (defect states) is easier than by modifying the core composition (intrinsic states). In other words, it is efficient to remove non-radiative electron-hole recombination centers such as 19 epoxy/carboxylic acid groups and improve quantum yield (QY) by reduction and passivation of CDs’ surface [69]. Passivating agents are usually amino-containing molecules or polymers such as 4,7,10-trioxa-1,13-tridecanediamine (TTDDA), polyethylene glycol (PEG), polyethylenimine (PEI), octadecylamine, 1-hexadecylamine, and others [70]. Surface passivation with these polymers or organic molecules are required for the enhancement of PL intensity, for example, CDs before and after passivation with TTDDA showed QYs of 1% and 13% respectively [60]. The passivation process can be simplified from two steps to one step through the proposed methods of co-pyrolysis or one-pot hydrothermal route of carbon precursors and passivation agents such as ammonia solution [66], branched PEI [71], dimethylformamide [72], and ethylenediamine [73].

CDs were also revealed to have size-dependent PL properties as demonstrated by Kim and co-workers in 2012, when the size of CDs is increased from 5 to 17 nm the emission wavelength shifts from 450 nm (2.75 eV) to 486 nm (2.55 eV) [65]. Furthermore, CDs showed pH dependent PL properties which suggests that their PL may originate from free zigzag sites with a carbine-like triplet ground state [74]. In CDs, PL originates from the transition of the particle’s lowest unoccupied molecular orbital (LUMO) to the highest occupied molecular orbital (HOMO). The HOMO-LUMO gap changes according to the size of the fragments. Based on a study performed by Li et al., (2010), if the gap increases inversely proportional to the increases in the size of C-dots, various sizes of C-dots would have different excitation and emission spectra [75]. Moreover, smaller CDs tend to have stronger PL intensity.

Regardless of the method of preparation, most of the reported CDs possess oxygen containing group including carbonyl, epoxy/ether, and carboxylic acid on their surfaces [70]. The surface states of CDs are formed from these functional groups, with energy levels between π and π* states [62]. When CDs were oxidised through the supply of high voltage, the oxygen-related surface increased and resulted in the red-shift of the maximum emission wavelength [76]. On the other hand, CDs reduced with NaBH4 causes a blue-shift of the maximum emission wavelength from 520 to 450 nm and an increase in QY [77]. It has also been discovered that doping CDs with nitrogen and sulfur to create new surface states could efficiently shift the PL emission and increase QY [78].

2.2.3 Top-down Approaches The synthetic methods for CDs can be divided into two categories, namely the top- down and bottom-up synthetic routes. Depending on the method of synthesis and carbon source, CDs synthesised are generally rich with surface functional groups such as carboxyl, hydroxyl, amino, and others that can help with further modifications to conjugate with proteins or other compounds. The top-down methods involve the cleavage of larger carbonaceous materials such carbon nanotubes, graphite, and activated carbon into smaller pieces, usually by subjecting them under harsh physical or chemical conditions [56, 79, 80].

2.2.3.1 Arc Discharge Method This method has been applied to produce CDs with a QY of 1.6% from crude carbon nanotube soots [7]. After oxidising the crude material (soot) with nitric acid, the oxidised materials were extracted with an alkaline solution (pH 8.4). Then, the extracted products were purified by gel electrophoresis.

2.2.3.2 Laser Ablation A simple laser ablation approach using nano-carbon materials as the starting material in a solvent as the liquid media was introduced by Li and co-workers in 2010 [81]. The typical procedure started with dispersing a small amount of nano-carbon material (0.02 g), in 50 mL of solvent such as ethanol, acetone, or water. The solution was then ultrasonicated before dropping 4 mL of the suspension into a glass cell for laser irradiation. The suspension was irradiated using a neodymium-yttrium-aluminum garnet (Nd:YAG) pulsed laser with a second harmonic wavelength of 532 nm. Subsequently, the solution was centrifuged to obtain the supernatant containing the CDs. Another rendition of CDs produced via laser ablation was carried out by laser irradiating a carbon target in the presence of water vapour with argon as a carrier gas at

900°C and 75 kPa [7, 82]. It was then refluxed in nitric acid (HNO3) for 12 h and surface passivated by attaching simple organic species such as poly(ethylene glycol)

(PEG1500N) and poly(propionylethyleneimine-coethyleneimine) (PPEI-EI) resulting in a production CDs with bright PL emission.

2.2.3.3 Electrochemical Methods CDs can be prepared through a powerful method which is electrochemical soaking of various bulk carbon materials as precursors. However, reports on CDs produced via electrochemically carbonising small molecules are very limited. In a report by Deng and co-workers, alcohols were used as precursors and transited into carbon-containing particles by subjecting them through electrochemical carbonization under basic conditions [83]. The working and auxiliary electrode consisted of two platinum sheets, and the reference electrode was a calomel electrode mounted on a freely adjustable Luggin capillary. When applied with increasing potential, the sizes and graphitisation degrees of the CDs were increased. CDs produced from this method exhibited excellent excitation and size-dependent PL properties without needing complicated purification and passivation steps. Demonstration of the CDs in human cancer cells indicated that the CDs have low toxicity.

2.2.4 Bottom-up Approaches In the bottom-up methods, CDs are prepared from small molecules or precursors such as glucose and fructose which are then carbonised via ultrasonification, microwave pyrolysis, hydrothermal treatment and heating [78, 84-86].

2.2.4.1 Thermal Routes Soot collected from burning candles had been used as starting materials for CDs [87]. It was then treated with oxidants such as HNO3 and hydrogen peroxide/acetic acid

(H2O2/AcOH) to produce CDs. When using polyacrylamide gel electrophoresis to separate the as-prepared CDs, it was observed that the migrating CDs with higher mobility had PL at shorter emission wavelengths. The QY values of these CDs ranged from 0.8% to 1.9%.

2.2.4.2 Microwave-assisted Methods Microwave irradiation is a quick and low-cost method to synthesise CDs. Green luminescent CDs were obtained by microwaving sucrose as the carbon source in diethylene glycol (DEG) as the reaction media for 50 s [88]. These DEG-stabilised CDs disperse well in water with a transparent appearance. The intensity of the emission first increased to its maximum at 360 nm excitation wavelength and then decreased as the excitation wavelength went higher up. In addition, the PL peak showed no perceptible 22 shift over an excitation range from 320 to 380 nm. Optical labelling was successfully performed on C6 glioma cells, indicating the low cytotoxicity of these CDs. In another study carried out by Zhai and co-workers, highly luminescent CDs were produced by one-step microwave-assisted pyrolysis of citric acid in the presence on various amine molecules [89]. The primary amine molecules served dual functions, as N-doping precursors and surface passivation agents for the CDs, which could considerably enhance the PL. The increase in nitrogen (N) content for the CDs fabricated from citric acid and 1,2-ethylenediamine resulted in the increase of QY values for up to 30.2%. Moreover, these CDs showed good biocompatibility.

2.2.4.3 Hydrothermal/Solvothermal Routes Hydrothermal or solvothermal carbonisation has the advantages of being a low-cost, environmentally friendly and nontoxic method of producing CDs from various precursors. A solution of organic precursor is typically sealed and reacted in a hydrothermal reactor/autoclave at high temperature. Many kinds of precursors have been used to prepare CDs using hydrothermal treatment such as citrate [90], ascorbic acid [91], glucose [60], orange juice [92], pomelo peels [93], gelatine [94] and grass [95]. Highly photoluminescent CDs with QY of 26% were synthesised through one- step hydrothermal treatment of orange juice followed by centrifugation [92]. The particle size of obtained CDs was between 1.5-4.5 nm. The CDs also exhibited high photostability and low toxicity making them suitable for bioimaging. In another study carried out by Yang and co-workers, one-step synthesis of amino-functionalised CDs were prepared by hydrothermal treatment of chitosan at 180°C for 12 h [96]. These CDs were used to bioimage human lung cancer cells and showed excellent biocompability and low cytotoxicity. The amino-functionalised surface of CDs meant no further passivation or modification is necessary for their use as bioimaging agents.

Another popular approach to fabricate CDs is the solvothermal carbonisation of percursors followed by extraction with an organic solvent which was performed by Bhunia and co-workers [97]. Carbonaceous compounds are usually heat-treated in organic solvents with high boiling point, followed by extraction and concentration. In their study, hydrophobic and hydrophilic CDs were synthesised from carbonisation of a variety of carbohydrates in different solvents. Hydrophobic CDs were fabricated by mixing different amounts of carbohydrate with octadecylamine and octadecene before

23 being carbonised at temperature up to 70-300°C for 10-30 min. The hydrophilic CDs, on the other hand, were prepared by heating up aqueous solution of carbohydrate within wide pH ranges. In the same study, red and yellow emitting hydrophilic CDs were also prepared by heating together aqueous solution of carbohydrate and concentrated phosphoric acid at 80-90°C for 60 min.

2.2.5 Surface Functionalisation Modifying surface of CDs through functionalisation is a very effective way to tune its surface properties for desired applications. Surface functionalisation of CDs can be achieved through surface chemistry or interactions such as covalent bonding [96], coordination [98], π-π interaction [99] and sol-gel technology [100].

Synthesised CDs are mostly rich with oxygen-containing groups, which make them feasible for covalent bonding. A common approach to improve PL of CDs includes surface passivation via covalent bonding of amine-rich agents. One of the simplest and first applied surface functionalisation methods is treatment with acid solutions. When CDs were refluxed for a few hours with concentrated nitric acid solution, useful surface defects were created and also various oxygen-containing functional groups such as hydroxyl, carboxyl, and carbonyl groups were introduced [101].

EDC chemistry is a functionalisation method that is based on carboxyl-to-amine crosslinking using 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) and N- hydroxysuccinimide (NHS) or sulfo-NHS. Huang et al. (2012), used EDC and NHS to activate the carboxyl groups of chlorine e6, also known as talaporfin which is used in photodynamic therapy as a photosensitizer, in order to react with the amine groups of

CDs passivated with polyethylene glycol-2000 (PEG2000N) [102]. This process allowed the CDs to have covalently grafted chlorine e6 through amide bonds.

Aside from covalent bonding to CDs, another useful strategy is coordination. An off-on fluorescence probe that is capable of phosphate (Pi) detection were developed using europium-adjusted CDs and has been successfully applied to detect Pi in complicated matrixes such as artificial wetlands system [98]. The fluorescence of the CDs was turned off when the surface carboxyl groups on the CDs were coordinated with Eu3+ inducing the CDs to aggregate (Figure 2.10). The fluorescence could be switched on again when Eu3+ was specifically coordinated with Pi. Since the Eu3+ ions display

24 higher affinity with oxygen-donor atoms from Pi that the ones from the surface carboxylate groups on CDs, this process allowed the aggregated CDs to dissociate.

Figure 2.10 Schematic diagram of Pi detection based on the off-on fluorescence probe of CDs adjusted by Eu3+. [98]

Another promising approach to fabricate functional molecules on the surface of CDs is the sol-gel technique. Highly luminescent amorphous organosilane-functionalised CDs (QY=47%) with diameters of ~0.9 nm were synthesised using organo-silane as a coordinating solvent [100]. The CDs that benefited from the surface methoxysilyl groups can be easily fabricated into pure CDs fluorescent films or monoliths by the simple method of heating at 80°C for 24 h. In addition, the hydrophobic CDs can be further modified into water-soluble CDs with good biocompatibility and low toxicity.

2.2.6 Applications

2.2.6.1 Biosensing CDs have high solubility in water, flexible surface modification, low toxicity, good biocompatibility, high photostability, good cell permeability and excitation-dependent multicolour emission which make them good for utilisation as biosensors. Biosensors based on CDs have been developed to visually monitor glucose [103], phosphate [98], iron [73], pH [104], and nucleic acid [99].

Li and co-workers demonstrated that carbon nanoparticles can be effectively applied as a fluorescent sensing platform to detect nucleic acid with selectivity down to a single- base mismatch as shown in Figure 2.11 [99]. Two steps were employed to detect DNA: First, the dye-labelled single-stranded DNA (ssDNA) probe was adsorbed onto the surface of the CDs via π- π interaction, leading to the fluorescence quenching of the dye. In the second step, the dye-ssDNA hybridised with its specific target to form a double- 25 stranded DNA (dsDNA) hybrid which then desorbed itself from the CDs, recovering the dye fluorescence.

Figure 2.11 A schematic illustration of the CD-based fluorescent nucleic acid detection. [99]

A tunable ratiometric pH sensor to measure intracellular pH was developed using CDs labelled with two fluorescent dyes [104]. The preparation of CD-based fluorescent pH started with creating amino-coated CDs by heating citric acid in glycerol at 220°C for 3 h under argon in the presence of a surface coating agent. Then, the amino–coated CDs were treated with various molar ratios of pH-sensitive fluoresceinisothiocyanate (FITC) to pH-insensitive rhodamine B isothiocyanate (RBITC) dyes to produce the dual- labelled CDs. After purification using dialysis and gel chromatography, the modified CDs were incubated with HeLa cells in different pH environments. Standard MTT assay test indicated low cytotoxicity and good biocompatibility of the CDs. Fluorescence imaging analysis of the HeLa cells were used to visualise the CDs colour change with pH (Figure 2.12).

Figure 2.12 Fluorescent images of HeLa cells clamped at pH 6.0, 6.5, 6.8, 7.2, 7.5, and 8.0, respectively. The images of the first row (FITC channel) and second row (RBITC channel) were collected in the ranges of 510–550 nm and 570–610 nm, respectively. The third row shows the corresponding differential interference contrast images. Olympus software (FV10-ASW) was used to generate the images of the fourth row (the ratio channel). The bottom colour strip represents the pseudocolour change with pH. Scale bar, 20 mm. [104]

2.2.6.2 Chemical Sensing Monitoring changes in the fluorescence of CDs under external chemical or physical stimuli have been used to detect substances and quantities like temperature, pH [105], Fe3+, nitrite [106], Ag+ [107], and heavy metals such as Hg2+ [107] . These CDs can be applied to monitor conditions especially pollution of the environment, water systems, and food.

Copper(II) ion (Cu2+) detecting CDs were developed by Vedamalai and co-workers [108]. The CDs were synthesised from o-phenylenediamine (OPD) using a simple hydrothermal method and had a peak emission at 567 nm when excited at 420 nm. The colour of the CDs changed from yellow to orange with an increased PL intensity when

Cu2+ ions were present due to the formation Cu(OPD)2 complexes on the surfaces of CDs. The CDs showed a linear detection of Cu2+ ions over a concentration range of 2- 80 nM and the limit of detection for Cu2+ ions was 1.8 nM with a signal-to-noise ratio of

S/N= 3. In addition, the CDs showed good biocompatibility and has been employed for the localisation of Cu2+ ions in cancer cells treated with Cu2+ ions.

A peroxynitrous acid-carbon dots system was developed for the detection of nitrite [106]. The changes in chemiluminescent property of the CDs in the presence of peroxynitrous acid allowed for the sensing of nitrite. Peroxynitrous acid was formed by mixing nitrite and acidified hydrogen peroxide. The CDs exhibited a linear increase in CL intensity with nitrite concentration in the range from 1.0× 10-7 M to 1.0×10-5 M, and the detection limit was 5.3×10-8 M with a signal-to-noise ratio of S/N= 3.

2.2.6.3 Imaging The emergence of fluorescent CDs drew the attention on their application as a viable alternative in the field of bioimaging for their biocompatibility, low toxicity, high PL, chemical stability, and photo stability.

One of the first applications of CDs for bioimaging was carried out by Yang and co- workers in 2009 where in vitro and in vivo imaging was demonstrated [109]. In 2006, Sun et al. pioneered biolabeling using CDs by incubating E. coli ATCC 25922 with PEGylated CDs and the multicolour fluorescence were imaged with a confocal microscope under different excitation lengths [59]. CDs have also performed the same or even better for in vivo imaging in mice models when compared to other commercial fluorophores such as QDs. The first ever use of CDs as fluorescence imaging agents for in vivo imaging of mice was reported by Yang et al., (2009) [6]. The study involved injecting aqueous PEGylated CDs solution subcutaneously into mice and observing the fluorescence under different excitation wavelengths. The resulting images indicated sufficient contrasts were present for both red and green emissions. In addition, CDs were also shown to be biocompatible and performed competitively against commercially supplied CdSe/ZnS QDs. In another relevant study, high doses of PEGylated CDs were administered to mice and histopathological analyses of the liver spleen and kidneys were carried out [58]. After 6 h of intravenous exposure to CDs, the liver and spleen of the mice were harvested. Histopathological analyses indicated that CDs were present in the liver and spleen at relatively higher amounts (Figure 2.13). Furthermore, mice subjected to extended period of time (28 days) showed no signs of toxicity.

Figure 2.13 Top: Images from histopathological analyses of liver, spleen, and kidneys. Bottom: Fluorescence images (two-photon excitation at 800 nm) of sliced liver and spleen harvested from mice 6 h after intravenous exposure to C-dots. [58]

Numerous studies to monitor cellular uptake of various CDs using fluorescence microscopy were also demonstrated. In a study carried out by Qiao et al. (2010), CDs were surface passivated using 4,7,10-trioxa-1,13-tridecanediamine (TTDDA) and tested for their photostability and cell permeability on African green monkey kidney fibroblast-like (COS-7) cells [80]. After the COS-7 cells were incubated with the passivated CDs for 24 h, the COS-7 cells became bright, indicating that the CDs had permeabilised into the cells. Confocal fluorescence microphotographs of the treated cells showed that the CDs were accumulated in both the cytoplasm and cell membrane 29 of the COS-7 cells. When a continuous excitation of 10 min was exerted on the cells, no obvious PL intensity reduction was observed which indicated that the CDs were highly photostable with low photobleaching. Similarly, Yang and co-workers (2012) synthesised amino-functionalised CDs by hydrothermal treatment of chitosan [96]. After imaging of A549 cells incubated with these CDs, fluorescence spots were only observed in the cell membrane and cytoplasmic region with very weak signal in the cell nucleus area. This observation suggested that the CDs entered the cell easily but did not enter the nuclei.

Many studies on the imaging of CDs incubated cells suggested that internalised CDs are generally localised in the cytoplasm [59, 110, 111]. However, there are reports that demonstrated the localisation of CDs in the cell nucleus. For example, fluorescent carbon nanoparticles were synthesised from nitric acid oxidation of carbon soot without any additional surface functionalisation and incubated with Ehrlich ascites carcinoma (EAC) cells [112]. The incubation was done for only 30 min and subsequent imaging using a conventional microscope showed the CDs were internalised into both of the cytoplasm and nucleus of the cell. The EAC cells exhibited bright blue-green light under UV excitation and yellow light under blue excitation. The whole cells were lighted brightly under UV excitation while under the blue excitation the yellow light in the cytoplasm was more defined than in the nucleus. The mechanisms of various CDs internalising in different compartments in cells is still not clear and more high- resolution imaging studies are needed to comprehend the intracellular distribution of CDs.

2.2.6.4 Catalysis Another interesting use of CDs is in photocatalysis and electrocatalysis. In a drive for greener alternatives in organic synthesis, advanced materials for electrocatalytic and photoelectrochemical (PEC) hydrogen evolution (HER) reaction are sought after. The rising interest in photocatalysis is partly motivated by the fact that sunlight is an effective renewable source. However, organic compounds may be adversely damaged by high UV energy and short wavelength visible light [113]. Solution species of CDs demonstrated the capability of harnessing long wavelength light and energy exchange making them potentially usable as photocatalysts in organic synthesis. A study by Li et al., (2013) has demonstrated that small CDs (1-4 nm) are effective NIR light driven

30 photocatalysts for the effective oxidisation of benzyl alcohol to benzaldehyde with high conversion (92%) and selectivity (100%) in the presence of antioxidant hydrogen peroxide (H2O2) [114]. Control catalytic experiments confirmed that excellent catalytic activity is due to the effective photocatalytic activity for H2O2 decomposition and NIR light induced electron transfer property.

CDs have also shown to improve photocatalytic performance of existing photocatalysts in the form of nanocomposites. A popular photocatalyst, titanium dioxide (TiO2) has been used in organic pollutants removal and water splitting to generate hydrogen gas

(H2) [115]. Despite being a promising photocatalyst, TiO2 has an ineffective utilisation of visible light as the irradiation source which affects its photocatalytic efficiency. This is due to the bandgap of bulk TiO2 (3.0-3.2 eV) that lies in the UV region, indicating that only less than 5% of sunlight is utilised by TiO2. Consequently, one of the approaches to enhance TiO2 photocatalytic performance is through bandgap engineering by possible modification of TiO2-based materials. Taking into account of the excellent upconversion luminescence properties of CDs, a nanocomposite complex of CDs and

TiO2 was designed to harness the full spectrum of sunlight as illustrated in Figure 2.14

[75]. Demonstrating on methylene blue (MB) as a model compound, the TiO2–CD nanocomposites fabricated by Li et al., (2010) were able to completely degrade MB (50 -1 mg mL ) within 25 min under visible light irradiation compared to when pure TiO2 is used as a the photocatalyst where only less than 5% of MB was degraded.

Figure 2.14 Possible catalytic mechanism of the TiO2–CD nanocomposites under visible light. [75]

One of the important electrocatalytic reactions in renewable energy technologies is the oxygen reduction reaction (ORR) which is applied in fuel cells and water splitting [116]. Nitrogen-doped CDs aggregates synthesised from the hydrothermal treatment of willow leaves performs excellent electrocatalytic activity for the ORR via a dominant four- electron oxygen reduction pathway in 0.1 M potassium hydroxide (KOH) aqueous solution. In the addition, the CDs showed great stability (even after 20 000 cycles) as well as a methanol and carbon monoxide (CO) tolerance that is superior to a commercial platinum on carbon (Pt/C) catalyst. Not only do these heteroatom-doped CDs exhibit highly efficient electrochemical activity, they also showed excellent stability and immunity towards methanol and CO in practical applications, which are common problems for metal-based catalysts.

2.3 Fluorescence In Situ Hybridisation

Fluorescence in situ hybridisation (FISH) is a branch of bioimaging method that is widely used in molecular cytogenetics [117, 118]. It is a powerful technique that utilises fluorescent probes which can bind selectively to complementary sequence of a DNA. In addition, FISH can also be used to detect and localise RNA targets such as mRNA, lncRNA and miRNA in cells and tissue samples. Molecular cytogenetics is the study of chromosome biology and the application of molecular cytogenetic techniques varies from clinical diagnostics to basic genome research [119].

FISH technology has been developing continuously hitherto since its first appearance in the 1980s [117]. During its early years, FISH was developed as an alternative to radiolabelled probes used in older methods for visualisation of nucleic acids [120]. The older methods of isotopic detection used non-specific labelling strategies like the random incorporation of radioactive modified bases into growing cells which is then accompanied with autoradiography. There are several drawbacks of isotopic hybridisation that paved the way for new and better techniques. Firstly, the isotope decays over time due to the radioactive nature of the material in the probe, causing instability and inconsistency in the specific activity of the probe. Secondly, the resolution of radiography is limited despite its sensitivity. Thirdly, to produce measurable signals on radiography film, long exposure is needed which delays the results of assay. Lastly, radiolabelled probes are quite costly and hazardous. The transportation, storage, disposal and handling of this material have to be carried out in 32 accordance with regulations. With FISH techniques, the resolution, speed and safety of nucleic acid detection have vastly improved and have allowed for more complex detection methods such as multiplexed bioimaging [121], quantitative analyses [122], and live-cell imaging [123, 124].

Fluorescent in situ detection was first realised by Bauman et al., (1980) when they directly labelled a fluorophore on the 3’ end of an RNA and used it as a probe for specific DNA sequences [125]. The development of amino-allyl modified bases [126], which allowed for conjugation to any kind of hapten or fluorophore, was central to the advancement of in situ technologies as it led to the production of various low-noise probes by simple chemistry. Assessment of nucleic acids with low copy number using FISH is prevented by low probe specific activity. Ergo, indirect detection methods were established to allow artificial increase in signal output by the use of secondary reporters that bind to the hybridisation probes. During the early 1980s, detection of DNA [127] and mRNA [128] targets were achieved using assays that featured nick-translation, biotinylated probes, and secondary detection by fluorescent streptavidin conjugates. Around a decade later, chemical preparation of hybridisation probes carrying enough fluorescent molecules to allow direct detection [129] became feasible and is attributed to the improvements on the labelling of synthetic, single-stranded DNA probes. Since then, many variations on the methods for direct and indirect labelling have been introduced which allow for a wide selection of detection schemes to choose from.

2.3.1 Mechanism of FISH In 1953, Watson and Crick proposed the double helix structure of DNA which is two strands of bases connected anti-parallel to each other by hydrogen bonds [130]. The bases paired in these strands are complementary; where A bases are always paired with T bases, and G bases are always paired with C bases (Figure 2.15). The many hydrogen bonds that hold the bases together make the double helix a very stable structure. A DNA denatures to break into two single strands when heat or chemicals disrupt the hydrogen bonds that hold the double helix together. Under more favourable conditions, the helix can re-form or renature. The ability of the DNA helix to renature becomes the basis for molecular hybridisation.

Figure 2.15 Base pairing in DNA. T and A are connected by two hydrogen bonds whereas G and C are connected by three hydrogen bonds. The sugar-phosphate backbones (grey) run anti-parallel to each other, so that the 3’ and 5’ ends of the two strands are aligned. [131]

FISH relies on exposing the chromosome to a small DNA sequence called a probe that has a fluorescent molecule attached to it. Under optimised conditions, the probe will only anneal to its targeted complementary sequences. Hybridisation occurs when the probe meets the target as shown in Figure 2.16. In the figure, the probe sequence, usually a short strand of cloned DNA, is represented in red. The chromosomes on the glass slide are the target DNA, shown in blue (right column). The hydrogen bonds that connect the double-stranded DNA helix are represented by black lines.

In general, the first step of FISH is to make a fluorescent copy of the probe sequence (Figure 2.16b, middle column) or a modified copy of the probe sequence that can later be labelled with a fluorescent material (Figure 2.16b, left column). In the next step, both the target and the probe sequence have to be denatured with heat or chemicals (Figure 2.16c) before any hybridisation steps can take place. The denaturation step is crucial to allow for new hydrogen bonds to form between the target and the probe during hybridisation. In Figure 2.16d, the probe and target sequence are then mixed together to allow the probe to specifically hybridise with its complementary sequence on the 35 chromosome. The probe that is already fluorescently labelled (middle column) may be used to detect the site of hybridisation directly. On the other hand, the probe sequence that has not been fluorescently labelled (left column) requires an additional step to visualise the hybridised probe. Once hybrids are formed between the probes and their target, a fluorescent microscope can be used for detection. The process of FISH is quicker when using directly labelled probes whereas the use of indirectly labelled probes provides an advantage of signal amplification by using multiple layers of antibodies. It may therefore offer a brighter signal compared with background levels.

2.3.2 FISH Applications

2.3.2.1 Gene Positioning FISH provides the means to identify the location of a cloned DNA sequence on metaphase chromosomes. In this example of a typical FISH experiment (Figure 2.17), a cloned DNA was hybridised to normal metaphase chromosomes [133]. The detection at hybridisation sites on two homologous chromosomes are localised by the characteristic banding patterns in red. On closer examination, each red band is made up of two spots that corresponds to the two sister chromatids in a mitotic chromosome.

Figure 2.17 (a) Fluorescent signals are seen at cytogenetic bands (grey) where fragments of a sequence- tagged BAC hybridise (red) via FISH. (b) The breakpoint of a translocation involving chromosomes 11 and 19 in a patient with multiple congenital malformations and mental retardation (DGAP012) were mapped out using clones selected based on band location. Clone CTD-3193o13 spans the breakpoint on chromosome 19 where the red signal is split between the derivative chromosome 11 and derivative 19 chromosomes. The red signal is also present on the normal chromosome 19. The GTG-banded karyotype for this patient is 46,XY,t(11;19)(p11.2;p13.3). [133]

During the course of the Human Genome Project (HGP), an international project that began in 1990 and concluded in 2003, three billion nucleotide base pairs were sequenced and a complete gene mapping of human chromosomes were realised [134]. The results from FISH and other in situ hybridisation experiments were then collected and compiled in databases that are useful for the HGP during its annotation phase. After the completion of the HGP, FISH and related in situ hybridisation are rarely used for identifying chromosomal location of a human gene anymore. However, these techniques still provide useful gene mapping data for genomes of other species that have not been sequenced yet.

2.3.2.2 Multicolour-FISH Detecting chromosome rearrangements using site-specific probes (see Figure 2.17b) can be a very lengthy process, especially if the rearrangements are complex or the rearranged regions are difficult to identify by their banding patterns. In order to overcome this, multifluor FISH, or spectral karyotyping, were developed for rapid scanning of a set of metaphase chromosomes for potential rearrangements [135]. Using multifluor FISH, a karyotype in which each chromosomes are labelled with different colours can be generated, which is why it is also alternately termed as ‘multicolour FISH’. Each ‘colour’ consists of a collection of hybridisation probes for sequences that span the length of a specific chromosome.

Multifluor FISH requires the researchers to prepare a collection of DNA sequences to be used as probes for each chromosome beforehand [136]. As seen in Figure 2.18a, the probe chromosomes were sorted out by flow cytometry. Nowadays, it is easier for researchers to just obtain commercially available DNA probes collections for each chromosome. Next, fluorochromes are attached to the DNA samples to produce a unique colour for each chromosome. The fluorescent probes are subsequently mixed with and hybridised to metaphase chromosomes. An example of what a microscope image of interphase or metaphase chromosomes would look like after hybridisation is shown in Figure 2.18b. Several of the metaphase chromosomes may appear the same through human eyes but with the help of digital image processing, the spectral differences between the chromosomes can be determined. A uniform colour will form along the lengths of a normal human chromosome while a rearranged chromosome will show a striped appearance.

Figure 2.18 Spectral karyotyping and multicolour-FISH paint each human chromosome in one of 24 colours. Cytogenetic localization of DNA sequences via FISH. [136]

2.3.2.3 Diagnosing Chromosomal Abnormalities FISH is a crucial tool in clinical diagnosis of several chromosomal abnormalities such as deletions, duplications, and translocations, which are referred to as ‘chromosomal breakpoints’. Figure 2.17b shows an example of FISH used together with standard karyotyping to analyse translocation in a patient. The segment of chromosome 19 where the probe was hybridised to was suspected to contain the translocation breakpoint. In the fluorescent image, three areas of hybridisation can be seen: a spot that corresponds to the patient’s normal copy of chromosome 19 (nl19) and another two spots that corresponds to the altered versions of chromosomes 11 and 19 that were made during the translocation. Using this information, the breakpoint region on chromosome 19 could be narrowed down and the second chromosome involved in the translocation were identified.

Trisomy 21 or Down syndrome, is a genetic disorder caused by the presence on a third copy of chromosome 21. Diagnosis of Down syndrome in prenatal stage were carried out by interphase FISH of amniocytes collected from pregnant women [137]. Interphase cells obtained from the uncultured amniotic fluid samples were hybridised to Ls121 DNA probe that corresponds to the D21S259, D21S341, and D21S342 loci located in the 21q22.13 to 21q22.2 region on chromosome 21. The results of the diagnosis were confirmed with a karyogram (Figure 2.19).

Figure 2.19 Interphase FISH for uncultured amniocytes (right) using a fluorescent probe for chromosome 21 (orange), and chromosome 13 (green) showing 3 signals of chromosome 21 giving the diagnosis of trisomy 21 or Down syndrome. The karyogram (left) done on cultured amniotic fluid cells confirms the FISH diagnosis. [137]

2.3.2.4 Diagnosing Diseases Another important role that FISH has played is in the detection of diseases that are caused genetically or by infection. Moreover, FISH probes can be employed on many forms of cellular targets such as nuclei derived from cell suspension, formalin-fixed paraffin-embedded tissue sections, fixed cell lines, interphase chromosomes, and metaphase chromosomes.

Although there are many available methods of detecting cancer, FISH provides better analysis by allowing simultaneous detection and localisation of the cancerous mass or genes. In 2010, Chen and co-workers developed a FISH technique using red fluorescent quantum dot labelled oligonucleotide probe (QD-FISH) for the detection of Epstein- Barr virus (EBV) in paraffin-embedded nasopharyngeal carcinoma tissue blocks (Figure 2.20) [138]. Nasopharyngeal carcinoma (NPC) is closely associated with EBV infection due to the presence of EBV DNA found in majority of NPC cells. Once inside the cell, Epstein-Barr virus encoded small RNA (EBER) is replicated abundantly by the virus; hence, serves as a marker for EBV in cells. Thus, FISH can be used for the detection 39 and localisation of EBV in infected tissue sections by using fluorescently labelled EBER oligonucleotide probe.

Figure 2.20 Detection of EBER by QD-FISH. NPC tissue sections were subjected to QD-FISH and observed under blue light excitation in a fluorescence microscope. (A–C) NPC specimens showed positive signals for EBER. (D) In the hybridization mixture containing EBER oligonucleotides without biotin, negative signals were detected. Original magnifications: A, D: 200×; B, C: 400×. [138]

FISH assays test kit were developed to detect malaria infection caused by parasitic Plasmodium knowlesi in blood samples [139]. The kit uses a P.knowlesi-specific probe labelled with Alexa 488 green fluorescent dye and also a Plasmodium genus-specific probe with Atto 550 orange fluorescent dye. After hybridisation using the PK-FISH assay, a dual fluorescence microscope was used to observe the probe. Under LED illumination (Figure 2.21), P.knowlesi parasites would appear green under a green filter (excitation 492 nm; emission 530 nm band pass) while all Plasmodium species, including P.knowlesi would appear orange under an orange filter (excitation 560; emission 630 nm long pass).

Figure 2.21 Fluorescence observed in the PK-FISH assay with the P. knowlesi-specific probe (green) and the Plasmodium genus-specific probe (orange) on monkey blood smears containing P. knowlesi. Dual colour fluorescence seen in a single microscope field at ×1000 magnification using the two different filters is shown in each set of paired photographs. The four sets of paired photographs are from four different fields (i) - (iv). Scale-bars: 5 μm. [139]

3 Synthesis and Characterisation of Carbon Dots

3.1 Materials and Instruments

3.1.1 Materials Carboxymethylcellulose (CMC) sodium salt, medium viscosity was bought from MP Biomedicals, Inc. (CAT NO. 150560). Ortho-phosphoric acid, 85%, was purchased from R&M Chemicals/1502-80. All the metals used for metal test; copper(II) nitrate

(Cu(NO3)2), aluminium nitrate (Al(NO3)3), cadmium chloride (CdCl2), lead(II) nitrate

(Pb(NO3)2), magnesium nitrate (Mg(NO3)2), iron(III) nitrate (Fe(NO3)3), chromium(II) nitrate (Cr(NO3)2), zinc nitrate (Zn(NO3)2), sodium arsenite (NaAsO2), cobalt(II) nitrate

(Co(NO3)2), nickel(II) nitrate (Ni(NO3)2), mercury(II) chloride (HgCl2), and tin(II) chloride (SnCl2) powder were purchased from R&M Chemicals.

3.1.2 Instruments The instruments used in this study are the Varian Cary Eclipse Fluorescence Spectrophotometer, JEOL JSM-6390LA Analytical Scanning Electron Microscope (SEM) by UNIMAS and Perkin Elmer Thermoscientific Smart Omni Transmission Nicolet IS10 Fourier Transform Infrared Spectrometer (FTIR) by UNIMAS.

3.2 Hydrothermal CD Synthesis

For this research project, CDs were prepared through a bottom-up method in which a carbon precursor was carbonised via hydrothermal treatment. The starting precursor material that had been chosen for the CDs synthesis was carboxylmethylcellulose (CMC), a cellulose derivative commonly used in food products as thickeners and stabilisers. It is also used as a treatment for irritation caused by dry eyes. CMC has a backbone that consists of glucopyranose monomers with their hydroxyl groups attached with carboxymethyl groups (Figure 3.1). Since it is non-toxic and contains carboxyl groups in its structure, CMC is a suitable starting material for synthesis of the CDs in this project.

Figure 3.1 Chemical structure of sodium carboxymethylcellulose.

For bioimaging applications, it is preferable to use CDs that do not emit in the blue region. This is mainly to prevent interference with the biological samples that can sometimes autofluoresce in the blue region. Most of the early reported synthesised CDs are in the blue-emitting region and this has gained some interests among researchers to explore CDs with red-shifted emission [140]. In a previous work reported by our group, we have successfully red-shifted blue emitting CDs by using phosphoric acid during the synthesis, which resulted in the production of orange emitting CDs [141]. The addition of acid was also found to help in reducing the time and temperature needed for the hydrothermal treatment. Hence, CMC was hydrothermally treated with phosphoric acid in this work.

The starting material used was a 25.0 mg/mL stock CMC solution prepared by dissolving CMC powder in distilled water and phosphoric acid (85%). Hydrothermal treatment of starting materials were carried out in Teflon lined hydrothermal autoclave reactors. The parameters used for hydrothermal treatment of CDs were optimised based on the most favourable emission yield and wavelength of the raw CDs produced.

In each Teflon lined hydrothermal autoclave reactor, 5.0 mL CMC solution was mixed with 5.0 mL phosphoric acid to make up a total volume of 10.0 mL. The mixture was heated in a furnace oven. The general formulation for the synthesis of CDs was repeated with varying CMC concentration, phosphoric acid concentration, temperature and heating time for optimisation study.

3.3 Optical Measurement

The synthesised CDs were filtered against 110 mm filter paper, 0.45 µm, and 0.22 µm syringe filters to remove bigger char and aggregates. The resultant solution obtained is referred to as ‘raw CD’. The emission profile of the CDs was measured using a spectrophotometer with 10 nm for emission and excitation slit size unless stated otherwise. The CD samples were prepared by diluting 100 µL of the raw CD with 3.0 mL of distilled water.

Neutralisation and isolation of the raw CD were carried out to make it suitable for further reaction like EDC-NHS conjugation and FISH. These steps were also done to remove contaminants and salt from the CDs for subsequent FTIR and SEM analysis.

The raw CDs were neutralised with 10 M NaOH to reach a pH 7 in a 50.0 mL falcon tube. Salt and other impurities produced during neutralisation were removed by salting out extraction method. This was done by reducing solubility of molecules in a solution with high ionic strength. With the aid of an organic solvent as the extractant, the more hydrophobic CDs can be separated from the aqueous mixture [142]. Approximately 1.0 mL of acetone solution was added to 1.0 mL of the neutralised CDs solution to isolate the CDs. The mixture was shaken well until two layers could be seen and the salt precipitates would start to form. The solution was left to settle for 15-20 min under the fume hood with occasional shaking in between until the salting out process was completed (Figure 3.2a). The upper layer in the mixture was comprised of acetone mixed with CDs solution. The bottom layer contained the salt precipitates. The upper layer was then filtered through a filter paper and collected into a new clean falcon tube. At this point, the emission profile of the filtered CDs was measured before drying. Subsequently, the tube was transferred into an oven and dried overnight at 80°C to remove the excess acetone and water. The CDs were dried and formed into a dark brown solid layer that could be scrapped off and collected as shown in Figure 3.2d.

Figure 3.2 Salt removal from CDs solution using salting out extraction. (a) Two layers formed after mixing acetone with the neutralised CD solution. The top layer consists of acetone mixed with CDs while the bottom layer is salt. (b) Salt precipitates filtered out from the CDs solution. (c) Isolated CDs layer mixed in water and acetone. (d) Solid CDs after drying out in oven at 80°C overnight.

The isolated solid CDs were further characterised by using SEM and FTIR analyses. For SEM, CDs samples were analysed using the JEOL JSM-6390LA Analytical Scanning Electron Microscope (UNIMAS). The dried neutralised and isolated solid CDs were mounted on aluminium stubs using a double-sided tape. Then, the samples were coated with a thin film of gold using JEOL JFC-1600 Auto Fine Coater before SEM analysis.

3.4 Optimisation of Parameters for CDs Synthesis

3.4.1 Temperature Temperature was one of the first parameters to be optimised for the synthesis of CDs through hydrothermal method. In each Teflon chamber, 5.0 mL of 25.0 mg/mL CMC and 5.0 mL of concentrated phosphoric acid, 85 wt. % were mixed and heated in the furnace for 2 h. The temperatures used for the set-up were 100, 150, 200, 250, and 300°C.

3.4.2 Duration of Synthesis The following parameter to be optimised was the heating duration of the hydrothermal chambers at the optimised temperature of 200°C. In each Teflon chamber, 5.0 mL of 25.0 mg/mL CMC and 5.0 mL of concentrated phosphoric acid, 85 wt. %, were mixed and heated in the furnace for 1.0, 1.5, 2.0, 2.5, and 3.0 h.

3.4.3 Effect of Acid Ratio In each Teflon chamber, 5.0 mL of phosphoric acid with different concentrations of 17, 34, 51, 68, and 85 wt. % were used. Then, the acid was mixed with 5.0 mL of CMC solution and heated in the furnace at 200° for 2 h. The different concentrations of phosphoric acid were prepared by mixing distilled water with stock phosphoric acid (85 wt. %) as detailed in Table 3.1.

Table 3.1 Preparation of different phosphoric acid concentrations.

Final phosphoric acid Volume of stock Volume of Phosphoric acid concentration in total phosphoric acid distilled water concentration (wt. volume of 10.0 mL (wt. (mL) (mL) %) %) 1.0 4.0 17 8.5 2.0 3.0 34 17.0 3.0 2.0 51 25.5 4.0 1.0 68 34.0 5.0 0.0 85 42.5

3.4.4 CMC Concentration The CMC used in this study comes in the form of sodium salt and it is highly viscous when dissolved in water. CMC salt is soluble up to 50.0 mg/mL with the help of heat but it is normally used at concentrations of less than 20.0 mg/mL due to the high viscosity nature. Thus, the highest concentration prepared for this study was 25.0 mg/mL. A stock solution of 25.0 mg/mL CMC solution was prepared by slowly adding 1 g of CMC sodium salt powder into 40.0 mL of warm distilled water while stirring using a spatula. This thick and clear solution was then mixed with distilled water to a total volume of 5.0 mL to prepare different concentrations of 2.5, 5.0, 7.5, 10.0, and 12.5 mg/mL as described in Table 3.2. In each Teflon chamber, 5.0 mL of the prepared CMC solution was mixed with 5.0 mL of 85 % phosphoric acid and then heated at 200°C for 2 h.

Table 3.2 Preparation of different CMC concentrations.

Volume of stock Volume of Final CMC CMC concentration CMC, 25.0 mg/mL distilled water concentration used for (mg/mL) (mL) (mL) the synthesis (mg/mL) 1.0 4.0 5.0 2.5 2.0 3.0 10.0 5.0 3.0 2.0 15.0 7.5 4.0 1.0 20.0 10.0 5.0 0.0 25.0 12.5

3.5 External Effects

CDs are often known to show response to pH changes and in the presence of metal ions. As part of the characterisation of the obtained CDs, the effect of different pH conditions and various metal ions on the fluorescence emission were studied.

3.5.1 Effect of Metal Ions The metal ions involved in the CDs samples testing were Cu2+, Al3+, Cd2+, Pb2+, Mg2+, Fe3+, Cr2+, Zn2+, As3+, Co2+, Ni2+, Hg2+, and Sn2+. All the metal solutions except for

SnCl2 were prepared by dissolving the metal salt powder in clean, distilled water to

47 make 0.1 M of each metal solution. SnCl2 has low solubility in water, so pure ethanol was used as solvent to prepare a 0.1 M solution of SnCl2.

The effect of metals test was performed to investigate the effect of their presence on the fluorescence intensity of CDs. 50 µL of each metal solution was added and mixed into the CDs sample. The emission of each CDs sample was measured at 450 and 595 nm. Only metal ions that induced noticeable response in the CDs sample were used for further metal test. This was done by comparing the change in intensity of CDs between the metal ions added. Then, for each of these chosen metal test, 20 µL of 0.1 M metal solution was mixed into the CDs sample and measured for changes in emission. The addition of 20 µL of the metal solution was repeated until a total of 100 µL had been added to the mix.

3.5.2 pH CDs samples in pH ranging from 1-14 were prepared by mixing 200 µL of raw CDs with 1 M NaOH solution and topped up to a total volume of 6.0 mL with distilled water as shown in Table 3.3. The pH of the samples were measured using the Mettler Toledo pH meter. From each sample, 3.0 mL of the CDs solution was collected to be measured using the spectrofluorometer.

Table 3.3 Preparation of CDs in different pH conditions.

CDs solution 1 M NaOH Distilled Total volume pH (mL) (mL) water (mL) (mL) 0.2 0.0 6.0 6.2 1.0 0.2 1.0 5.0 6.2 2.7 0.2 1.2 4.8 6.2 5.6 0.2 2.0 4.0 6.2 7.1 0.2 2.2 3.8 6.2 8.1 0.2 2.5 3.5 6.2 10.9 0.2 4.0 2.0 6.2 13.6

3.6 Results and Discussion

3.6.1 Optical Properties

3.6.1.1 Neutralisation and Isolation of CDs Figure 3.3 shows the emission profile of CDs in the acetone solution layer after salt and other contaminants had been filtered out. It appeared that neutralising the CDs to pH 7 had shifted the orange emission peak from 595 nm wavelength to 570 nm. This observation showed the CDs solution emitting yellow-coloured fluorescence when exposed under UV light.

Figure 3.3 Emission profile of neutralised CDs (pH 7) in acetone solution after salt precipitates were filtered out.

After drying the CDs solution, solid CDs were weighed and reconstituted in distilled water to make 30.0 mg/mL CDs solution and the emission profile was measured. The pH of this reconstituted CDs solution was 7. From Figure 3.4, it is clearly shown that the emission profile of CDs had shifted to a shorter emission wavelength with the highest emission peak now appearing 470 nm. As the excitation wavelengths was increased above 360 nm the emission peak shifted towards longer emission wavelengths with decreased intensity. Observation under UV light revealed that the CDs were emitting in green colour. The further heating of the CDs overnight for drying and removal of acetone is likely the cause of the altered fluorescence properties of CDs.

Figure 3.4 Emission profile of CDs reconstituted in water (30.0 mg/mL) after neutralisation, isolation and drying process.

3.6.1.2 SEM Analysis Neutralised and isolated solid CDs were analysed to determine particle size. Figure 3.5 and 3.6 depict the SEM images of CDs samples that were prepared on two different days using the same synthesis parameters, the only difference being that an extra filtration step was done on the second CDs sample in Figure 3.6. SEM analyses were performed using Smile View™ image analysis software. Due to the limitation of the SEM magnification and image quality, it was difficult to identify and measure particles smaller than 100 nm. As all the CDs samples were filtered through a 0.22 µm syringe filter immediately after they were synthesised, the CDs were expected to be near 220 nm or smaller in diameter. In Figure 3.5, however, the particle sizes of the CDs are in the range of 30 to 1230 nm. The average particle size of the CDs is 194 nm with a standard deviation of 187 nm. The vast difference in sizes of the CDs may have been due to the aggregation of CDs and the larger sized clumps that were observed in the SEM image may possibly also be CDs in their aggregate form. In the second CDs sample prepared, an extra filtration step using a 0.22 µm syringe filter was carried out after the neutralisation of CDs solution as an attempt to minimise aggregated CDs. As seen in Figure 3.6, the particle sizes of the CDs now vary in a smaller range of 40 to 340 nm. The average particle size of the CDs is 134 nm with a standard deviation of 55 nm.

Figure 3.5 (a) SEM image and (b) size distribution of CDs after neutralisation and isolation. Mean particle size= 0.194 µm. Standard deviation= 0.187 µm. The size distribution represents 95% of measured particles.

Figure 3.6 (a) SEM image and (b) size distribution graph of CDs after neutralisation and isolation prepared on a different day using the same parameters but with an extra filtration step using 0.22 μm syringe filter after neutralisation. Mean particle size= 0.134 µm. Standard deviation= 0.055 µm. The size distribution represents 100% of measured particles.

3.6.1.3 FTIR Analysis FTIR spectroscopy was carried out to investigate the functional groups of CDs. It is worth noting that the CDs were required to be conjugated onto a modified probe via carbodiimide crosslinking later on in this study. This chemistry requires the presence of carboxyl groups on the CDs. Therefore, the FTIR would be used not only for characterisation of the CDs but also for confirming the presence of the carboxylic acid group. Solid samples of the neutralised and isolated CDs were analysed.

Carboxylic acids usually exists as hydrogen-bonded dimers, which gives the broad appearance of the O-H stretch band in the region of 3300-2500 cm-1 centred at about 3000 cm-1 which can be observed in Figure 3.7 [143]. The region of this O-H stretch

51 overlaps with the C-H stretching bands of both alkyl and aromatic groups, thus giving carboxylic acid the jagged absorption pattern in the region 3300-2500 cm-1 which is also observed in the FTIR spectrum of the CDs. Another characteristic of this dimer that can be seen in this spectrum is the carbonyl stretch (C=O) of a carboxylic acid which appears as a strong band at 1760-1690 cm-1. Additionally, O-H bend peak from carboxylic acid is also observed at around 1440-1395 cm-1. This observation confirms the appearance of carboxylic acid functional group in CDs.

Figure 3.7 FTIR spectra of the neutralised and isolated CDs.

3.6.2 Optimisations

3.6.2.1 Temperature After the synthesis of CDs at different temperatures, the CDs solutions were filtered to remove any char and large aggregates that had formed. The filtered CDs solution were observed under normal light and UV light, revealing different colours (Figure 3.8). The starting solution, consisting of CMC mixed with phosphoric acid, that was treated at 100°C had no char or aggregates after the hydrothermal process and remained similar to its initial colour indicating that the carbonisation process had yet to start. Hence, no CDs were formed yet at this temperature. Above 100°C, at temperatures of 150, 200, 250 and 300°C, the process of carbonisation started to happen. This process had caused the increase of char and aggregates in the CDs solutions as the temperature was increased. After filtration, clear solutions were obtained and the colour the raw CDs solutions were revealed as dark brown, reddish-brown, orange and pale yellow (Figure 3.8a). The change of the solution from colourless to coloured and the production of char indicated that “polymerisation” and “carbonisation” steps were in progress, which were similar to what was reported in other studies [73, 144-146]. These two processes are crucial for the formation of CDs. Polymerisation refers to the process of small molecules or monomers combining to form a larger chain of molecules. The dark brown, reddish-brown, orange and pale yellow coloration of the solution indicated the formation of aromatic compounds and oligosaccharides during polymerisation [146]. As the solution approached critical supersaturation during hydrothermal treatment, a short single burst of nucleation occurred, resulting in the formation of nucleated carbon particles. The nuclei subsequently grew in a uniform and isotropic manner by diffusion of solutes in the solution towards the particle surface until a final size was reached. This process is known as carbonisation [147]. Further observation made under a UV lamp showed that no fluorescence was observed for the solution treated at 100°C while raw CDs solutions synthesised at 150, 200, 250 and 300°C fluoresced in yellow, orange, green and blue colour respectively (Figure 3.8b). These emissions were further studied in detail by using a fluorescence spectrophotometer.

Figure 3.8 Filtered raw CDs solutions synthesised at different temperatures of 100, 150, 200, 250 and 300°C under (a) normal light and (b) UV light.

From each sample, 100 µL of CDs solution was diluted with 3.0 mL of distilled water in a quartz cuvette and then measured using a fluorescence spectrophotometer. The optimal temperature was chosen based on the highest intensity, emission wavelength and shape of the emission curve of the CDs (preferably to be sharp and distinct from other emission peaks). Optimum fluorescence spectra for each temperature were measured (Figure 3.9). CDs solution treated at 100°C showed no fluorescence (not shown), indicating that the condition was not suitable for carbonisation to occur. At 150°C, green fluorescence with an emission peak at 515 nm was observed (Figure 3.9a), indicating that the carbonisation and polymerisation processes were started at this temperature. At 200°C, the CDs solution showed an increase in fluorescence intensity and revealed two fluorescent peaks with the first peak at λem=450 nm (blue) and the second peak at λem=595 nm (orange) (Figure 3.9b). Interestingly, both peaks showed different PL properties with changes in excitation wavelength (not shown). The first peak was red-shifted with increasing excitation wavelength, but the second peak did not shift and merely changed in PL intensity. At 250°C, the emission peak at 595 nm had 54 disappeared and only one peak was observed at 450 nm which was similar to the first peak from the previous temperature (200°C) but with a decrease in fluorescence intensity. At the final temperature of 300°C, an excitation-dependent fluorescence could be observed with the emission peak increasing and shifting continuously with the increase in excitation wavelength. In a similar finding from a study by Wang and co- workers (2015), it was reported that the formation of larger carbon particles at higher temperatures and longer heating duration could lead to the excitation-dependent fluorescence of CDs [148]. The excitation-dependent fluorescence is common feature in various types of CDs that is due to the surface state affecting the bad gap of CDs and also their carbogenic cores [144, 149].

Figure 3.9 Fluorescent emission spectra of CDs synthesised at (a) 150°C; (b) 200°C; (c) 250°C and (d) 300°C at their optimum emission peaks.

By using the orange emission at 595 nm as reference to determine the optimal temperature, the emission showed a maximum intensity at 200°C before disappearing at higher temperature (Figure 3.10). The hydrothermal treatment at 200°C provided the best intensity for emission wavelength at 595 nm and was set as the default temperature in subsequent CD syntheses.

Figure 3.10 The effect of temperature on the emission of CD at at λex = 400 nm, λem = 595 nm.

3.6.2.2 Duration of Synthesis CD solutions were synthesised at 200°C with different heating durations (1.0, 1.5, 2.0, 2.5, and 3.0 h) and then filtered, revealing different colours under normal light and UV light (Figure 3.11). CDs solution treated for 1.0 h was obtained as a pale yellow solution while the colour obtained for the CDs solutions treated for 1.5 h, 2.5 h, and 3.0 h was orange. CDs solution treated for 2.0 h was obtained as a reddish-brown solution. The orange or red colour of the solutions indicated the formation of aromatic compounds and oligosaccharides as result of the “polymerisation” step [150]. An observation under UV lamp (Figure 3.11b) revealed a pale green fluorescence for sample treated for 1.0 h, orange fluorescence for sample treated for 2.0 h and yellow fluorescence for the other samples treated for 1.5, 2.5, and 3.0 h. When treated at constant concentration and temperature, the diameter of the CDs produced would increase as the heating time was increased [146]. Emission wavelengths of CDs were discovered to be dependent on particle size, even under the same excitation light [87]. It is likely that the change in the emission colours of these CDs were caused by the change in particle size attributed to the heating time. 56

Figure 3.11 Filtered raw CDs solutions synthesised with heating durations of 1.0, 1.5, 2.0, 2.5 and 3.0 h for time optimisation under (a) normal light and (b) UV light.

Emission profile from CDs synthesised under different heating duration was investigated (Figure 3.12). CDs heated for 1.0 h showed a peak emission at 515 nm while CDs treated for 1.5 h revealed three emission peaks at wavelengths 430, 520, and 595 nm. CDs treated for 2.0 h showed that the number of emission peaks were now reduced to two at wavelengths 450 and 595 nm, with the latter emission showing higher intensity. CDs synthesised for 2.5 h showed two emission peaks similar to the previous CDs (2.0 h) but this time having the 450 nm peak with the lesser intensity. CDs synthesised for 3.0 h exhibited similar emission peak patterns with CDs synthesised for 2.5h. The only difference observed was the overall emission intensity for CDs synthesised for 3.0 h was decreased. Overall, the increase in heating time has induced a red-shift of CDs fluorescence. In a study of CDs prepared by hydrothermal method by Liu and co-workers (2007), they found that the emission wavelength is gradually red- shifted as the particle size increased [87]. It is suspected that the red-shift observed in this study is also caused by the increasing size of CDs. 57

Figure 3.12 Fluorescent emission spectra of CDs synthesised at (a) 1.0 h; (b) 1.5 h; (c) 2.0 h; (d) 2.5 h and (e) 3.0 h, at their optimum emission peaks.

A graph of emission at 595 nm against time was plotted (Figure 3.13). The emissions showed a steady increase up to 2.0 h before decreasing on further heating time. Hydrothermal treatment for 2.0 h provided the best emission at 595 nm wavelength and was set as the default time in subsequent CD syntheses.

Figure 3.13 The effect of heating time on the emission of CD at λex = 400 nm, λem = 595 nm.

3.6.2.3 Effect of Acid Ratio Emission profile from CDs synthesised with different phosphoric acid concentrations was observed (Figure 3.14). When using 8.5 % phosphoric acid, two apparent peaks at 425 nm and 520 nm were produced with excitation at 340 and 360 nm, respectively. At excitation wavelength of 390 nm, a third emission peak near 600 nm was recorded. When using 17.0 % phosphoric acid, the fluorescence emission of CDs under the typical measurement settings had exceeded the 1000 a.u.scale limit. Thus, the emission and excitation slits of the fluorometer were reduced from 10 nm to 5 nm in order to obtain the complete emission profile. From Figure 3.14b, the three peaks obtained were similar to the one in Figure 3.14a with the third emission peak becoming more noticeable at 595 nm. As the concentration of phosphoric acid used was increased to 25.5, 34.0, and 42.5 %, the emission peak at 520 nm became less and less noticeable and finally disappeared. Meanwhile, the third emission peak at 595 nm appears to be more defined. From these observations, it is noticed that increasing the concentration of phosphoric acid had overall shifted the fluorescence colour of the CDs from green to yellow and finally to orange. It is suggested the addition of phosphoric acid helped red-shift the CDs emission. A study by Bhunia and co-workers (2013) reported red-shifted emission profiles from a phosphoric acid-mediated CD synthesis [97]. The team demonstrated that the use of sulphuric acid with a carbohydrate would generally produce blue-green emission, whereas carbohydrates dissolved in phosphoric acid as the dehydrating agent produced CDs with red-shifted emission. They attributed the red-shift to a number of 59 factors including the various particle sizes as the key. Controlled by the dehydrating agent, as the size of CDs increase, the size of the sp2 domains can also increase. It is theorised that the increase in concentration of phosphoric acid in this study allows for larger particle sizes which in turn caused the red-shifting in CDs (Figure 3.14).

Figure 3.14 Fluorescent emission spectra of CDs synthesised with phosphoric acid concentrations of (a) 8.5 %; (b) 17.0 %; (c) 25.5 %; (d) 34.0 % and (e) 42.5%, at their optimum emission peaks. Cary Eclipse emission and excitation slit settings used for (a), (c), (d) and (e) is 10 nm while for (b) the slit size used is 5 nm. 60

A graph for the emission at 595 nm against phosphoric acid concentration used was plotted as shown in Figure 3.15. The emission showed a steady increase with regard to the increase in the concentration of phosphoric acid (8.5 to 25.5 %) before reaching a plateau at higher concentrations. In the optimisation studies for temperature, duration of synthesis and concentration of phosphoric acid used, the best conditions were chosen based on the highest intensity that can be obtained at 595 nm emission wavelength. In this case, raw CDs from phosphoric acid concentrations of 25.5, 34.0, and 42.5 % gave a similar intensity at 595 nm. The most noticeable difference between the CDs synthesised from these three concentrations is the appearance and intensity of the emission peak at 520 nm (Figure 3.14). At the highest phosphoric acid concentration (42.5%), the disappearance of the second peak gave a cleaner emission profile. This would provide less interference with the orange peak at 595 nm emission wavelength. Therefore, the use of phosphoric acid with the concentration of 42.5 % was preferable for subsequent CDs syntheses.

Figure 3.15 The effect of phosphoric acid concentration on the emission of CD at λex = 400 nm, λem = 595 nm.

3.6.2.4 CMC Concentration After filtration, 100 µL from each of the raw CDs synthesised from different CMC concentrations was added with 3.0 mL of distilled water and the emission profiles were obtained. From Figure 3.16a, three small emission peaks at 425, 520, and 595 nm were observed. When increasing the CMC concentration from 2.5 to 12.5 mg/mL (Figure 3.16a-e), only emission peaks at 425 and 595 nm showed an increase in the intensity

61 while the peak at 520 nm had disappeared when 7.5 mg/mL of CMC was used. Overall, changing of CMC concentrations during the synthesis did not cause shifts in emission peaks. However, increasing the CMC concentration increased the intensity of the CDs emission peaks. It is possible that at higher CMC concentrations, the CDs only increased in concentration, with little to no effect on the particle size and emission properties.

Figure 3.16 Fluorescent emission spectra of CDs synthesised from CMC of increasing concentrations (a) 2.5 mg/mL; (b) 5.0 mg/mL; (c) 7.5 mg/mL; (d) 10.0 mg/mL and (e) 12.5 mg/mL, at their optimum emission peaks.

A graph for emission at 595 nm against CMC concentration was plotted as shown in Figure 3.17. The increase of CMC concentration had caused a steady increase in the intensity of the emission peak. Although the result of the graph suggests that concentrations of CMC higher than the one used in this study (12.5 mg/mL) could provide brighter CDs, this would not be possible due to the solubility limit of CMC salt. The CMC concentration that provided the highest intensity, 12.5 mg/mL, was used for the subsequent syntheses of CDs.

Figure 3.17 The effect of concentration of CMC on the emission of CDs at λex = 400 nm, λem = 595 nm.

3.6.3 Effect of External Surroundings

3.6.3.1 Effect of Metal Ions As part of the characterisation of CDs synthesised in this study, the effect of metal ions on CDs were investigated. From the results in Figure 3.18, there were noticeable response from CDs when added with 50 µL of 0.1 M SnCl2, Al(NO3)3 and Fe(NO3)3 solution. The response seen in Figure 3.18a was mainly on the change of emission peak intensity at 450 nm. Upon addition of Sn2+, an increase in intensity (enhancement effect) was observed while a decrease in intensity (quenching effect) could be seen when Fe3+ was added. In Figure 3.18b, Al3+ and Sn2+ caused an enhancement response from CD at 490 nm emission peak. In Figure 3.18c, Fe3+ ions induced a quenching effect on the 595 nm emission peak.

Figure 3.18 Comparisons of CD samples added with 50 µL water and various metal ions solution under (a) 450 nm, (b) 490 nm, and (c) 595 nm emission wavelengths.

The increase in concentration of Al3+ had caused an enhancement of CDs intensity at 490 nm emission wavelength (Figure 3.19). There are numerous reports on the addition of Al3+ that causes enhancement on the photoluminescence of CDs, however, the exact mechanism behind the enhancement may vary depending on the functional groups of the CDs [151-153]. The presence of oxygen functional groups in the CDs was confirmed by FTIR in this study (Figure 3.7) and these functional groups were postulated to induce the enhancement effect of Al3+ ions via chelation-enhanced fluorescence (CHEF) mechanism. In a study by Fu and co-workers (2017), enhancement effect of Al3+ ions on CDs synthesised from electrolysis of graphite in NaOH solution have been reported [151]. They attributed the enhancement effect to CHEF mechanism. It was suggested that the abundant oxygen functional groups on the surface of CDs had chelation interaction with Al3+ ions that increased the rigidity of the CDs, which then led to fluorescence enhancement.

Figure 3.19 Normalised intensity of CD at λex=390 nm, λem=490 nm against concentration of Al(NO3)3.

The increase of Fe3+ concentration had caused a steady quenching effect on both emission peaks at 450 and 595 nm (Figure 3.20). The quenching caused by the addition of Fe3+ is quite commonly reported CDs. Since the CDs in this study were synthesised with phosphoric acid, it is likely that the surface of the CDs were doped with phosphate groups that may have interacted with Fe3+ ions and caused the quenching effect. Shangguan and co-workers (2017) suggested that the quenching interaction between CDs and Fe3+ may have originated from dynamic or static quenching [154]. Their group synthesised N/P co-doped CDs via hydrothermal method using adenosine-5'- 65 diphosphate (ADP) and adenosine-5'-monophosphate (AMP) as precursors, mixed with different phosphorus doping content. It was speculated that the CDs were quenched by Fe3+ ions due to the strong binding preference of Fe3+ towards the phosphate group of the surface of the co-doped CDs. The formation of Fe-O-P bond due to the Fe3+ complexing with the surface functional groups of CDs were confirmed with FTIR and static quenching was confirmed by carrying out fluorescence lifetime investigation. A similar complex may have occurred between the Fe3+ ions and CDs synthesised in this study, which subsequently quenched the fluorescence.

Figure 3.20 Normalised intensity of CD at (a) λex=350 nm, λem=450 nm and (b) λex=390 nm, λem=595 nm against concentration of Fe(NO3)3.

Increasing the concentration of Sn2+ had a linear enhancement effect on the blue peak of CDs at 450 nm emission wavelengths, even under different excitation wavelengths (Figure 3.21). Under the excitation wavelength of 390 nm, CDs showed a more significant enhancement effect with the intensity more than doubled after increasing the concentration of Sn2+ as compared to when the CDs are excited under 350 nm. To date, 66 there are no external reports on Sn2+ enhancement of CDs and the exact mechanism behind this enhancement remains to be explored. There was, however, a report on Sn2+ detecting CDs, although it was via the quenching of CDs fluorescence [155]. This report proposed that the complex formed between Sn2+ ions and the surface of CDs interferes with the initial energy transition that emit fluorescence; hence the quenching of the CDs. The Sn2+-induced fluorescence enhancement of CDs in Figure 3.21 may be ascribed to metal-enhanced fluorescence (MEF) in which the Sn particles could increase the local incident field of the CD and subsequently increasing the emission intensity [156]. Nevertheless, further study is still required to understand the detailed fluorescence enhancement mechanism.

Figure 3.21 Normalised intensity of CD at (a) λex=350 nm, λem=450 nm and (b) λex=390 nm, λem=450 nm against concentration of SnCl2.

3.6.3.2 pH The CDs in this study had showed a pH-dependent emission (Figure 3.22). There was an enhancement on the CDs when pH was increased from 1 to 3 and the intensity subsequently decreased gradually to its initial intensity at pH 7. The intensity then continued to decrease as the pH was further increased from pH 7. The decrease in intensity of the orange peak was caused by blue-shift; the shifting of CDs emission to shorter wavelengths. As a result, neutralised CDs fluoresced in yellow or green colour as shown in Figure 3.3 and 3.4 in the previous sub-chapter 3.6.1. It has been indicated that the pH conditions are able to modify the nature of energy levels and consequently modify the type of electronic transitions involved in CDs [157]. It is postulated that the effect was due to the deprotonation of the oxygen containing functional groups that are present on the surface of CDs, which consequently leads to the gradual creation of chemically different CDs species as pH condition is increased. Depending on the behaviour of CDs towards pH change, they can be used for pH sensing via ratiometric changes in fluorescence emission. Further studies can be done to explore the potential use of this CD as a pH sensor.

Figure 3.22 Graph of pH curve of CDs at λex = 400 nm, λem = 595 nm.

3.7 Conclusions

Synthesis of CDs with carboxylic acid groups from CMC as the carbon precursor was successfully carried out using optimised parameters. The addition of phosphoric acid in the synthesis resulted in the production of orange coloured CDs which is desirable and preferable for use in bioimaging applications. However, the CDs produced was acidic and had to be neutralised before they could be used for further characterisations and modifications. The orange emitting property did not remain in the CDs after going through neutralisation, isolation and drying process as the orange emission peak shifted to lower wavelengths, which resulted in blue-green emitting CDs. Although the CDs do not possess the desired optical properties after isolation, the carboxylic groups present on its surface indicated the CDs are still suitable for conjugation purposes.

4 Conjugating Carbon Dots with GAPDH Probe

4.1 GAPDH Probe Design

The gene chosen for this project was preferred to be commonly and constantly expressed in the model cultured cell line. Housekeeping genes are typically genes that are involved in the maintenance of basic cellular functions and are expected to maintain a constant level of expression in all cells regardless of developmental stage, tissue types, cell cycle state, or external signal [158]. Therefore, housekeeping genes are often used as control genes in experiments such as reverse transcription polymerase chain reaction (RT-PCR) and in situ hybridisation.

Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) is a commonly used housekeeping gene for gene expression data in human tissues. It is expressed as GAPDH mRNA in at least 72 types of tissues in the human body, which include skin, fat, neurological, reproductive, respiratory, cardiovascular, urinary, endocrine, and other tissue groups [159]. GAPDH serves as an enzyme that catalyses the sixth step in the metabolic pathway of glycolysis which converts glyceraldehyde 3-phosphate to D- glycerate 1,3-bisphosphate. Other than glycolysis, GAPDH has multifunctional properties associated with DNA repair [160], cell apoptosis [161], tRNA transport [162], membrane fusion [163], and vesicular transport [164].

A short oligonucleotide probe was designed by choosing a suitable target sequence from the Homo sapiens glyceraldehyde-3-phosphate dehydrogenase isoform 1 (GenBank accession no: NM_002046) gene sequence to complement the GAPDH mRNA [165]. The target sequence chosen for this study is 25 bases long with reference to the GAPDH gene specific probe sequence used previously in a paper [166]. In this paper, the GAPDH probe used is: GAPDH TaqMan probe 5'-VIC- TCCGACGCCTGCTTCACCACCTTCT-MGBNFQ-3'.

The GAPDH sequence used for this project is similar to the sequence mentioned above. A different modification was done to the probe instead by attaching a primary amine on the 5’ end of the probe with the 5’Amino Modifier C6 with spacer arm of 6-7 atoms. The probe synthesised for this project is: 5’-AmMC6- TCCGACGCCTGCTTCACCACCTTCT-3’. The GAPDH probe ordered is a 25 nmole

70 scale, standard desalted, DNA oligo purchased from Integrated DNA Technologies Pte. Ltd.

4.2 CD-aniline EDC-NHS Conjugation

Conjugation of the CDs to the amine-modified GAPDH mRNA probe was performed via carbodiimide crosslinker chemistry. This reaction involves the use of carboxyl- reactive chemical groups such as 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) to crosslink carboxylic acids to primary amines via amide bond formation. The presence of carboxylic acid functional groups on the CDs had been established through FTIR analysis; however, to minimise wastage of the GAPDH probe, the optimisation of the EDC-NHS conjugation protocol had to be carried out beforehand using a substitute compound for the GAPDH probe. The amount and concentration of GAPDH probe was also too low for analysis.

The successful conjugation of CDs to an amine group was confirmed using FTIR analysis. Aniline (Figure 4.1), C6H5NH2, was used as a substitute for the GAPDH probe due to its simple structure of having a single amino group attached to a phenyl group.

Figure 4.1 Chemical structure of aniline.

N-hydroxysuccinimide (NHS) and EDC were both purchased from Sigma-Aldrich. Activation buffer or MES buffer (0.1 M) was prepared by dissolving 0.8 g of 2-(N- Morpholino) ethanesulfonic acid (MES) (purchased from Sigma-Aldrich) in 30.0 mL of distilled water with adjustment to pH 6 using 0.5 M NaCl and 10 M NaOH. Aniline (analytical grade EMSURE® purchased from Merck Sdn. Bhd.) with concentration of 10.0 mg/mL was prepared by dissolving 100.0 mg aniline dissolved in 10.0 mL 1X PBS and stored at 2-8°C.

Firstly, 20.0 mg of dried CDs were dissolved in 2.0 mL activation buffer to make 10.0 mg/mL. After that, 0.8 mg of EDC and 1.2 mg NHS were added separately and mixed 71 well. The reaction was left to complete for 15 min at room temperature. Then, 2.0 mL of 10.0 mg/mL aniline was mixed into the activated CDs solution and let sit for 2 h at room temperature. The CD-aniline solution was then stored at 2-8°C. Before sending the CD-aniline sample for FTIR analysis, the solution was dried overnight in a drying oven at 75°C. The solid CD-aniline was compared with dried CDs and aniline.

The utilisation of carbodiimide EDC and Sulfo-NHS for carboxyl-to-amine crosslinking is shown in Figure 4.2. In this reaction, CD with carboxylic group is represented by molecule (1) and aniline with a primary amine group is represented by molecule (2). Often included in this reaction is N-hydroxysuccinimide (NHS) or its water-soluble analogue (Sulfo-NHS) to improve reaction efficiency or create dry-stable (amine reactive) intermediates for storage or later use. The EDC reaction forms an active intermediate that is unstable and readily hydrolysed by water. This competing reaction with water can cleave the active intermediate to regenerate the carboxyl group (see middle pathway). The addition of NHS prevents rapid hydrolysis by allowing the formation of a more stable NHS ester intermediate that slowly reacts with primary amines to form a stable amide bond (see bottom-most pathway). Activation using EDC and NHS is most efficient between pH 4.5 and 7.2. Therefore, MES buffer with pH 6 is preferred for the activation reaction [167]. The activation buffer also has to be free from any primary amine or carboxyl groups, which would compete with the activation reaction. The final product expected from the crosslinkage of CD and aniline is a stable conjugate with amide bond referred to as ‘CD-aniline’ (see top-most pathway). The formation of this amide bond in CD-aniline was confirmed by FTIR analysis.

Figure 4.2 Schematic of Sulfo-NHS plus EDC (carbodiimide) crosslinking reaction [168].

Figure 4.3 shows a comparison of FTIR spectra for CD-aniline, aniline and CD. The FTIR spectrum for CD in Figure 4.3 showed the presence of carboxylic acid group characterised by the O-H stretch overlapping with C-H stretch in the region 3300-2500 cm-1, the carbonyl C=O stretch at 1700-1690 cm-1 and the O-H bend from 1440-1395 cm-1 [143]. Next, the characteristic peaks of aniline in its FTIR spectrum were also labelled. In the aniline spectrum in Figure 4.3, the N-H stretching of a primary amine at 3500 cm-1 and 3350 cm-1 was observed. The peak at 1620 cm-1 corresponds to N-H bending of the amine group and the peak at 1500 cm-1 corresponds to the C=C stretching for benzenoid ring of the aniline [169]. In addition, the C-N stretch for aromatic amine at 1270 cm-1 could be observed and the N-H peak at 750 cm-1 is attributed to the N-H wag vibration. According to the amide bond of the stable conjugate in Figure 4.2, the expected vibrations in the final CD-aniline product are N-H stretching for secondary amine and C=O stretching for secondary amide. From the CD- aniline FTIR spectrum in Figure 4.3, the presence of these vibrations were confirmed by the peak at 3350 cm-1 corresponding to the N-H stretch vibration for secondary amine and also the appearance of peak 1680 cm-1 corresponding to C=O stretch vibration for secondary amide. After establishing the amide formation, conjugation of CD with GAPDH using EDC-NHS were carried out.

Figure 4.3 FTIR spectra of CD-aniline, aniline and CD.

4.3 CD-oligonucleotide EDC-NHS Conjugation

GAPDH oligonucleotide probe stock solution were prepared according to the manufacturer’s manual by mixing 267 µL nuclease-free water into the vial to make a 100 nM probe solution. The probe solution were later on split into 20 µL aliquots and stored in -20°C for later use. Similar to the protocol used on CD-aniline conjugation, the volumes and concentrations for activation buffer, EDC, and NHS used were the same.

Three different concentrations of CDs, 10.0, 15.0, and 20.0 mg/mL were used to prepare the CD-GAPDH. Firstly, 10.0 mg of CDs solid was weighed in a microcentrifuge tube and 1.0 mL of activation buffer was added to dissolve the CDs. This CDs solution was then filtered through a 0.22 µL syringe filter. Then, 1.0 mg of EDC and 2.4 mg of NHS was added separately into the solution and mixed well. The mixture was left to sit at room temperature for 15 min to allow reaction to complete. Meanwhile, a GAPDH aliquot was thawed. Finally, 20 µL (100 nM) of the GAPDH probe was transferred and mixed into the activated CDs solution and left to incubate at room temperature for 2 h. The CD-GAPDH probe was stored in 4°C until further use.

4.4 Conclusions

An oligonucleotide probe specific to housekeeping gene, GAPDH mRNA, was designed. A primary amine group was included to the 5’ end of the probe for conjugation purposes. Conjugation of CDs and GAPDH probe was carried out using EDC-NHS crosslinking protocol. Due to limited amount of GAPDH probe and limitations on the FTIR spectrometer, direct confirmation of CD-GAPDH conjugation using this method could not be done. Aniline was used as the substitute for the probe in this crosslinking protocol for its simple chemical structure consisting of a primary amine attached to a phenyl ring. The amide bond formation between CDs and aniline using carbodiimide crosslinker chemistry was confirmed through FTIR analysis, suggesting successful conjugation.

5 CD-FISH

5.1 Preliminary Testing for CDs Imaging

Before any imaging studies using the CD-GAPDH probe were employed, preliminary tests were carried out to assess for penetration of CDs across the cell membrane by looking at the presence of fluorescence in the cell plasma. Mammalian cell line cultures require frequent subculturing for maintenance, which are costly and tedious. Hence, single layer onion cells were used as a substitute for mammalian cells in this preliminary testing. Since a fluorescence microscope was not readily available at the time of this experiment, a simple fluorescence imaging set-up was prepared using a normal microscope with different types of cameras for imaging to view neutralised CD in cells.

Cells to be incubated with CDs were prepared by carefully peeling out single layer onion epidermis using a pair of tweezers and then drying them overnight at room temperature. 20.0 mg of dried CD was weighed and dissolved in 1.0 mL distilled water to make 20.0 mg/mL. Dried onion layer was incubated overnight in the CDs solution. After incubation, the onion epidermis was laid flat on a glass slide and covered with a coverslip. The cells were viewed under an inverted microscope.

5.1.1 Webcam and Smartphone Camera The addition of a UV light source on a Nikon Eclipse Ti inverted microscope was done by placing a commercially available 390 nm wavelength UV light above the condenser unit (Figure 5.1). A black box was used to cover the whole microscope to prevent stray light interference. Images were taken using a Logitech C160 5 MP webcam and an iPhone SE 12 MP camera focusing through the eyepiece of the microscope and the comparisons were made.

Figure 5.1 (a) Nikon Eclipse Ti inverted microscope. (b) Placement of 390 nm UV torchlight above condenser unit.

The images of CDs incubated onion cells were captured and compared (Figure 5.2). Under white light, both Logitech webcam and iPhone SE camera showed similar images with the only difference being the higher image quality was captured using phone camera. Under UV light, despite showing the outline of onion cells both cameras showed oversaturation of blue and violet colour. Furthermore, the intensity of the UV light was non-adjustable. These limitations made viewing of the fluorescence of CDs in the cells impossible. The inability of both cameras to manually focus on images also made photo taking difficult. Hence, these cameras were deemed unsuitable for the preliminary imaging test and an alternative camera that allows for manual focusing and exposure control was used for capturing images.

Figure 5.2 Images of onion cell epidermis after incubation with CDs solution. Top: Onion cells image under white light (left) and UV light (right) taken with Logitech webcam. Bottom: Onion cells image under white light (left) and UV light (right) taken with iPhone SE camera.

5.1.2 Microscope Camera A compatible microscope camera, Nikon SMZ 745T, was installed on the microscope (Figure 5.3). To overcome oversaturation of UV light, light source was placed on the side of the sample instead of through the condenser. Focus and exposure adjustments of the camera were done on a computer via NIS-Elements D program. The samples used for this test were onion epidermis incubated in highlighter solution and onion epidermis incubated with 20.0 mg/mL CDs solution for comparison. Captured images were later on adjusted for colour and brightness enhancement using the lookup table (LUT) available in the NIS-Elements D program.

Figure 5.3 Nikon Eclipse Ti inverted microscope with Nikon SMZ 745T microscope camera installed.

5.2 HaCaT Cell Culture

Human adult low calcium high temperature keratinocytes (HaCaT) cells are immortalised normal keratinocytes that stably expresses GAPDH gene [170]. The HaCaT cell line was cultured in a 96-well plate. Subsequently, the fixation and hybridisation steps of FISH technique were applied on the cells in the same well plate.

5.2.1 Reviving Cryo-preserved HaCaT Cells DMEM (10% FBS and 1% penicillin/streptomycin) media was pre-warmed at 37°C for at least 15 min before use. The cryovial containing the frozen HaCaT cells was removed from the liquid nitrogen storage and immediately placed into a 37°C water bath to be thawed. In a biosafety cabinet (BSC), the vial was wiped with 70% ethanol before it was opened. The cell suspension was briefly homogenised by slowly pipetting the content of the tube and then transferred into a falcon tube with 10.0 mL of DMEM. The content of the tube was briefly mixed and the cell pellet was collected by centrifugation at 1,500 rpm for 5 min. Then, supernatant was discarded and the cell pellet was re- dispersed in 10.0 mL of fresh DMEM. The centrifugation step was repeated another time to collect the cell pellet. Subsequently, the supernatant was discarded and the cell pellet was re-dispersed in 10.0 mL DMEM which later on was transferred into a T-25

79 cell culture flask. The cells were incubated in a cell culture incubator at 37°C with 95% air and 5% CO2.

5.2.2 Sub-culturing HaCaT Cells DMEM (10% FBS and 1% penicillin/streptomycin) media was pre-warmed at 37°C for at least 15 min before use. Existing media was discarded from cell culture flask through aspiration. The cells were rinsed with 10.0 mL of 1X PBS. The PBS was aspirated off and 2.5 mL of trypsin-EDTA was added. The flask was then incubated for approximately 10 min to fully detach the cells. Later on, the flask was rinsed with an aliquoted 10.0 mL DMEM media to pool the cells in the supernatant and to deactivate trypsin. From this suspension, 1.8 mL was transferred into a T-75 flask containing 10.0 mL fresh DMEM. The remaining cell suspension was used for sub-culturing into a 96 well plate. The cell concentration was calculated using a haemocytometer beforehand to determine the amount to be added in each well. The stock cell concentration were diluted with DMEM to an optimum cell concentration of 20,000 cells/mL. Then, 100 µL of the diluted cell suspension was added to each well. Cells were incubated in the incubator until confluency was reached.

5.3 Fluorescence In Situ Hybridisation

The following reagents were used for FISH protocol. TE buffer was prepared using 10 mM Tris, adjusted to pH 8.0 with HCl, mixed with 1 mM EDTA. Hybridisation buffer was prepared by mixing 0.9 M NaCl, 20 mM Tris-HCl with pH 7.2, 50% formamide with 0.01% SDS added last. Pepsin working solution (0.5% in 5 mM HCl) was prepared by mixing 0.05 mL of 1% pepsin stock solution with 4.95 mL of distilled water. Washing buffer was prepared by mixing 20 mM Tris-HCl with pH7.2, 0.01% SDS, 40 mM NaCl and 5 mM EDTA. Mounting oil were made fresh before use each time using 1.0 mL of 1X PBS mixed in 9.0 mL of glycerol. Another optional set of mounting oil with antifade was prepared by adding 10.0 mg of p-phenylenediamine (PPD) and mixing it thoroughly into the solution. Mounting oil with added PPD were stored in - 20°C when not in use and covered in aluminium foil to prevent exposure to light.

5.3.1 Fixation of Cells All buffer used were pre-warmed at 37°C beforehand. In each well, the media was removed and replaced with 100 µL of TE buffer to wash the cells. The well plate was given a little shake and the supernatant was later on removed. Then, the cells were fixed with 50 µL PBS and 150 µL of 10% buffered formalin at room temperature for 1 hour. After that, formalin was removed and the cells were washed with 100 µL 1x PBS. The supernatant was discarded and 50% (v/v) ethanol/PBS solution was added. The fixed cells were stored at -20°C until further analysis.

5.3.2 Hybridisation Supernatant wasremoved from fixed cell cultures before undergoing dehydration in a series of ethanol. This was carried out by covering each wells with about 100 µL of 50, 80, and 100% ethanol solution for 3 min each and then air-dried. The cells were then treated with 100 µL 0.01% pepsin for 10 min at 37°C. The reaction was stopped by adding the same volume of ice-cold PBS. Supernatant was removed and cells were subsequently air-dried. Next, the probe solution (10 µL CD-GAPDH probe mixed with 40 µL of hybridisation buffer) was added to the cells and left to incubate overnight in a dark, humid chamber at 45°C. In addition to this, a negative control was prepared by mixing 10 µL of distilled water with 40 µL of hybridisation buffer. The probe solution would also be compared with a solution of 10 µL CD (15.0 mg/mL) mixed with 40 µL of hybridisation buffer. On the following day, the supernatant was removed and unbound probes were washed off by incubating the cells in 100 µL of pre-warmed washing buffer for 15 min at 45°C. The cells were then gently rinsed with 200 µL of clean dH2O and air-dried. Cell samples were embedded in mounting oil before being imaged with BioTek™ Cytation™ 3 Cell Imaging Multi-Mode Reader with GFP filter cube.

5.4 Results and Discussion

5.4.1 Preliminary Testing for CD Imaging A comparison of the visibility of CDs and highlighter dye in onion cells under UV excitation at 390 nm are shown in Figure 5.4. In image 5.4a, the cells could be faintly seen but showed no fluorescence, which was expected for distilled water. The commercial orange highlighter dye used for this work normally contains a mixture of xanthene and coumarin derivatives namely basic red 1 and basic yellow 40 [171]. Basic red 1, also known as rhodamine 6G, has a maximum absorption at 530 nm wavelength with maximum emission near 565 nm wavelength [172]. Basic yellow 40 is a coumarin- based dye that fluoresces in bright yellow or green between 365 and 485 nm when excited under UV light [173]. These properties were observed in Figure 5.4b where the green florescence visible in the cell walls of the sample originated from the coumarin- based dye, while fluorescence from rhodamine 6G was low to none as the excitation wavelength 395 nm used in this study did not fall into the optimum absorption spectrum of rhodamine 6G. In Figure 5.4c, no fluorescence was observed from the CDs solution incubated with the onion cells. Under 395 nm excitation wavelength, the CDs solution should fluoresce at 500 nm emission peak. The microscope camera, however, did not capture this property as the emission from CDs was not bright enough to be detected and the system was not sensitive for capturing simple fluorescence. Consequently, imaging work was proceeded with utilising a fluorescence microscope equipped with special optical filters to enhance sample fluorescence.

Figure 5.4 Fluorescence images of onion epidermis incubated in (a) distilled water, (b) highlighter dye, and (c) 20.0 mg/mL CDs solution illuminated under UV light. Photos were taken with Nikon SMZ 745T microscope camera.

5.4.2 Fluorescence In Situ Hybridisation The use of mounting media was optimised beforehand for fluorescence imaging. Generally, mounting media may be added with an antifading agent or free radical scavenger to help bring out a brighter signal from samples and doing this also prevents fluorescent dyes from photobleaching. P-phenylenediamine (PPD) was added as the antifade agent in the mounting oil for its strong antioxidising property. The use of mounting oil with and without added PPD were compared in Figure 5.5. On HaCaT cells with no probe added (Figure 5.5a), no fluorescence signal was observed when mounting oil with no PPD was used whereas with PPD used, the cells showed fluorescence signal and the background was stained. In Figure 5.5b, cell samples incubated with CD-GAPDH probe viewed in mounting oil both with and without PPD showed fluorescence signal. The cells in mounting oil with PPD appeared brighter and has significant background staining. The use of PPD in mounting oil was unfavourable as the background staining made it difficult to discern fluorescence signal from the probe. Moreover, the fluorescence from the probe was bright enough to be detected without requiring any antifade agent. Thus, only mounting media with no PPD were used on the cell samples.

Figure 5.5 A comparison of fluorescence images of fixed HaCaT cells incubated in (a) hybridisation medium without probe and (b) 15.0 mg/mL CD-GAPDH probe viewed in mounting oil without p- phenylenediamine and with p-phenylenediamine. All images were taken under the same exposure settings. 83

The efficacy of the prepared CD-GAPDH probe was tested. FISH procedure were performed on three sets of HaCaT cell line by adding hybridisation buffer with no probe, CDs and CD-GAPDH probe. A comparison made in Figure 5.6 showed that fluorescence signal could be observed from cells treated with CDs and CD-GAPDH probe. The sample with hybridisation buffer with added CDs in Figure 5.6b was expected to not yield any signal, similar to cells added with hybridisation buffer without probe in Figure 5.6a. The reason being that there was no surface modification done on the CDs to attach with specific components in the cell and the washing step in the FISH procedure would have removed any unbound components from the fixed cell line. However, the fluorescence signal showed uniform staining of CDs in whole cells inclusive of the nucleus and cytoplasm, which suggests that the bare CDs attach non- selectively to these cell components (Figure 5.6b). Although it is unclear why CDs stain, a somewhat similar observation was reported in an experiment carried out by Kong and co-workers (2014) where synthesised CDs were reported to permeate and stain cells [174]. In their study, green CDs were prepared from ethylene glycol and polyethelyne glycol using sodium hydroxide-assisted reflux method with no further functionalisation steps. The CDs were cultured with HeLa cells, which were later washed and fixed. Their fluorescence images showed the CDs specifically targeting the nucleus and nucleolus but the mechanisms behind the staining were not postulated. A possible explanation for the non-specific staining of the CDs used in this study could be the electrical or coulumbic interactions, which are involved in the acidic and basic dyeing of fixed tissues also known as electrostatic binding or salt links. Under acidic conditions, most proteins in fixed tissues carry an overall positive charge and when applied, acidic dyes stain protein non-specifically [175]. The affinity and staining selectivity are controlled by the electric charge of the dyes. The CDs used for this study contains carboxylic acid groups which are negatively charged and were likely to have electrical interactions to the biomolecules in the fixed HaCaT cells.

Figure 5.6 Fluorescence images of fixed HaCaT cells incubated with different samples: (a) hybridisation medium without probe, (b) 15.0 mg/mL CDs, and (c) 15.0 mg/mL CD-GAPDH probe. All images were taken under the same exposure settings.

Interestingly, the cells treated with CD-GAPDH probe (Figure 5.6c) displayed brighter fluorescence compared to the cells stained by bare CDs (Figure 5.6b) indicating that a higher amount of CDs were present in the cells. The fluorescence images of cells with CDs and CD-GAPDH probe were zoomed in and compared to show a clearer distinction between the fluorescence signals in Figure 5.7. Cells treated with CDs showed a dimmer and more uniformly distributed fluorescence when compared to cells treated with CD-GAPDH probe. In Figure 5.7b, the brightness of fluorescence signal was uneven across the cells and clusters of small bright spots could be seen. This observation is postulated to be due to both the non-selective staining property and selective GAPDH labelling property of the CD-GAPDH probe manifesting together. Although the CD-GAPDH probe solution consisted of modified CDs, there were possibly presence of CDs that were not conjugated to any oligonucleotide in the mix. The bright dots pointed by the arrows were likely to be spots where denser amount of GAPDH mRNA were present and had hybridised with the probe. In addition, the brighter fluorescence of the whole cells may also be attributed to the presence of GAPDH mRNA in the plasma membrane and cytosol. In general, GAPDH are mainly localised to the plasma membrane and cytosol while in certain types of cells such as lung epithelial cells, they are also present in the nuclear membrane and vesicles such as shown in the samples in Figure 5.8 [176].

Figure 5.7 A zoomed in image comparison between a) CDs and b) CD-GAPDH probe in HaCaT cells. Arrows are pointed to clusters of bright spots likely to be GAPDH mRNA. Same contrast adjustments were applied on both images to enhance fluorescence signal.

In Figure 5.8, specialised antibodies were used to show GAPDH (in green) localised to the plasma membrane and cytosol in U-251 MG human malignant glioblastoma cells. In A549 adenocarcinomic human alveolar basal epithelial cells, GAPDH was localised to the nuclear membrane, vesicles and plasma membrane. GAPDH localised to the vesicles in A549 cell line was seen as clusters of bright spots. This characteristic is reminiscent to the bright spots seen in the HaCaT cells in Figure 5.7b implying that GAPDH was also localised to the vesicles in HaCaT cells. Despite the interference from non-selective staining, the irregular fluorescence signals and bright spots suggested that CD-GAPDH probe had been successfully hybridised to GAPDH mRNA.

Figure 5.8 Localisation of GAPDH in U-251 MG cell line (left) and A549 cell line (right). The nucleus is stained in blue, microtubules in red and GAPDH in green. [176]

5.5 Conclusions

A preliminary test to check for penetrability of CDs through cell membrane was performed by incubating onion cells in neutralised CDs solution. A simple fluorescence imaging set-up was attempted by shining UV light on the onion sample viewed under a normal microscope with a camera system installed. The CDs, however, could not be viewed under this set-up as it was not sensitive enough to capture the fluorescence.

HaCaT cell line was cultured, fixed, and then incubated with the as-prepared CD- GAPDH probe for in situ hybridisation. Cell images were taken with a fluorescence microscope and comparisons were made between CDs and CD-GAPDH probe. The images reveal that the CDs have stained the whole cells non-selectively. This property interfered with the fluorescence signal coming from the CD-GAPDH probe. Consequently, localisation of GAPDH mRNA in cells could be difficult to achieve due to the overlapping signal with unforeseen non-selective staining property of the CDs. On closer inspection, unlike with just CDs, HaCaT cells incubated with CD-GAPDH probe showed a brighter fluorescence signal with clusters of small spots which were postulated to be GAPDH localised to the plasma membrane, cytosol and vesicles.

Despite the limitations, this outcome suggests that the conjugation of CDs to GAPDH oligonucleotide probe was successful and the hybridisation of CD-GAPDH probe with target nucleotide in the cell had occurred. For improvement, this technique could be applied to other CDs that do not stain non-specifically on cell components. Thus, CDs have great potential as fluorescent labels for FISH probes.

6 Conclusion

6.1 Summary

The viability of CDs as fluorescent label in in situ hybridisation probe was assessed in this study. This study have presented the synthesis of CDs with carboxylic acid functional groups from hydrothermal treatment of carboxymethylcellulose in phosphoric acid solution. The as-prepared CDs were then characterised using spectrofluorometer, FTIR, and SEM. Subsequently, the CDs were neutralised, isolated and conjugated to an oligonucleotide probe designed specifically for GAPDH mRNA. The conjugated CD-GAPDH probe were applied on cultured HaCaT cell line for fluorescence in situ hybridisation. Probe hybridisation in HaCaT cells was confirmed using fluorescence microscopy.

The CDs synthesised under optimised parameters had an orange emission that peaked at 595 nm wavelength when excited at 400 nm wavelength. SEM analysis revealed that the particle size of CDs averaged at 0.134 µm with 0.055 µm standard deviation. FTIR spectroscopy confirmed the presence of carboxylic acid groups on the CDs. External effects on CDs such as the presence of different metal ions and pH were also assessed. CDs showed change in signal intensity when Al3+, Fe3+, and Sn2+ ions were introduced. Al3+ and Sn2+ ions induced enhancement in the fluorescence intensity of CDs while Fe3+ ions caused quenching. The CDs were also revealed to have pH-dependent emission. As the pH was changed from acidic to basic, blue-shifting occurred to the CDs resulting in the change of emission colour. Consequently, after the CDs were neutralised and isolated, the emission became yellow in colour. The neutralised CDs were then dried for conjugation purposes later on. When the dried CDs were reconstituted in water, the CDs emission became green.

A 25 base pairs long GAPDH specific oligonucleotide probe was designed with a primary amine attached to the 5’ end. CDs were conjugated to the oligonucleotide probe via carbodiimide crosslinker chemistry using coupling agents EDC and NHS. The conjugation procedure was tested using aniline as a replacement for the oligonucleotide probe and FTIR analysis confirmed the successful formation of carbodiimide.

This conjugated CD-GAPDH probe was incubated in fixed HaCaT cell line for hybridisation with GAPDH. Comparisons were made between cell samples incubated 88 with neutralised CDs and CD-GAPDH probe. Fluorescence images revealed that the CDs stain the cell components non-specifically which appeared as a uniform fluorescent stain across the whole cells. This property interfered with the fluorescence signal from CD-GAPDH probe, which complicated the confirmation of hybridisation and localisation of GAPDH in the HaCaT cell. Nevertheless, clusters of fluorescent spots and a brighter, uneven fluorescence signal appeared in cells incubated with CD-GAPDH probe indicating that probe hybridisation had successfully taken place. On closer observation, the GAPDH may likely be localised to the plasma membrane, cytosol and vesicles in HaCaT cells.

Although improvements are still needed to remove the non-specific staining property, CDs have shown great potential as biosensors and can be used as an alternative to traditional fluorophores. The same conjugation and in situ hybridisation procedure can be repeated with other types of suitable CDs to detect specific biomolecules. Furthermore, the CDs this in study also showed potential as pH sensor and metal ions sensor.

6.2 Future Directions

A study on surface capping of CDs to hinder non-specific staining should be carried out to improve the performance of CD-oligonucleotide probe. This would also be useful for other types of CDs that could stain on cells. In addition, the CDs from this study are versatile and can be explored further for other practical uses particularly in pH sensing and metal sensing applications. A more detailed study on the efficiency and limit of detection for Al3+, Fe3+, and Sn2+ metal ions are required.

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