Nanoarchitectured Point-Of-Care Detection System for Clinically Relevant Biomarkers Mostafa Kamal Masud Master of Science (MS)

Nanoarchitectured Point-of-Care Detection System for Clinically Relevant Biomarkers Mostafa Kamal Masud Master of Science (MS)

A thesis submitted for the degree of Doctor of Philosophy at The University of Queensland in 2020 Australian Institute for Bioengineering and Nanotechnology (AIBN)

Abstract The detection of disease-specific biomarkers such as proteins (particularly autoantibody) and microRNA (miRNA) are prerequisites to understanding their physiological and biological functions, early diagnosis, the prognosis of the disease and their treatment. Despite the numerous molecular biology and analytical techniques, their detection approaches are largely narrowed to laboratory-based molecular biology techniques. While the analytical performance and reliability of these approaches are admirable, most of these methods necessitate enzymatic amplification, cumbrous sample pre-treatment, multi-step assay protocol, sound technical personnel and expensive maintenance. The cutting-edge electrochemical and optical approaches are relatively simple and rapid and provide sensitive detection in a portable arrangement, however, target-specific transduction surface modification is vital to achieving the selectivity of a functional biosensor. With the progress of nanotechnology, nanostructured magnetic materials have aroused enormous interest in the arena of biosensing and biomedicine owing to their flexible and modular structure, nanosize, low toxicity, biocompatibility, intrinsic functionalities. Magnetic nanoparticles inimitably combined with the dimension of more modest size or the same size of molecular analytes and henceforth retain enormous potentiality in isolation and purification of target molecules, signal transduction, signal generation and signal enhancing steps in biosensing. Nonetheless, engineered multifunctional magnetic nanostructure-based diagnostics that could offer the advantages of excellent stability in a complex biomatrix, easy and alterable biorecognition of ligands, antibodies, and receptor molecules and unified point-of-care integration have yet to be achieved. This PhD project endeavours to engineer the nanostructure-based strategies for developing an inexpensive, specific, sensitive, and portable point-of-care diagnostic platform for clinics. This thesis intensively studies the bio-favourable nanostructure synthesis, their biofunctionalisation, intrinsic properties and cutting-edge nanostructure-based strategies for the detection of clinically-relevant biomarkers. Besides, the biogenesis, diagnostic, and prognostic potential of miRNA biomarkers followed by a comprehensive evaluation of recent progress in the development of nanostructure-based electrochemical miRNA biosensors are reviewed. I initially reported on the synthesis of mesoporous iron oxides with two different crystal phases named α- and γ-Fe2O3 to examine and understand the phase-dependent behaviours towards the magnetism and peroxidase mimetic activity. The cubic γ-Fe2O3 phase exhibited much higher activity and presented their superior aptness for biosensing. I have then designed and developed a new class of gold-loaded superparamagnetic mesoporous iron oxide nanocube, which superbly shows intrinsic nanozyme (peroxidase-mimetic) activity and

electrocatalytic activity towards different redox molecules. Based on these promising activities, I have developed a set of biosensing platform (three novel readout scheme) that facilitates simple, rapid, and inexpensive analysis of autoantibody and miRNA biomarkers. First, considering the promising nanozyme activity of developed nanocube a specific and sensitive autoantibody sensor was developed for detecting different stages of ovarian cancer, where the nanozyme potentially replaces the natural enzyme (horseradish peroxidase-HRP). The nanocube is purposefully loaded with nanosized (~ 2 nm) gold so that the nanocube can achieve good biocompatibility, well dispersibility in body fluids and can easily immobilise reporter or capturing antibody. Besides, the nanocube can easily achieve miRNA capture through favourite gold-DNA/RNA affinity interactions. By employing this gold-RNA affinity interaction and the electrocatalytic activity of this nanocube, I have then developed an amplification-free electrochemical miRNA sensor for the detection of esophageal cancer-specific mRNA (miR- 21). This forthright sensor enables the 10 pM level of detection by using the direct adsorption of magnetically isolated and purified miRNAs onto nanocube-modified disposable electrodes. Succeeding in advance of this proof-of-concept detection system, I endeavoured to address the growing thirst for detecting the ultralow levels of miRNAs from the complex biological sample through the development of another novel readout system. In this approach, the electrocatalytic activity of nanocube towards the redox reaction of methylene blue (MB) was coupled with 3-/4- 3-/4- [Fe(CN)6] to form an MB/[Fe(CN)6] electrocatalytic redox cycle. The combination of both electrocatalytic activity and redox cycling, the sensor achieved detection of 100 aM of miRNAs from motor neuron disease (MND) patient samples. In my final detection strategies, I prolonged the approach towards a translational- focused assay platform, where a mesoporous gold electrode was engineered which will be used for direct adsorption of magnetically isolated and purified miRNA followed by differential pulse voltammetric (DPV) interrogation. All of the reported readout systems have revealed admirable analytical performance with high specificity and sensitivity. The applicability of the assays was also established in complex biological samples (a cohort of cancer and MND patient samples) with high reproducibility. The analytical performance of all miRNA assays was validated using the standard RT-qPCR approach. I have faith in that our research efforts will lead to the design and development of a translational-focused point-of-care platform for clinically relevant autoantibody and miRNA analysis, which in sequence will hold the significant potential to improve the patient care in clinics as well as the outcomes may substantiate to be a venture of enormous commercial implications.

Declaration by author This thesis is composed of my original work and contains no material previously published or written by another person except where due reference has been made in the text. I have clearly stated the contribution by others to jointly-authored works that I have included in my thesis.

I have clearly stated the contribution of others to my thesis as a whole, including statistical assistance, survey design, data analysis, significant technical procedures, professional editorial advice, financial support and any other original research work used or reported in my thesis. The content of my thesis is the result of work I have carried out since the commencement of my higher degree by research candidature and does not include a substantial part of work that has been submitted to qualify for the award of any other degree or diploma in any university or other tertiary institution. I have clearly stated which parts of my thesis, if any, have been submitted to qualify for another award.

I acknowledge that an electronic copy of my thesis must be lodged with the University Library and, subject to the policy and procedures of The University of Queensland, the thesis be made available for research and study in accordance with the Copyright Act 1968 unless a period of embargo has been approved by the Dean of the Graduate School.

I acknowledge that copyright of all material contained in my thesis resides with the copyright holder(s) of that material. Where appropriate I have obtained copyright permission from the copyright holder to reproduce material in this thesis and have sought permission from co-authors for any jointly authored works included in the thesis.

Publications included in this thesis 1. Masud, M.K., Na, J., Younus M., Hossain, M.S.A., Bando Y., Shiddiky, M.J.A., and Yamauchi Y., Superparamagnetic Nanoarchitectures for Disease-Specific Biomarker Detection, Chemical Society Review 2019, 48, 5717-5751. – Incorporated in Chapter 2.1. Contributor Statement of contribution Mostafa Kamal Masud (Candidate) Conception and design (90%) Drafting (85%) Jongbeom Na Drafting (10%) Muhammad Younus Critically reviewing (10%) Md. Shahriar A. Hossain Critically reviewing (20%) Yoshio Bando Critically reviewing (20%) Muhammad J. A. Shiddiky Conception and design (5%) Critically reviewing (20%) Yusuke Yamauchi Conception and design (5%) Drafting (5%) Critically reviewing (30%)

2. Masud, M.K., Umer, M., Hossain, M.S.A., Yamauchi, Y., Nguyen, N.T. and Shiddiky, M.J.A., Nanoarchitecture Frameworks for Electrochemical miRNA Detection. Trends in Biochemical Sciences 2019, 44, 433-452. – Incorporated in Chapter 2.2. Contributor Statement of contribution Mostafa Kamal Masud (Candidate) Conception and design (90%) Drafting (85%) Muhammad Umer Drafting (10%) Critically reviewing (10%) Md. Shahriar A. Hossain Critically reviewing (20%) Yusuke Yamauchi Critically reviewing (20%) Nam-Trung Nguyen Critically reviewing (20%) Muhammad J. A. Shiddiky Conception and design (10%) Drafting (10%) Critically reviewing (30%)

3. Masud, M.K., Billah, M.M., Kim, J., Wood, K., Shiddiky, M.J., Nguyen, N.T., Parsapur, R.K., Selvam, P., Hossain, M.S.A., and Yamauchi, Y. Nanoarchitectured peroxidase-mimetic nanozymes: mesoporous nanocrystalline α- or γ-iron oxide?, Journal of Material Chemistry B 2019, 7, 5412-5422. – Incorporated in Chapter 3. Contributor Statement of contribution Mostafa Kamal Masud (Candidate) Conception and design (70%) Analysis and interpretation (70%) Drafting (80%) Jeonghun Kim Drafting (5%) Critically reviewing (5%) Md. Motasim Billah Drafting (5%) Kathleen Wood Analysis and interpretation (5%) Mohammad J. A. Shiddiky Critically reviewing (10%) Nam-Trung Nguyen Critically reviewing (10%) Rajesh Kumar Parsapur Analysis and interpretation (10%) Yusuf Valentino Kaneti Analysis and interpretation (5%) Critically reviewing (5%) Abdulmohsen Ali Alshehri Drafting (5%) Yousef Gamaan Alghamidi Critically reviewing (10%) Khalid Ahmed Alzahrani Critically reviewing (10%) Murugulla Adharvanachari Critically reviewing (10%) Parasuraman Selvam Conception and design (10%) Critically reviewing (10%) Md. Shahriar A. Hossain Analysis and interpretation (10%) Critically reviewing (10%) Yusuke Yamauchi Conception and design (20%) Drafting (5%) Critically reviewing (20%)

4. Masud, M.K., Islam, M.N., Haque, M.H., Tanaka, S., Gopalan, V., Alici, G., Nguyen, N.T., Lam, A.K.Y., Hossain, M.S., Yamauchi, Y. and Shiddiky, M.J.A., Gold-loaded nanoporous superparamagnetic nanocubes for catalytic signal amplification in detecting miRNA, Chemical Communications 2017, 53, 8231-8234. – Incorporated in Chapter 4. Contributor Statement of contribution Mostafa Kamal Masud (Candidate) Conception and design (70%) Analysis and interpretation (70%) Drafting (80%) Md. Nazmul Islam Drafting (5%) Analysis and interpretation (10%) Md. Hakimul Haque Drafting (5%) Analysis and interpretation (10%) Shunsuke Tanaka Drafting (5%) Vinod Gopalan Critically reviewing (10%) Gursel Alici Critically reviewing (10%) Nam-Trung Nguyen Critically reviewing (10%) Alfred K. Lam Critically reviewing (10%) Md. Shahriar A. Hossain Analysis and interpretation (10%) Critically reviewing (10%) Yusuke Yamauchi Conception and design (10%) Critically reviewing (20%) Muhammad J. A. Shiddiky Conception and design (20%) Drafting (5%) Critically reviewing (30%)

5. Masud, M.K., Yadav, S., Islam, M.N., Nguyen, N.T., Salomon, C., Kline, R., Alamri, H.R., Alothman, Z.A., Yamauchi, Y., Hossain, M.S.A. and Shiddiky, M.J.A., Gold- Loaded Nanoporous Ferric Oxide Nanocubes with Peroxidase-Mimicking Activity for Electrocatalytic and Colorimetric Detection of Autoantibody. Analytical Chemistry 2017, 89, 11005-11013. – Incorporated in Chapter 5. Contributor Statement of contribution Mostafa Kamal Masud (Candidate) Conception and design (70%) Analysis and interpretation (80%) Drafting (80%) Sharda Yadav Drafting (5%) Analysis and interpretation (10%) Md. Nazmul Islam Drafting (5%) Nam-Trung Nguyen Critically reviewing (10%) Carlos Salomon Analysis and interpretation (10%) Critically reviewing (10%) Richard Kline Critically reviewing (10%) Hatem R. Alamri Critically reviewing (10%) Zeid A. Alothman Critically reviewing (10%) Yusuke Yamauchi Conception and design (10%) Critically reviewing (10%) Md. Shahriar A. Hossain Drafting (5%) Critically reviewing (10%) Muhammad J. A. Shiddiky Conception and design (20%) Drafting (5%) Critically reviewing (30%)

Submitted manuscripts included in this thesis 1. Masud, M.K. Rabbee, G.M. Aziz, N.B., Stevens, C.H., Do-Ha, D. Yang, S. Blair, I.P. Hossain, M.S.A, Shim, Y-B, Ooi, L., Yamauchi, Y. and Shiddiky, M.J.A., Electrochemical detection of motor neuron disease (MND) derived exosomal miRNA at Attomolar Sensitivity, 2020. (Submitted in Integrative Biology) – Incorporated in Chapter 6. Contributor Statement of contribution Mostafa Kamal Masud (Candidate) Conception and design (70%) Analysis and interpretation (80%) Drafting (80%) Rabbee G. Mahmudunnabi Drafting (5%) Nahian Binte Aziz Drafting (5%) Claire H. Stevens Analysis and interpretation (10%) Dzung Do-Ha Critically reviewing (10%) Shu Yang Critically reviewing (10%) Ian P. Blair Critically reviewing (10%) Md. Shahriar A. Hossain Drafting (5%) Critically reviewing (10%) Yoon-Bo Shim Critically reviewing (10%) Lezanne Ooi Analysis and interpretation (10%) Critically reviewing (10%) Yusuke Yamauchi Conception and design (10%) Critically reviewing (10%) Muhammad J. A. Shiddiky Conception and design (20%) Drafting (5%) Critically reviewing (30%)

2. Masud, M.K., Na, J., Sina, A.A.I., Wood, K., Billah, Kani, K., Nguyen, N.-T., Shiddiky, M.J.A., Trau, M., Hossain, M.S.A., and Yamauchi, Y. Mesoporous Gold Biosensor for Electrochemical Detection of MicroRNA at Attomolar Level, 2020. (Submitted in ACS Applied Materials and Interfaces) – Incorporated in Chapter 7. Contributor Statement of contribution Mostafa Kamal Masud (Candidate) Conception and design (70%) Analysis and interpretation (80%) Drafting (80%) Jongbeom Na Drafting (5%) Analysis and interpretation (10%) Abu Ali Ibn Sina Critically reviewing (10%) Kathleen Wood Analysis and interpretation (10%) Mutasim Billah Drafting (5%) Critically reviewing (10%) Kenya Kani Drafting (5%) Nam-Trung Nguyen Critically reviewing (10%) Muhammad J. A. Shiddiky Conception and design (10%) Critically reviewing (10%) Matt Trau Critically reviewing (10%) Md. Shahriar A. Hossain Critically reviewing (20%) Yusuke Yamauchi Conception and design (20%) Drafting (5%) Critically reviewing (30%)

Other publications during candidature 1. Phan, H.P.§, Masud, M.K.§, Vadivelu, R.K., Dinh, T., Nguyen, T.K., Ngo, K., Dao, D.V., Shiddiky, M.J., Hossain, M.S.A., Yamauchi, Y., and Nguyen, N.T, Transparent crystalline cubic SiC-on-glass electrodes enable simultaneous electrochemistry and optical microscopy, Chemical Communication, 2019, 55, 7978-7981 (§equal contributor). 2. Hossain, T.,1 Mahmudunnabi, G.,1 Masud, M.K.,1 Islam, M.N., Ooi, L., Konstantinov, K., Al Hossain, M.S., Martinac, B., Alici, G., Nguyen, N.T. and Shiddiky, M.J., Electrochemical biosensing strategies for DNA methylation analysis, Biosensors and Bioelectronics, 2017, 94, 63-73. (1equal contributor). 3. Tanaka, S.,¥ Masud, M.K.,¥ Kaneti, Y.V., Shiddiky, M.J., Fatehmulla, A., Aldhafiri, A.M., Farooq, W.A., Bando, Y., Hossain, M.S.A. and Yamauchi, Y., Enhanced Peroxidase Mimetic Activity of Porous Iron Oxide Nanoflakes. ChemNanoMat. 2019, 5, 506-513. (¥equal contributor) 4. Yadav, S., Kashaninejad, Masud, M.K., N., Nguyen, N.T., Shiddiky, M.J.A., Autoantibodies as Diagnostic and Prognostic Cancer Biomarker: Detection Techniques and Approaches, Biosensor and Bioelectronics, 2019, 139, 111315. 5. Boriachek, K., Masud, M.K., Palma, C., Phan, H.P., Yamauchi, Y., Hossain, M.S.A., Nguyen, N.T., Salomon, C. and Shiddiky, M.J., Avoiding Pre-Isolation Step in Exosome Analysis: Direct Isolation and Sensitive Detection of Exosomes Using Gold-Loaded Nanoporous Ferric Oxide Nanozymes. Analytical chemistry. 2019, 89, 11005-11013 6. Azhar, A., Li, Y., Cai, Z., Zakaria, M.B., Masud, M.K., Hossain, M.S.A., Kim, J., Zhang, W., Na, J., Yamauchi, Y. and Hu, M., Nanoarchitectonics: A New Materials Horizon for Prussian Blue and Its Analogues. Bulletin of the Chemical Society of Japan, 2019, 92, 875-904. 7. Islam, M.N., Masud, M.K., Nguyen, N.T., Gopalan, V., Alamri, H.R., Alothman, Z.A., Al Hossain, M.S., Yamauchi, Y., Lam, A.K.Y. and Shiddiky, M.J., 2017. Gold-loaded nanoporous ferric oxide nanocubes for electrocatalytic detection of microRNA at attomolar level. Biosensors and Bioelectronics, 2018. 101, 275-281. 8. Bhattacharjee, R., Tanaka, S., Moriam, S., Masud, M.K., Lin, J., Alshehri, S.M., Ahamad, T., Salunkhe, R.R., Nguyen, N.T., Yamauchi, Y. and Hossain, M.S.A. Porous nanozymes: the peroxidase-mimetic activity of mesoporous iron oxide for the colorimetric and electrochemical detection of global DNA methylation, Journal of Material Chemistry B, 2018, 6, 4783-4791.

9. Gorgannezhad, L., Umer, M., Masud, M.K, Hossain, M.S.A., Tanaka, S., Yamauchi, Y., Salomon, C., Kline, R., Nguyen, N.T. and Shiddiky, M.J., Detection of FGFR2: FAM76A Fusion Gene in Circulating Tumor RNA Based on Catalytic Signal Amplification of Graphene Oxide‐loaded Magnetic Nanoparticles. Electroanalysis, 2018, 30, 2293–2301 10. Islam, M.N., Gorgannezhad, L., Masud, M.K., Tanaka, S., Hossain, M.S.A., Yamauchi, Y., Nguyen, N.T. and Shiddiky, M.J., 2018. Graphene‐Oxide‐Loaded Superparamagnetic Iron Oxide Nanoparticles for Ultrasensitive Electrocatalytic Detection of MicroRNA. ChemElectroChem, 2018, 5, 2488-2495. 11. Yadav, S., Masud, M.K., Islam, M.N., Gopalan, V., Lam, A.K., Tanaka, S., Nguyen, N.T., Hossain, M.S., Li, C., Yamauchi, Y. and Shiddiky, M.J., Gold-Loaded Nanoporous Iron Oxide Nanocubes: A Novel Dispersible Capture Agent for Tumor- Associated Autoantibodies Analysis in Serum, Nanoscale, 2017, 9, 8805-8814. 12. Islam, M.N., Masud, M.K., Haque, Hossain, M.S., Yamauchi, Y., Nguyen, N.T. and Shiddiky, M.J., RNA Biomarkers: Diagnostic and Prognostic Potentials and Recent Developments of Electrochemical Biosensors, Small Methods, 2017, 1, 1700131. 13. Haque, M.H., Gopalan, V., Islam, M.N., Masud, M.K., Bhattacharjee, R., Al Hossain, M.S., Nguyen, N.T., Lam, A.K. and Shiddiky, M.J., Quantification of gene-specific DNA methylation in oesophageal cancer via electrochemistry, Analytica Chimica Acta, 2017, 976, 84-93. 14. Islam, M.N., Gopalan, V., Haque, M.H., Masud, M.K., Al Hossain, M.S., Yamauchi, Y., Nguyen, N.T., Lam, A.K.Y. and Shiddiky, M.J., A PCR-free electrochemical method for messenger RNA detection in cancer tissue samples, Biosensors and Bioelectronics, 2017, 98, 227-333

Contributions by others to the thesis No contributions by others.

Statement of parts of the thesis submitted to qualify for the award of another degree No works submitted towards another degree have been included in this thesis.

Research Involving Human or Animal Subjects No animal or human subjects were involved in this research.

Acknowledgements The quest of my doctoral degree has been an exhilarating journey and more enjoyable due to the guidance, support and present of so many gracious and beautiful individuals. With my following few words, I would like to thank and convey my wholehearted gratitude and acknowledgement of your support, inspiration and influence in the completion of my dissertation. To the greatest extent, I wish to extend my heartfelt gratitude to my PhD supervisors Professor Yusuke Yamauchi and Dr Shahriar Hossain for their guidance, enormous support, enthusiasm, research support and environment-that smoothly drive my research. With the deepest gratefulness, I would like to thank Dr Muhammad Shiddiky (Collaborative supervisor, Griffith University-GU) for considering me and spotting my aptness for research from the beginning. It is a great bliss to work under you and learning so much, I am always grateful for what you have done to achieve this degree. I would like to thank Prof. Nam- Trung Nguyen (GU) for his scientific advice and research support at Queensland Micro- and Nanotechnology Centre (QMNC). I also wish to thank and acknowledge Dr Na, Motassim Billah and the entire Yamauchi group at AIBN, Dr Nazmul Islam (ICL, UK), Dr Hoang-Phuong Phan (GU), Dr. Shuvashis Dey (UQ), all members of Shiddkiky research group (GU) and the supportive UQCCR collaborators Dr Carlos Solomon. I am respectful to all of your optimistic encouragement and advice. I would like to extend my acknowledgement to Dr Katy Wood, Australia's Nuclear Science and Technology Organisation (ANSTO) for your support in neutron scattering and AINSE PGRA award. I will also like to extend my gratitude to my companions, friend and colleagues at the University of Queensland (UQ). I would like greatly acknowledge the AIBN for providing me with the resources looked- for my research along with helping me with countless administrative stuff. I would like to acknowledge the facilities and the scientific and technical assistance, of the Australian Microscopy & Microanalysis Research Facility at the Centre for Microscopy and Microanalysis and Queensland node of the Australian National Fabrication Facility, UQ. I would like to acknowledge and thank the government of the People’s Republic of Bangladesh for providing high-quality education from primary school to university. I am also very much obligated to my amazingly supportive and loving family. Especially, my parents, brother, sisters, parents-in-law are the leading blessings of my life. Last but not least, my inestimable gratefulness and love for the two most invaluable assets of life, my wife Sajiya Akther and my daughter Marnia Kamal Tehzeeb, whose sacrifice, unrivalled love and encouragement have made me come to this far.

Financial support This research was supported by an Australian Government Research Training Program Scholarship. I would like to thank AINSE Limited for providing financial assistance (top-up) through post-graduate research award (PGRA) and support to conduct the experiment at ANSTO.

Keywords autoantibody, biosensor, electrocatalytic activity, electrochemical detection, mesoporous gold electrode, mesoporous iron oxide, nanoarchitecture, disease diagnostic, nanozyme, microRNA

Australian and New Zealand Standard Research Classifications (ANZSRC) ANZSRC code: 030102 Electroanalytical Chemistry, 30% ANZSRC code: 100402 Medical Biotechnology Diagnostics (incl. Biosensors), 40% ANZSRC code: 100703 Nanobiotechnology, 30%

Fields of Research (FoR) Classification FoR code: 0301 Analytical Chemistry, 30% FoR code: 1004 Medical Biotechnology, 40% FoR code: 1007 Nanotechnology, 30%

Table of Contents 1. General Introduction 2 1.1 Background 2 1.2 Aims 6 1.3 Significance of the project 7 1.4 Structure of the thesis 10 1.5 References 13 2. Literature Review: Nano-architectures for disease-specific biomarker detection 17 2.1. Superparamagnetic nanoarchitectures 17 2.1.1. Synthesis of superparamagnetic nanoparticles 21 2.1.1.1. General synthesis 21 2.1.1.2. Template-based synthesis 23 2.1.1.3. Multicomponent (hybrid) magnetic nanoparticle synthesis 27 2.1.2. Surface functionalization of superparamagnetic nanoparticles 29 2.1.2.1. Small organic molecules, functional groups, and surfactants 29 2.1.2.2. Polymers 31 2.1.2.3. Bioactive molecules 32 2.1.2.4. Inorganic materials 33 2.1.3. The function of superparamagnetic nanoparticles in biosensor development 36 2.1.3.1. Magnetic capture and separations 36 2.1.3.2. Development of a detection platform (sensor and biosensor) 39 2.1.3.2.1. Immobilization of biomolecules 41 2.1.3.2.2. Electrocatalytic signal amplification 43 2.1.3.2.3. Signal-generating probes 45 2.1.3.3. Nanocarriers 48 2.1.3.4. Natural enzyme mimetics: Nanozymes 49 2.1.4. Application of superparamagnetic nanoparticles in biomolecular detection 52 2.1.4.1. Electrochemical biosensors 52 2.1.4.1.1. Nucleic acid assay 52 2.1.4.1.2. Immuno assay 56 2.1.4.1.3. Cytosensors 56 2.1.4.2. Optical biosensors 59 2.1.4.2.1. Colorimetry and fluorescence detection 59 2.1.4.2.2. SPR biosensors 60

2.1.4.2.3. LSPR biosensor 62 2.1.4.2.4. SERS biosensors 63 2.1.4.3. Biosensors based on nanozymes 65 2.1.5. Point-of-care testing: The impact of superparamagnetic particles 72 2.1.6. Future perspectives and conclusion 77 2.1.7. References 78 2.2. Nanoarchitecture frameworks for electrochemical miRNA detection 112 2.2.1. Challenges and considerations 114 2.2.2. Nanoarchitectures for miRNA detection 117 2.2.3. Nanoarchitecture based electrochemical miRNA detection 122 2.2.4. Concluding remarks and future perspectives 136 2.2.5. References 137 3. Nanoarchitectured peroxidase-mimetic nanozymes: mesoporous nanocrystalline α- or γ-iron oxide? 150 3.1. Introduction 150 3.2. Experimental 152 3.2.1. Materials 152 3.2.2. Characterization 152 3.2.3. Synthesis of CMK-3 153

3.2.4. Synthesis of α-Fe2O3 and γ-Fe2O3 153 3.2.5. Nanozyme activity 153 3.2.6. Investigation of nanozyme activity through scavenging hydroxyl radical 154 3.2.7. Naked-eye observation and colourimetric estimation of Glucose 154 3.3. Result and discussion 155 3.3.1. Structural Characterization of the Nanozymes 155 3.3.2. Nanozyme activity 160 3.3.3. Detection and estimation of glucose 169 3.4. Conclusion 173 3.5. References 173 4. Gold-Loaded Nanoporous Ferric Oxide Nanocubes with Peroxidase-Mimicking Activity for Electrocatalytic and Colorimetric Detection of Autoantibody 180 4.1. Introduction 180 4.2. Experimental section 182 4.2.1. Materials and instrumentations 182

4.2.2. Study group and ovarian cancer samples 183 4.2.3. Determination of the surface area of the electrode 183

4.2.4. Preparation of IgG/Au-NPFe2O3NC nanocatalyst 184

4.2.5. The peroxidase-mimetic activity of Au-NPFe2O3NC 184 4.2.6. Colorimetric and amperometric detection of serum p53 autoantibody 185 4.3. Result and discussion 186

4.3.1. The peroxidase-mimetic activity of Au-NPFe2O3NC 186

4.3.2. Steady-State Kinetic for Au-NPFe2O3NC 190 4.3.3. Colourimetric and amperometric detection of autoantibody 194 4.4. Conclusions 199 4.5. References 200 5. Gold-loaded nanoporous superparamagnetic nanocubes for catalytic signal amplification in detecting miRNA 207 5.1. Introduction 207 5.2. Experimental 208 5.2.1. Materials and instrumentations 208 5.2.2. Electrochemical measurement of catalytic activity 210 5.2.3. Determination of the surface area of the electrodes 210 5.2.4. Preparation of the miRNA recognition interface 210 5.2.5. RNA Extraction from cell lines and tissue samples 211 5.2.6. Isolation of target miRNA 211 5.2.7. Electrochemical detection of adsorbed microRNA 212 5.2.8. Quantitative reverse-transcription polymerase chain reaction 213 5.3. Result and discussion 214

5.3.1. Synthesis and Characterization of Au@NPFe2O3NC 214

5.3.2 Electrocatalytic Activity of Au@NPFe2O3NC 216 5.3.3. microRNA detection 221 5.4. Conclusion 228 5.5. References 228 6. Electrochemical Detection of Motor Neuron Disease (MND) Derived Exosomal miRNA at Attomolar Sensitivity 234 6.1. Introduction 234 6.2 Experimental 237 6.2.1. Materials and Instrumentation 237

6.2.2. Electrocatalytic activity 238 6.2.3. Cell culture and cell culture media collection 238 6.2.4. Exosome isolation 239 6.2.5. Isolation of exosomal (total) small RNA 239 6.2.6. Probe hybridization and magnetic isolation of target miRNA (miR-338-3p) 240 6.2.7. Electrochemical detection of adsorbed microRNA 240 6.2.8. Statistical analysis 241 6.2.9. qRT-PCR validation 241 6.3. Result and discussion 242 6.3.1. Electrocatalytic activity of nanocubes towards Methylene Blue (MB) 242 6.3.2. Electrocatalytic detection of miRNA 245 6.3.3. Assay Specificity 247 6.3.4. Assay sensitivity 248 6.3.5. Analysis of miR-338-3p in exosomes from ALS patient 250 6.4. Conclusions 252 6.5. References 253 7. Mesoporous Gold Biosensor for Electrochemical Detection of MicroRNA at Attomolar Level 259 7.1 Introduction 259 7.2. Experimental 256 7.2.1. Reagents and materials 262 7.2.2 Instrumentations 262 7.2.3 Synthesis and characterisation of the mesoporous gold electrode (MPGE) 263 7.2.4. Calculation of electrochemical surface area (ECSA) and roughness factor 268 7.2.5. Hybridization and magnetic isolation of target miRNA (miR-9-2) 268 7.2.6. Electrochemical detection of adsorbed target miRNA 269 7.3. Results and discussion 269 7.4. Conclusion 282 7.5. References 282 8. Conclusions and Future Recommendations 289 8.1 Conclusions 289 8.2 Future recommendations 291

List of Figures Figure 2.1.1: Schematic representation of the building blocks of nanoengineered 20 superparamagnetic nanoparticles, their surface functionalization, and their functions for integration into biomolecular biosensors. Figure 2.1.2: Schematic illustration of the fate of PB nanoparticles at different 26 applied calcination temperatures. Figure 2.1.3: Schematic representation of the conversions of selected multilayer 28

FexOy@Au composites. Figure 2.1.4: Schematic representation of the functionalization of magnetic 31 nanoparticles. Figure 2.1.5: Schematic representation of the thiolate–oleylamine exchange 35 reaction followed by cross-linking via different linkers (A and B)

for the assembly of Fe3O4@Au NPs. Figure 2.1.6: Schematic representation of superparamagnetic NP–based 37 bioseparation of biomolecules. Figure 2.1.7: The function of superparamagnetic nanomaterials in a biomolecule 39 detection platform Figure 2.1.8: General strategies for the immobilization of MNPs with 42 biomolecules. Figure 2.1.9: Schematic representation of an SPR immunosensor based on 44 clustered magnetic NMs for signal amplification. Figure 2.1.10: Schematic representation of a biobarcode-functionalized MNP label– 46 based chemiluminescence detection of DNA hybridization (A); the flow injection chemiluminescence readout system for the quantification of Fe3+ (B). Figure 2.1.11: Magneto-controlled reversible translocation of the functionalized 47 MNPs between the organic phase above the aqueous electrolyte and the electrode surface. (A) A magnet below the electrode surface pulls the hydrophobic MNPs, forming a membrane-like layer on the electrode surface (“off” state); (B) a magnet positioned above the electrode returns the MNPs to the organic phase, resulting in the generation of an electrochemical reaction (“on” state).

Figure 2.1.12: Schematic representation of the detection of p53 autoantibodies 49

based on a superparamagnetic Au@NPFe2O3 nanocube as a dispersible nanocarrier.

Figure 2.1.13: A magnetic Fe3O4-based sandwich-type detection strategy for two- 54 base mismatched DNA detection. Figure 2.1.14: Superparamagnetic nanoparticle-based miRNA sensor. 55

Figure 2.1.15: Schematic illustration of (A) the fabrication of Fe3O4@Ag-Pd 58 hybrid NPs and (B) the steps involved in sensor design for the detection of CTCs.

Figure 2.1.16: Schematic representation of a GO-sensing film and Fe3O4-HGNPS- 62 hybrid probe-based magnetic field-assisted SPR biosensor for IgG detection. Figure 2.1.17: Schematic illustration of a magnetic SERS immunosensor for sensitive 64 detection of avian influenza virus. Figure 2.1.18: Schematic representation of the assay for p53 autoantibody 66 detection using the peroxidase mimetics of gold-loaded nanoporous ferric oxide nanocubes. Figure 2.1.19: Schematic demonstration for the detection of human hCG using 73 magnetic nanozyme–based LFIA strips. Figure 2.1.20: Schematic representation of the detection of Ebola, Lassa, and 74 malaria using magnetic particles and SERS nanotags. Figure 2.2.1: Commonly used nanoarchitecture in miRNA biosensing. 119 Figure 2.2.2: Nanoarchitecture in miRNA biosensing. 123 Figure 2.2.3: Nanoarchitecture based electrochemical miRNA biosensing. 126 Figure 2.2.4: Triple signal amplification strategy for ultrasensitive determination 134 of miRNAs based on duplex-specific nuclease (DSN) and bridge DNA−Gold NPs.

Figure 2.2.5: Schematic representation of Au-Fe2O3NC catalyzed miR-107 assay 136 3+ 3 using [Ru(NH3)6] /[Fe(CN)6] .

Figure 3.1: (a) Low- and wide-angle XRD patterns of mesoporous γ-Fe2O3 and 156

α-Fe2O3. (b) N2 adsorption-desorption isotherms and pore size

distribution curves of mesoporous γ-Fe2O3 and α-Fe2O3 (c) TEM

image with ED patterns and (d) high-resolution TEM image of

mesoporous γ-Fe2O3.

Figure 3.2: (a, b) SEM image of (a) mesoporous γ-Fe2O3 and (b) mesoporous 157

α-Fe2O3. (c and d) Enlarged TEM images of mesoporous γ-Fe2O3.

Figure 3.3: Rietveld refined XRD patterns of (a) mesoporous γ-Fe2O3 and (b) 158

mesoporous α-Fe2O3. The calculated lattice parameters are a, b, c =

8.374(7) Å (for mesoporous γ-Fe2O3) and a, b = 5.0327(9) Å; c =

13.761(3) Å (for mesoporous α-Fe2O3), respectively.

Figure 3.4: M-H curves of (a1) of mesoporous γ -Fe2O3, (a2) bulk-γ-Fe2O3, (b1) 159

mesoporous α-Fe2O3, and (b2) bulk-α-Fe2O3.

Figure 3.5: M-T curves of (a) mesoporous γ-Fe2O3 and (b) bulk γ-Fe2O3. 160 Figure 3.6: (a) Schematic illustration of the nanozyme activity of mesoporous 161

IO for the oxidation of TMB in the presence of H2O2. (b) The peroxidase-like activities of the two mesoporous IO samples in comparison with the two control samples

Figure 3.7: The nanozyme activity of non-porous and mesoporous γ-Fe2O3 163 towards the oxidation of TMB. Figure 3.8: The absorbance spectra obtained for the oxidation of ABTS by 163 peroxidase mimetic activity of iron oxide nanozymes. Figure 3.9: The responses of absorbance change (at 652 nm) after the addition 164 of hydroxide radical scavengers. Figure 3.10: Investigating hydroxyl ion (·OH) radical formation through the 165 fluorescence spectra of 2-hydroxyterephthalic acid produced from the oxidation of terephthalic acid (TA) by ·OH in the presence of

both (a) mesoporous γ-Fe2O3 and (b) mesoporous α-Fe2O3. Figure 3.11: The pH-dependent peroxidase-like activities of (a) mesoporous γ- 166

Fe2O3 and (b) α-Fe2O3 samples (5 µg).

Figure 3.12: The peroxidase-like activities of (a) mesoporous γ-Fe2O3 and (b) 166

mesoporous α-Fe2O3 samples with different loading amounts of the samples. Figure 3.13: Steady-state kinetics and catalytic mechanism of mesoporous 169

Fe2O3.

Figure 3.14: (a) Schematic representation of nanozyme-based glucose detection. 171 (b)The mean responses of absorbance obtained for the assay with

mesoporous α-Fe2O3 versus γ-Fe2O3 and a control sample. (c) The absorbance spectra obtained for the designated concentration of standard glucose samples ranging from 1 µM to 1000 µM. (d) The corresponding bar diagram for standard glucose concentration. (e)

Calibration curve for glucose detection using γ-Fe2O3 as nanozyme with the range of glucose concentration from 1 µM to 1000 µM Figure 4.1: Bright-field TEM image and HAADF-STEM image for gold-loaded 186

nanoporous ferric oxide nanocubes (Au-NPFe2O3NC).

Figure 4.2: (a) SEM image and (b-d) elemental mapping for Au-NPFe2O3NC 187

Figure 4.3: XRD pattern for Au-NPFe2O3NC. 187 Figure 4.4: (a) Schematic illustration of peroxidase-mimicking activity of Au- 188

NPFe2O3NC for the oxidation of TMB in the presence of H2O2. Mean values of (b) absorbance (UV-vis) and (c) amperometric current signals for the negative and positive control samples (inset in (b) and (c) are the corresponding photos for the naked eye evaluation and i-t curves respectively). Figure 4.5: Optimization of the pH and nanocube concentration for the 190

peroxidase-like activity of Au-NPFe2O3NC nanocubes. Mean values of absorbance (a1, b1) and amperometric current density (a2 and b2) at designated pH (a1 and a2) of NaAc buffer solution and amount of nanocubes (b1 and b2). Figure 4.6: Dependence of the UV-vis absorbance (at 652 nm) of 191

Au@NPFe2O3NC catalysed TMB/H2O2 reaction on the

concentration of (a) H2O2, (b) TMB in the range from 0.01 to 1.1 M and 0.01 to 1.0 mM respectively. Figure 4.7: Steady-state kinetic analyses using Michaelis-Menten model (main 192 panel) and Lineweaver-Burk model (inset panel) for the Au-

NPFe2O3NC nanocubes by varying concentration of (a) H2O2 (0.01 to 1.1 M) and (b) TMB (0.01 to 1.0 mM) with fixed amount of (a)

TMB (800 µM) and (b) H2O2 (700 mM).

Figure 4.8: Schematic representation of the assay for the detection of tumor- 194 associated plasma (and serum) p53 auntoantibody. Figure 4.9: Mean responses of (a) absorbance and (b) steady-state 196 amperometric current obtained for the assay with one positive (presence of p53 autoantibodies in serum) with three negative control samples. Figure 4.10: Concentration dependent curve for p53 autoantibody standards 197 provided in the commercial p53 autoantibody ELISA kit. Mean responses of (a) absorbance and (b) steady-state amperometric currents obtained for the designated concentration of standard samples. Figure 4.11: Mean responses of (a) absorbance and (b) steady-state 198 amperometric current obtained p53-specific autoantibodies present in plasma samples obtained from patients with epithelial ovarian cancer high-grade serous subtype and their non-cancerous healthy patients. Figure 5.1: (A-B) SEM images of (A) PB nanocubes and (B) the calcined PB 215 nanocubes. (C-D) Wide-angle XRD patterns of (C) PB nanocubes and (D) Au-loaded nanoporous iron oxide nanocubes. Figure 5.2: (A) Elemental mapping images (O, Fe, and Au), and (B) EDX 216

spectrum of Au@NPFe2O3NC.

Figure 5.3: Magnetization curve measured at 300 K for Au@NPFe2O3NC 216 Figure 5.4: (A) CVs obtained at an unmodified GCE (top) and 217

Au@NPFe2O3NC-modified GCE (bottom) in 50µM RuHex (scan rate, 50 mVs-1). (B) Comparison of these two CVs. Figure 5.5: Cyclic voltammograms at GCE/Bare (left) and 217

GCE/Au@NPFe2O3NC (right) at designated pH from 3 to 11 in the presence of 50 µM RuHex (0.01M PBS, pH-7, scan rate 50 mVs-1). o o Figure 5.6: Cathodic peak currents of GCE/Au@NPFe2O3NC at 25 C, 37 C 218 and 50 oC in the presence of 50 µM RuHex (0.01 M PBS, pH-7, scan rate = 50 mVs-1).

Figure 5.7: (A) Cyclic voltammograms obtained at GCE/Bare (top, left) and 219

GCE/Au@NPFe2O3NC (top, right) electrodes at different scan rate (50µM RuHex, 0.01M PBS, pH 7.0). (B) Corresponding curves for 1/2 ipc and ipa (current density) as a function of ν .

Figure 5.8: Cyclic voltammograms of GCE/Au@NPFe2O3NC upon successive 220 addition of RuHex (a-0, b-25, c-50, d-100, and e-200 µM) to the 0.01M PBS (pH-7, scan rate = 50mVs-1).

Figure 5.9: (A) Amperometric responses of GCE/Au@NPFe2O3NC with the 220 successive addition of RuHex solution (10 to 1100µM) into the 0.01M PBS (pH-7); (B) the corresponding calibration plot.

Figure 5.10: Cathodic peak currents obtained by bare GCE, Au@NPFe2O3NC- 221

modified-GCE and NPFe2O3NC-modified GCE at room temperature, in presence of 50μM RuHex (0.01M PBS, pH-7, scan rate = 50mVs-1). Inset, corresponding cyclic voltammogram. Figure 5.11: Assay principle. miRNA was extracted from target cell lines or 222 tissue samples. After magnetic isolation and purification, the target

miR-21 was adsorbed onto the Au@NPFe2O3NC attached SPCE. An enhance electrochemical signals wer generated by the CC 3+/2+ interrogation of miRNA-bound Ru(NH3)6] complexes. Figure 5.12: (A) Charge density for the SPCE/Bare, SPCE/miR-21, 223

SPCE/NC/miR-21 (Qdl), SPCE/NC/buffer (NoT), SPCE/NC/miR- 107 (Wrong target) and SPCE/NC/miR-21 (Q) electrodes. Concentration of miR-21 and miR-107 were 10 nM. (B) Typical CC curves for the (c-j) 100 fM-1.0 μM of synthetic miRNA. Curves a,

and b are for the Qdl and NOT respectively. (C) Charge density- concentration profiles. (D) Charge density obtained for eight tissue samples derived from ESCC patients. The concentration of RuHex is 50 µM. Figure 5.13: CC charge generated by the extracted miR-21 from two HKESC-1 225 and HKESC-4 cell lines. Figure 5.14: RT-qPCR validation of miR-21 expression levels in the (A) two 225 ESCC cell lines (B) four tumor tissue samples obtained from the patients with ESCC.

Figure 6.1: Electrocatalytic activity of Au-Fe2O3NC in MB. Comparison of the 243 (a) CVs and (b) DPV obtained at an unmodified GCE and AuNP- -1 Fe2O3NC-modified GCE in 50 µM MB (scan rate, 50 mVs ). (c)

CV obtained at GCE/AuNP-Fe2O3NC-electrodes at different scan rate (50μM MB, 0.01M PBS, pH 7.0); (d) corresponding curves for 1/2 ipc and ipa (current density) as a function of ν .

Figure 6.2: (a) Represents the modification of a GCE by AuNPs-Fe2O3NC for 244 interrogating catalytic activity of NCs towards the redox process of MB; (b) and (c) depicts the calibration curve of the square root of potential (scan rate) vs. current density obtained from scan rate study of bare GCE towards reduction and oxidation process of MB respectively. The CV curve obtained from the interrogation of different scan rate ranging from 10 to 1500 mVs-1 using a bare GCE is figured in d. Figure e and f represents the calibration curve of the square root of potential (scan rate) vs. current density obtained from

the scan rate study of AuNPs-Fe2O3NC-modified GCE towards reduction and oxidation process of MB respectively.

Figure 6.3: (a) Amperometric responses of AuNP-Fe2O3NC-modified GCE 245 with the successive addition of MB solution (0 to 200 µM) into 0.01 M PBS (pH-7); (b) the corresponding calibration plot. Figure 6.4: Schematic representation of the assay. Exosomal RNA was isolated 246 from exosome that derived from preconditioned media of motor neurons (ALS). Target miRNA was hybridized with capture probe, and biotinylated magnetic beads were then added to the hybrid for magnetic isolation and purification. After magnetic purification, target miRNA were released from magnetic beads and allowed to

adsorb onto the gold NPs of AuNP-Fe2O3NC/SPCE surface for electrochemical detection. The presence of adsorbed miRNA interact with positively charged MB redox molecules, and the amount of charge intercalation is measured by CC readout in the 3- presence of 4 mM [Fe(CN)6] in 40mM Tris-HCl buffer (pH 7.4). Figure 6.5: Assay specificity; CC (inset) showing the charge density obtained 248 after assay was performed on different target sequences

(SPCE/miR-338-3p, SPCE/NC/ miR-338-3p (control), SPCE/NC , SPCE/NC/miR-338-3p (wrong target), SPCE/NC/miR-338-3p/MB in 40mM Tris-HCl buffer (pH 7.4) and SPCE/NC/miR-338-3p/MB 3- in 4mM [Fe(CN)6] in 40mM Tris-HCl buffer (pH 7.4) for redox cycle. Concentration of miR-338-3p and miR-338-3p were 10 pM

(NC: AuNP-Fe2O3NC). Figure 6.6: Improved assay sensitivity; CC curves showing amount of charge 249 generated to different concentration of starting miR-338-3p targets (a) before (a; a-control; b to g-10 fM to 1 nM) and (b) after (b; a- 3- control; b to I -100 aM to 1 nM) coupling with [Fe(CN)6] (redox cycle). The figure c and d show the analogous bar diagrams (inset; 3- linear calibration plots) before (and after coupling with [Fe(CN)6] respectively. Figure 6.7: Assay performance on clinical samples; (a) Electrochemical signal 251 obtained from sample C1, C2, and P1 to P3 in week 2 and week 4 intervals, where C1 and C2 represent the motor neuron samples from healthy patients and P1, P2 and P3 represent motor neuron samples from ALS patients. (b) RT-qPCR validation of exosomal miR-338-3p expression levels in the two healthy control samples and three ALS samples over the two and four weeks of motor neuron differentiation. Below: Corresponding p values obtained using two-way ANOVA comparing the control (healthy) samples and ALS patient’s samples. Figure 7.1: (a) Schematic representation of the preparation of mesoporous Au 261 electrode (MPGE) via electrodeposition of gold (III)-containing polymeric (block) micelles. The polymeric micelles are formed by dissolving polystyrene-b-poly(ethylene oxide)) (PS-b-PEO) diblock copolymer in THF followed by the addition of ethanol and aqueous gold (III) chloride solution; and. (b) Schematic representation of electrochemical detection of miRNA. Figure 7.2: FT-IR spectra during PS-b-PEO micellization with Au. 264 Figure 7.3: Small-angle neutron scattering (SANS) patterns of two types of 265 polymeric micelle solutions with corresponding debye fits. Sample

1 (a) was prepared by mixing PS-b-PEO in THF followed by the addition of ethanol; sample (b) was obtained by mixing PS-b-PEO

in THF followed by the addition of ethanol and HAuCl4 solution. Figure 7.4: Amperometric (i-t) plots for deposition of MPGE under a typical 266

electrolyte containing PS18000-b-PEO7500 micelles, THF, ethanol and gold (III) solution. Figure 7.5: FT-IR spectra before and after removal of PS-b-PEO template 266 during the preparation of MPGE. Figure 7.6: The SEM image of the top-surface of MPGE 267 Figure 7.7: The high-resolution XPS spectra of Au-4f. 268 Figure 7.8: Magnetic isolation and purification of target miRNA using 269 biotinylated capture probes and streptavidin-coated magnetic beads. Figure 7.9: Electrodeposition time versus ECSAs. (a) Cyclic voltammograms 270

(CVs) in 0.5 M H2SO4 of MPGE with different film thicknesses obtained by conducting electrodeposition in different timescale ranging from 0 s to 2000 s ; (b) the linear relationship between the ECSAs and the deposition time (film thickness), and (c) the roughness factor in response to deposition time. 4-/3- Figure 7.10: Electrocatalytic activity of MPGE in [Fe(CN)6] redox system. 272 (a) CV curves of SGE and MPGE and (b) DPV responses of SGE 3-/4- and MPGE in 2 mM [Fe(CN)6] solution. (c) CV curves at 3-/4- different concentrations of [Fe(CN)6] on MPGE and (d) bar- diagram of the corresponding currents (cathodic). (e) CV curves at different scan rates from 5 mVs-1 to 1000 mVs-1 on MPGE in 2 mM 3-/4- [Fe(CN)6] solution and (f) summary of the corresponding current responses upon the square-root of scan rates. (g) Amperometric 3-/4- responses on MPGE with the successive addition of Fe[(CN)6] solution into the 0.01 M PBS (pH-7) and (h) the corresponding calibration plot; inset- Lineweaver-Burk Model. 3-/4- Figure 7.11: (a) The CV’s in 2 mM [Fe(CN)6] for MPGEs prepared at 273 different deposition time (thickness); (b) the corresponding cathodic current, which follows a polynomial relationship with the increasing film thickness.

Figure 7.12: (a) The DPV responses of MPGE with increasing film thickness; 273 (b) the corresponding current density. The current follows the polynomial trend with the increasing deposition time. Figure 7.13: (a) The representation of both the current density (black line) and 274 ECSA’s (red line) obtained from different electrodeposited MPGE, where the film thickness (deposition) versus ECSA follows linear relationship and the film thickness (deposition) versus current density follows polynomial relationship (a characteristic of enzymatic electrocatalysis). 3-/4- Figure 7.14: Stability of MPGE over several CV curves in 2 mM [Fe(CN)6] 275 solution Figure 7.15: DPV responses after the adsorption of same quantity of miR-9-2 (10 276 pM) at MPGE with different deposition time (0 – 2000 s). Figure 7.16: (a,b,) DPV responses before and after 10 pM of miRNA adsorbed 278 on SGE and MPGE, respectively. (c) Sensitivity Comparison; both SGE and MPGE generates significantly higher amount of current in response to NoT (PBS is used instead of miRNA), and surprisingly MPGE generate 8-times higher current than that of SGE (0.384 vs. 0.061 mAcm-2) (d) Assay specificity where MPGE with the target miRNA generate pointedly higher responses from Bare (MPGE; without miRNA), NoT (PBS), wrong target (miR-338-3p), respectively.

Figure 7.17: (a) SEM and (b) AFM topography image miRNA-adsorbed MPGE, 279 respectively. (c) Assay sensitivity; the DPV responses with the different concentration of synthetic target miR-9-2 ranging from 1 fM to 1 nM. (d) The corresponding bar diagram of assay sensitivity.

List of Tables

Table 2.1: Magnetic nanoparticles used for magnetic isolation of target 40 biomolecules. Table 2.2: Engineered MNPs for disease-specific biomolecular (electrochemical 68 and optical) sensing Table 2.3: Magnetic nanostructure–based POC diagnostic system for rapid 75 analysis of disease biomarkers Table 2.4: Electrochemical miRNA detection based on nanoarchitectures. 127 Table 3.1: Structural information of SBA-15, CMK-3, and the resulting 155

mesoporous γ-Fe2O3 and α-Fe2O3. Table 3.2: Comparison of the peroxidase-mimicking activities (kinetic 172

parameters and conditions) of the as-prepared mesoporous α-Fe2O3

and γ-Fe2O3 with recently reported iron oxide-based nanostructures

and composites for TMB/H2O2 substrate.

Table 4.1: The comparison of the kinetic parameters of Au-NPFe2O3NC and 193 reported nanomaterials. Table 5.1: Oligonucleotide sequences (miR-21) 209 Table 5.2: Comparative Analytical performance of electrochemical sensors for 227 microRNA Table 6.1: Oligonucleotide sequences (miR-338-3p) 237 Table 7.1: Oligonucleotide sequences (miR-9-2) 262 Table 7.2. Comparison of nanostructure-based approaches of electrochemical 281 detection of miRNA

List of Abbreviation used in this thesis

µM Micro molar ABTS 2,2'-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid AFM Atomic Force Microscopy AIBN Australian Institute for Bioengineering and Nanotechnology AINSE Australian Institute of Nuclear Science and Engineering ALS Amyotrophic lateral sclerosis aM Attomolar BET Brunauer-Emmett-Teller CC Chronocoulometry CV Cyclic Voltammetry DPV Differential Pulse Voltammetry EC Electrochemical ESCC Esophageal squamous cell carcinoma (ESCC) fM Femtomolar FTIR Fourier-transform infrared spectroscopy GCE Glassy Carbon Electrode Km The Michaelis–Menten constant LSPR Localized surface plasmon resonance MB Methylene Blue mg Milligram miRNA MicroRNA mL Millilitre mM Millimolar MND Motor Neuron Disease MNPs Magnetic Nanoparticles MPGE Mesoporous Gold Electrode NC Nanocube nM Nanomolar NPs Nanopartilces PB Prussian Blue PBS Phosphate Buffer Saline pM Picomolar

RT-qPCR Quantitative reverse transcription PCR RuHex Ruthenium Hexaamine SEM Scanning Electron Microscope SERS Surface-enhanced Raman spectroscopy SGE Sputtered Gold Electrode SMNPs Superparamagnetic Nanoparticles SPCE Screen-printed Carbon Electrode SPR Surface Plasmon Resonance SSC Saline-sodium Citrate TEM Transmission Electron Microscopy TMB 3,3',5,5'-Tetramethylbenzidine UV-Vis Ultraviolet–visible spectroscopy

Vmax Maximum reaction rate XPS X-ray photoelectron spectroscopy XRD X-ray Powder Diffraction

Chapter 1

General introduction

1. General Introduction

1.1. Background

The advancements made in detection technology and medicine in the last decades are impressive. However, diseases not detected on time or not properly monitored are currently the main causes of death. Cancer is the foremost cause of illness in every country of the world and has a substantial health, social and economic impact. According to the American Institute for Cancer Research, the age-standardised rate for all cancers worldwide is 197.9 per 100,000 (2018), with Australia entailing the highest rate of 468 per 100,000. Moreover, it has estimated that there will be 21.7 million new cases of cancer and 13 million cancer-related deaths by 2030.1 For instance, in Australia during 2017, there were approximately 134,174 new cases of a cancer diagnosis with 47,753 deaths.2 Despite progress in treatment and detection technology, the burden associated with cancers is still high, as cancers are often only diagnosed at the most advanced stage (III or IV), especially in developing countries. Two main probable reasons can be attributed to the lack of efficient, easily accessible and precise diagnostic technologies and on-time detection as soon as symptoms are substantial. If detected on time, they can be prevented, and the burden will be reduced significantly. Moreover, having confidence in that early-stage detection of cancer increases the survival rate and reduces cancer-associated morbidity and mortality. For example, chemotherapy treatment at early stages of ovarian cancer leads to an 80% reduction in progression to an aggressive phenotype.3 Therefore, the development of point-of-care diagnostic technologies for detecting illness (cancer) in the earlier stage through the detection of clinically relevant disease biomarkers could provide a compelling resolution for patient treatment and monitoring and may reduce the disease- associated burdens meaningfully. Autoantibodies are predominantly produced in response to body’s antigen (self- antigen) by a small subset of the B cells known as B-1 cells or CD5+ B cells that may encompass nucleic acids (DNA, RNA), proteins, carbohydrates, lipids or several combinations of these biological constituents.4,5 They may be pathogenic, diagnostics and disease-specific, or of no ostensible significance. However, there is growing evidence that these autoantibodies are also tangled in chronic malignancies. Several mechanisms have already been proposed to explain for the production of autoantibodies in cancer comprising host-immune reactions to tumour-associated antigens (TAAs), antigenic stimulation as a consequence of the destruction of malignant cells, or immune dysregulation persuaded by the neoplastic process.6 They are formed by the patient’s immune system long before (quite a few months or years) the onset of

the disease symptoms. Moreover, they are relatively more stable (elongated half-lives owing to low proteolysis and clearance) and expressed at an increased level in very early stages of cancer and are witnessed in the patients with quite a lot of carcinomas including breast,7 lung,8 ovarian9 cancer, thereby presenting themselves as a potential early reporter (biomarker) of the atypical cellular mechanism involved in tumorigenesis. To date, numerous conventional methods such as enzyme-linked immunosorbent assay (ELISA) or sodium dodecyl sulphate- polyacrylamide gel electrophoresis (SDS/PAGE) have used to detect the autoantibodies in serum.4,10 The widely used method is ELISA, where HRP (horse-radish peroxidase)- conjugated protein-specific secondary antibodies are used to form sandwich immunoassay with the target and primary antibody to read the target autoantibody. However, these methods suffering from the expensive antibody, low sensitivity and qualitative or semiquantitative readout. Several advanced methods have also reported by integrating electrochemical readout; nonetheless, they still rely on HRP-based enzymatic reaction.11,12 With the revolutionary advancement in the next-generation sequencing technologies, the human transcriptome has been repeatedly interrogated, mainly due to the emerging functional and regulatory roles of a large fraction of coding and non-coding RNAs such as messenger RNA (mRNA), microRNA (miRNA), and long non-coding RNAs (lncRNA) as a novel class of stable and minimally invasive disease biomarkers.13-15 A growing body of reports suggested that dysregulated RNA molecules play a central role in the initiation and progression of numerous diseases, including cancers and motor neuron disease (MND).15-16 Being a critical regulator of post-transcriptional gene expression, microRNAs (miRNAs), a large group of small (~ 22 nucleotides) non-coding RNA, have garnered considerable recent interest as a diagnostic biomarker.17 Dysregulation of these highly conserved regulatory RNAs can result in etiology, progression, and prognosis of cancers. Several studies demonstrated that altered miRNA expression is a common feature of all human tumours, which is expressively altered from that of normal cells from the same tissue. Employing as a diagnostic and prognostic biomarker, circulating miRNA offer several distinct advantages such as the provision of early detection, better stability, and liquid biopsy for minimally invasive monitoring of cancer and other diseases. Despite the considerable potential for miRNAs in the diagnosis and prognosis of cancer, their detection approaches, are yet confined to several classic nucleic acid detection methods such as Northern blotting, microarray, RT-qPCR, and next-generation sequencing.11,18-19 Although there is no concern about their superiority for analytical performance and reliability, however, their scope is yet limited in the off-laboratory and resource-limited settings, where 3

sophisticated and expensive instruments might not be accessible. Also, these methods possess some inherent drawbacks. For instance, northern blotting consumed a large volume of samples and went through time-consuming steps and procedure.20 Microarray, being a high throughput technique, is usually more appropriate for discovery purpose instead of specific diagnostics.21 RT-qPCR, in comparison, is relatively sensitive; however, they rely on extensive laboratory infrastructure and technical support, lengthy assay steps (time-consuming sample preparation and purification, cDNA conversion, and qPCR).11,18 The development of a well-grounded platform that might be suitable for clinically relevant biomarker screening in resource-poor settings is, therefore, a priority. On that note, a great deal of research has been carried out to find relatively robust, sensitive, and specific methods, leading to several biosensor-based 11,18 miRNA sensing approaches coupled with optical and electrochemical readouts. Electrochemical detection methods are considered particularly attractive for bioanalysis, 22,23 because of their low cost, ease of automation, high sensitivity, and selectivity. However, the functionality of electrochemical miRNA sensors is still confined to the proof-of-concept studies, and several challenges to transform the technologies into routine clinical applications are yet to be addressed. These challenges include structural instability, non-specific interference from non-targets, and requirement of ultra-sensitivity. Besides, most of the assays involve complex electrode functionalization and rely on labelling agents and costly enzymatic amplification steps. One of the probable strategies to overcome those limitations and to enable a sensitive miRNA detection would be the integration of nanoarchitectures as signal transducer 18,24 and signal amplifier. Nanotechnology integrated electrochemical approaches generally offer relatively easy, robust, sensitive, multiplexed analysis in a portable point-of-care setting without labelling and signal amplification steps.25 Having smaller size, high surface-to-volume ratio, biocompatibility, and intrinsic electrocatalytic properties, nanoporous nanoarchitecture materials can overcome the barriers of structural miniaturization of diagnostics, leading to the prospect of designing inexpensive, sensitive portable devices.25,26 They provide pore-induced high surface area for uptake and release of biological molecules and redox species, a hybridization moiety for the efficient and faster analyte or target binding, biomimetic activity and cascade electrocatalytic signal amplification.27 In this PhD project, our research endeavour will be focused on the development of versatile biosensing platform consisting of a fundamental nanostructure-based study for obtaining optimum activity towards biosensing and four novel readout strategies for autoantibody and miRNA biomarker detection. Also to develop a platform that would have the high potentiality to address the criteria set by international experts for a point-of-care diagnostic 4

tool, i.e., a simple, accurate, and portable platform, which enables quick detection of miRNA- based biomarkers using the minimal equipment. Initially, we developed α and γ- iron oxides to find out which phase has a much higher potentiality for biosensing. Following that, a new class of gold-loaded iron oxide nanocube were synthesized, and activities for biosensing are comprehensively examined. By utilising the intrinsic nanozyme activity, a sensitive sensor was developed for autoantibody and based-on electrocatalytic event two amplification-free electrochemical approaches are developed for mRNA detection, which avoids any electrode modification, and relies on the direct adsorption of magnetically purified miRNA samples, on an unmodified disposable gold electrode. After realising this straightforward assay, we further exploited the design of mesoporous gold electrode for obtaining direct sample loading and significantly enhanced assay sensitivity while detecting cancer-associated miRNAs. Our final readout strategy will introduce a translational-focused assay platform, which enables sensitive electrochemical estimation of miRNAs in inexpensive portable devices for clinics.

1.2. Aims

The overall aim of this PhD study is to design and develop nanoarchitecture-based inexpensive, and simple biosensing platforms capable of efficiently detecting a range of crucial cancer- related biomarkers (autoantibody and micro RNA) in clinical samples with high specificity and sensitivity. After a comprehensive literature review on nanostructure synthesis, biofunctionalization, functionality towards biosensing for disease-related biomarkers, this research will investigate the development of optical (colourimetry) and electrochemical assays for autoantibody and miRNA biomarker detection. Moreover, an especial emphasize will also be given on nanoarchitecture and their intrinsic properties used for miRNA sensing. To address some of the existing limitations of natural enzyme HRP-based immunoassay for autoantibody detection and electrochemical miRNA sensing with poor sensitivity, a series of new functional nanostructure will be designed. The crystal-phase dependent superparamagnetism, nanozyme and electrocatalytic activity towards commonly used redox molecules will be studied. To obtain much simpler and ultra-sensitive detection, we have developed a mesoporous gold electrode which provides one-step detection of miRNA and translational direction of the developed sensor for routine analysis in clinics. This will be achieved with the following specific aims.

The specific aims are . To obtain a crystal structure for providing enhanced nanozyme (peroxidase-mimetic) activity, superparamagnetism, and electrocatalytic activity for biosensing . To develop a nanozyme-based optical and electrochemical assay for autoantibody detection and quantification; . To develop an electrochemical assay for ultra-sensitive miRNA detection . To produce a transduction surface (electrode) for electrochemical miRNA detection . To apply the assay platforms for clinical sample analysis

1.3. Significance of the project

The demand for point-of-care (biosensing) diagnostics and monitoring is proliferating with the concomitant increase in the prevalence of the chronic disease, increasing healthcare costs and unmet healthcare needs. Therefore, a great deal of research is going on in developing nanostructured-based novel methods and devices for diagnosis and monitoring of cancer via detecting levels of disease-specific biomarkers to meet this increasing demand. The importance of the research relates to the following scientific advances. Foremost, one of the significant limitations in disease diagnosis has been the lack of minimally invasive biomarkers, which can be detected and interrogated within complex biological samples with high sensitivity and specificity. In this regard, various species of clinically relevant autoantibody and miRNAs that are actively associated with the initiation and progression of cancers might prove to be a promising class of sensitive and specific biomarker. In this thesis, we have selected p53 autoantibody, which expresses in response to tumorigenesis in ovarian cancer, has been chosen. A set of miRNA, including miRNA miR-21 (specific to esophageal cancer), miR-9-2 (breast cancer) were also interrogated in this thesis. To check the versatility of our reported sensor, miRNA, named miR-338-3p from different disease state rather than cancer were chosen, which usually exhibits altered expression in a very low amount (need a highly sensitive sensor to detect it) in motor neuron disease (MND). Besides, these biomarkers can work as a liquid biopsy for non-invasive or minimally invasive interrogation of cancer. Secondly, the study reported in this thesis attempted to find the criterion for nanostructure design, i.e. the crystal phase-dependent magnetism and peroxidase mimetic activity, which are the crucial functions of nanomaterials required in biosensing for isolation, separation, purifications and signal readout. The gamma-phase nanostructure exhibits much higher superparamagnetism (depends on size of NPs and number of single domain present) and peroxidase mimetic activity (depends on exposed atomic sites and surface area of NPs). Though both two properties are independent to each other, however the as-prepared nanostructures exhibits a proportional relationship between them. Moreover, we have also reported on the design of multi-functional nanostructure which facilitate the adsorption of nucleic acids and showed cascade electrocatalysis towards popular redox marker. The integration of this multifunctional nanostructure in nucleic acid sensors eliminates time- consuming and cumbersome chemistries to obtain specific and sensitive detection of miRNA. Furthermore, we have reported on the design of the mesoporous gold electrode, which provides

a superior signal enhancement for miRNA facilitate sensitive nucleic acid detection without any cumbersome enzymatic amplification.

Thirdly, this thesis meets the increasing demand for detecting the ultralow levels of autoantibody and miRNAs from trace amounts of a complex biological sample. Gold-loaded nanoporous iron oxide nanocube was utilized to isolate target autoantibody for clinical samples and provide sensitive readout as nanozyme. This sensor design combined naked-eye, colourimetric and electrochemical readout. The LODs for both readout systems are better than that of the conventional p53-ELISA kit (0.08 vs ∼0.3 U/mL). The naked-eye discrimination of the autoantibodies described here holds huge potential for the development of a user-friendly and inexpensive bioassay in resource-limited settings, where sophisticated scientific equipment is unavailable. In particular, this approach can be exploited as a rapid first-pass screening (yes/no) tool to detect clinically relevant autoantibodies in a large population followed by more accurate and sensitive quantification of autoantibody via electrochemical readout. The assay also replaces natural enzymes for TMB oxidation, and thus reduces the cost, handling, and storage facilities generally required for natural enzymes. All the miRNA readout platforms have shown excellent sensitivity, ranging from picomolar to attomolar level (10 pM for miR- 21, 100 aM for miR-338-3p). Such low LOD is adequate to retrieve disease information from a minute amount of clinical sample. This high sensitivity could be ascribed to several reasonable assay components such as a) high catalytic activity and immense sample loading capacity of the newly synthesised gold-loaded nanoporous iron oxide nanocube based sample loading and electrocatalytic signal amplification; b) additional signal enhancement with multiple electrocatalytic cycles (redox-cycling); and c) the poor specificity due to high level of sequence overlap between various RNA molecules and presence of non-specific molecules, were addressed by using repetitive magnetic bead-based washing, isolation, and purification step in reported miRNA sensors. It can, therefore, be concluded that our advanced assay platforms have also improved the standard concept of autoantibody and miRNA biosensing technology, which often relies on sensor fabrications, signalling proteins, hybridisation of targets with surface-bound capture probes, and the use of electroactive ligands. One of the foremost significances here is the use of single-use disposable screen-printed electrodes, which successfully eliminates the utilisation of normal electrochemical cells, as well as counter and reference electrodes, thereby offering a relatively inexpensive (approximately AUD 5) and rapid platform. We have also successfully demonstrated a broad platform technology based on an own-prepared mesoporous gold 8

electrode, which could enable the one-step detection of miRNA after obtaining samples from clinics. Overall, we believe that our mesoporous gold electrode-based approach can be further extended towards the development of a multiplexed electrochemical device that could have huge potential in miniaturisation and portability of the biosensor with minimal cost, space, and power requirements. We also envisage that our studies might not only pave the way for improved patient care, specifically in impoverished regions, by developing inexpensive, sensitive, and specific detection platforms for personalised and point-of-care clinical applications but also prove to be a productive commercial venture.

1.4. Structure of the thesis

This thesis includes eight chapters. Chapters 2, 3, 4, 5, 6, and 7 are a collection of journal papers that have been published or are under consideration

Chapter 1. This chapter introduces the aims, background and significance of this research.

Chapter 2. This chapter presents a comprehensive literature review focusing on nanostructure- based sensing for clinically relevant disease biomarkers. Initially, we comprehensively reviewed the biomolecule specific magnetic nanostructure synthesis, their further biofunctionalization, intrinsic properties required for achieving robust biosensing and magnetic nanostructure-based electrochemical and optical biosensor (nucleic acid sensor, immunosensor, cytosensor, etc.) and nanostructure-based point-of-care diagnostics. With a comprehensive discussion on each section, this review will also chronicle the needs and challenges of nanoarchitecture-based detection. Coming next, we predominantly focused on nanostructure used for electrochemical miRNA Detection. We reviewed the significant contributions of engineered nanomaterials-based electrochemical biosensing strategies for the analysis of miRNAs. With a diverse emphasis on nanostructure-based detection, this review also accounts the needs and challenges of miRNA detection and provides a future perspective on the presented strategies. Chapter 2 provides the reasoning for the research described in the subsequent sections.

Chapter 3. In this chapter, we methodically investigate the influence of crystal phases (γ-Fe2O3 and α-Fe2O3) of mesoporous iron oxide (IO) on their peroxidase mimetic activity towards the oxidation of chromogenic substances, such as 3,3′,5,5′-tetramethylbenzidine (TMB) and 2,2'- azino-bis(3-ethylbenzothiazoline-6-sulphonic acid)-ABTS. Upon studying superparamagnetism and the apparent Michaelis–Menten constant (Km and Vmax) of crystal phase- materials towards the oxidation of TMB. Our mesoporous γ-Fe2O3 shows high nanozyme activities (and magnetism) toward the catalytic oxidation of TMB and ABTS. To check the findings in biosensing, we have demonstrated a proof-of-concept method for detecting glucose, where mesoporous γ-Fe2O3 exhibits higher responses for colourimetric detection of glucose than that of α-Fe2O and achieved a LOD of 0.9 µM. We believe that this in-depth study of crystal structure-based nanozyme activity will guide designing highly effective nanozymes

based on iron oxide nanostructures for chemical sensing, biosensing and environmental remediation.

Chapter 4. After the findings in the previous chapter, we have purposefully developed a new class gold loaded nanoporous ferric oxide nanocubes (Au-NPFe2O3NC) materials the development of a molecular sensor with enhanced electrocatalytic and colourimetric (naked- eye) detection of autoantibodies in clinical samples. The as-prepared Au-NPFe2O3NC exhibits excellent super-paramagnetism which was utilized for isolating p53 autoantibody (specific to ovarian cancer) from plasma and serum samples; enzyme-mimicking activity (nanozyme) of towards the catalytic oxidation TMB in the presence of H2O2 at room temperature (25°C) and follows the typical Michaelis–Menten kinetics. The intrinsic nanozyme activity of Au-

NPFe2O3NC was used as signal reporters specific to the target. The autoantibody sensor based on this intrinsic property of Au-NPFe2O3NC resulted excellent detection sensitivity (LOD = 0.08 U/mL) and reproducibility (% RSD = < 5% for n = 3) for analysing p53-specific autoantibodies using electrochemical and colorimetric (naked-eye) readouts. The clinical applicability of the sensor has been tested in detecting p53-specific autoantibody in plasma obtained from patients with epithelial ovarian cancer high-grade serous subtype (EOCHGS) and controls (benign).

Chapter 5. To design a nucleic acid sensor, we have then examined the electrocatalytic activity of Au-NPFe2O3NC nanocube towards the electrochemical redox reaction of ruthenium hexaammine (III) chloride (RuHex), a commonly used redox molecule for nucleic acid-sensing. With the higher catalytic activity, the nanocube provides a superior transduction surface for target miRNA immobilization and signal enhancement. This chapter demonstrated that this nanocube offers a nonenzymatic, amplification free, and highly sensitive platform for electrochemical detection of microRNA (miR-21) in clinical samples. The miRNA was directly adsorbed onto the exposed gold of the Au@NPFe2O3NC that were magnetically attached to the screen-printed carbon electrode preceded by magnetic isolation, separation and purification steps. The assay showed excellent detection sensitivity (up to 10 pM) and specificity for the analysis of miR-21 in oesophageal squamous cell carcinoma cell lines and clinical tissue samples without any enzymatic amplification or extension or cumbersome electrode surface modification step.

Chapter 6. Extending on the research from Chapter 5, this chapter discusses the further enhancement in the sensitivity of miRNA detection (LOD = 100 aM) assay. Exploiting the high surface loading and catalytic activity Au-NPFe2O3NC towards another redox molecule methylene blue (MB). To accelerate the detection signal, the chronocoulometric interrogation of MB (with surface-bound miRNA) was coupled with [Fe(CN)6]3-/4- and a [Fe(CN)6]3-/4- - MB redox cycling has been achieved. The applicability of the method was tested and validated in a set of motor neuron disease patient samples.

Chapter 7. After successful demonstration of highly specific, sensitive and specific biosensors for miRNA in Chapters 5 and 6, the research moved to the development of a new platform technology for detecting miRNA incorporating mesoporous gold electrode and novel differential pulse voltammetry readout. This approach attains a wide dynamic linear range from 100 aM to 1 nM with an ultra-low limit detection of 100 aM, exerting a notable augmentation in sensitivity. Moreover, the platform evades the cumbersome PCR and enzymatic amplification steps or transduction surface modification with enzyme, capture probe or any physical functionalization (e.g. nanostructure modification). Besides, it simplifies the assay building by circumventing multiple steps tangled in conventional biosensing approaches through recognition and transduction layers.

Chapter 8. Conclusions and Future Recommendations are provided in this chapter.

1.5. References

1. Society, A. C., Global Cancer Facts & Figures 3rd Edition. American Cancer Society 2015, 800, 1-64.

2. Australian Institute of Health and Welfare. Cancer in Australia: In brief 2019. Cancer series no. 122. Cat no. CAN 126. Canberra: AIHW. 3. Ebos, J. M.; Kerbel, R. S., Antiangiogenic therapy: impact on invasion, disease progression, and metastasis. Nature Reviews Clinical oncology 2011 8 (4), 210. 4. Yadav, S.; Kashaninejad, N.; Masud, M. K.; Yamauchi, Y.; Nguyen, N. T.; Shiddiky, M. J. A., Autoantibodies as diagnostic and prognostic cancer biomarker: Detection techniques and approaches, Biosensors and Bioelectronics 2019, 111315. 5. Aggarwal, A., Role of autoantibody testing. Best Practice & Research Clinical Rheumatology, 2014, 28 (6), 907-920. 6. Caron, M.; Choquet-Kastylevsky, G.; Joubert-Caron, R., Cancer immunomics using autoantibody signatures for biomarker discovery. Molecular & Cellular Proteomics 2007, 6 (7), 1115-1122. 7. Anderson, K. S.; Sibani, S.; Wallstrom, G.; Qiu, J.; Mendoza, E. A.; Raphael, J.; Hainsworth, E.; Montor, W. R.; Wong, J.; Park, J. G.; Lokko, N., Protein microarray signature of autoantibody biomarkers for the early detection of breast cancer. Journal of Proteome Research 2010, 10 (1), 85-96. 8. Chapman, C. J.; Thorpe, A. J.; Murray, A.; Parsy-Kowalska, C. B.; Allen, J.; Stafford, K. M.; Chauhan, A. S.; Kite, T. A.; Maddison, P.; Robertson, J. F., Immunobiomarkers in small cell lung cancer: potential early cancer signals. Clinical Cancer Research 2011, 17, 1474-1480. 9. Anderson, K. S.; Cramer, D. W.; Sibani, S.; Wallstrom, G.; Wong, J.; Park, J.; Qiu, J.; Vitonis, A.; LaBaer, J., Autoantibody signature for the serologic detection of ovarian cancer. Journal of Proteome Research 2014, 14 (1), 578-586. 10. Macdonald, I. K.; Parsy-Kowalska, C. B.; Chapman, C. J., Autoantibodies: opportunities for early cancer detection. Trends in Cancer 2017, 3 (3), 198-213. 11. Yadav, S.; Masud, M. K.; Islam, M. N.; Gopalan, V.; Lam, A. K. Y.; Tanaka, S.; Nguyen, N. T.; Hossain, M. S. A.; Li, C.; Yamauchi, Y.; Shiddiky, M. J., Gold-loaded nanoporous iron oxide nanocubes: a novel dispersible capture agent for tumor- associated autoantibody analysis in serum. Nanoscale 2017, 9, 8805-8814.

12. Garranzo-Asensio, M.; Guzman-Aranguez, A.; Povés, C.; Fernández-Aceñero, M. J.; Torrente-Rodríguez, R. M.; Ruiz-Valdepeñas Montiel, V.; Domínguez, G.; Frutos, L. S.; Rodríguez, N.; Villalba, M.; Pingarrón, J. M., Toward liquid biopsy: determination of the humoral immune response in cancer patients using halotag fusion protein- modified electrochemical bioplatforms. Analytical Chemistry 2016, 88(24), pp.12339- 12345. 13. Islam, M. N.; Masud, M. K.; Haque, M. H.; Hossain, M. S. A.; Yamauchi, Y.; Nguyen, N. T.; Shiddiky, M.J. A., RNA biomarkers: diagnostic and prognostic potentials and recent developments of electrochemical biosensors. Small Methods 2017, 1 (7), 1700131. 14. Ludwig, J. A.; Weinstein, J. N., Biomarkers in cancer staging, prognosis and treatment selection. Nature Reviews Cancer 2005, 5 (11), 845. 15. Esquela-Kerscher, A.; Slack, F. J., Oncomirs—microRNAs with a role in cancer. Nature Reviews Cancer 2006, 6 (4), 259. 16. Haramati, S.; Chapnik, E.; Sztainberg, Y.; Eilam, R.; Zwang, R.; Gershoni, N.; McGlinn, E.; Heiser, P. W.; Wills, A. M.; Wirguin, I.; Rubin, L. L.; miRNA malfunction causes spinal motor neuron disease. Proceedings of the National Academy of Sciences 2010, 107 (29), 13111-13116. 17. Nelson, K. M.; Weiss, G. J., MicroRNAs and cancer: past, present, and potential future. Molecular Cancer Therapeutics 2008, 7 (12), 3655-3660. 18. Kilic, T.; Erdem, A.; Ozsoz, M.; Carrara, S., microRNA biosensors: opportunities and challenges among conventional and commercially available techniques. Biosensors and Bioelectronics 2018, 99, 525-546. 19. Bertoli, G.; Cava, C.; Castiglioni, I., MicroRNAs: new biomarkers for diagnosis, prognosis, therapy prediction and therapeutic tools for breast cancer. Theranostics 2015, 5(10), p.1122. 20. Koshiol, J.; Wang, E.; Zhao, Y.; Marincola, F.; Landi, M. T., Strengths and limitations of laboratory procedures for microRNA detection. Cancer Epidemiology and Prevention Biomarkers 2010, 19, 907-911. 21. Babak, T.; Zhang, W. E. N.; Morris, Q.; Blencowe, B. J.; Hughes, T. R., Probing microRNAs with microarrays: tissue specificity and functional inference. RNA 2004,10 (11), 1813-1819. 22. Lautner, G.; Gyurcsányi, R. E., Electrochemical detection of miRNAs. Electroanalysis 2014, 26 (6), 1224-1235. 14

23. Ronkainen, N. J.; Halsall, H. B.; Heineman, W. R., Electrochemical biosensors. Chemical Society Reviews 2010, 39 (5), 1747-1763. 24. Zhu, C.; Yang, G.; Li, H.; Du, D.; Lin, Y., Electrochemical sensors and biosensors based on nanomaterials and nanostructures. Analytical Chemistry 2014, 87 (1), 230- 249. 25. Masud, M. K.; Umer, M.; Hossain, M. S. A.; Yamauchi, Y.; Nguyen, N. T.; Shiddiky, M. J. A., Nanoarchitecture frameworks for electrochemical miRNA detection. Trends in Biochemical Sciences 2019, 44 (5), 433-452. 26. Quesada-González, D.; Merkoçi, A., Nanomaterial-based devices for point-of-care diagnostic applications. Chemical Society Reviews 2018, 47 (13), 4697-4709. 27. Masud, M. K.; Islam, M. N.; Haque, M. H.; Tanaka, S.; Gopalan, V.; Alici, G.; Nguyen, N. T.; Lam, A. K.; Hossain, M. S. A.; Yamauchi, Y.; Shiddiky, M. J., Gold-loaded nanoporous superparamagnetic nanocubes for catalytic signal amplification in detecting miRNA. Chemical Communications 2017, 53 (58), 8231-8234.

Chapter 2

Literature Review

Nano-architectures for disease-specific biomarker detection

Sections of this chapter are based on Masud, M.K., et al., Chemical Society Reviews, 2019, 48 (24), 5717-5751* and Sections of this chapter are based on Masud, M.K., et al., Trends in Biochemical Sciences, 2019, 44 (5), 433-452.**

2. Literature Review: Nano-architectures for disease-specific biomarker detection

2.1. Superparamagnetic nanoarchitectures* Detection of a disease-specific biomolecular target, including DNA- and RNA-based biomarkers, circulating tumour cells, and small molecules (exosomes), is essential to understand its physiological and biological functions, disease diagnosis, and prognosis.1-5 These biomolecules carry out several anatomical and physiological functions: for instance, DNA encodes genetic information for the development and regulation of gene expression; RNA transmits that genetic information and translates it into proteins for structural and regulatory roles and therefore is involved in protein expression and different cellular functions.6-8 Exosomes, membrane-bound cargo that is enriched with proteins, lipid rafts, micro RNAs (miRNAs), messenger RNAs (mRNAs), and other noncoding RNAs (ncRNAs), exchange genetic information between neighbouring cells during their circulation.9 Moreover, they are now routinely used for the diagnosis and management of many diseases as the disease progresses, resulting in an alteration of the physiological state or genetic expression of these biomolecules. Consequently, there has been a rising demand for detection of those biomolecules, especially for early diagnosis and treatment. Early-stage detection enables effective therapies to reduce suffering and disease-related burdens. Over the last century, numerous detection platforms have been developed to detect these biomolecules. These methodologies range from classical molecular biology to advanced procedures, such as bisulphite sequencing,10 microarrays,11 quantitative real-time PCR (qRT-PCR),12 RNA sequencing,13 colourimetry,14 surface plasmon resonance (SPR),15 and surface-enhanced Raman spectroscopy (SERS),16 to more recent analytical approaches such as high-performance liquid chromatography (HPLC),17 mass spectrophotometry (MS),18 and electrochemical biosensors.6, 19 The classical molecular biology techniques, despite their robustness and high efficiency, nevertheless suffer from sensitivity and specificity (amplification biases) and require cumbersome sample pretreatment and expensive instrumentation.20-22 HPLC and MS can provide rapid, accurate, and selective detection, but they are limited in practical application due to their high cost, bulky equipment size, immobility, specialized operation, and throughput. Notably, they are not suitable when an ultrasensitive, miniaturized, portable detection system is looked for infield and wearable applications.23-24 Optical and electrochemical methodologies are relatively inexpensive and rapid, and they provide sensitive detection in a portable arrangement with a small volume input of clinical samples.6, 19, 25 Furthermore, tremendous advances in microfabrication and allied technologies have been improving the design and

development of electrical readout-based chemical sensors and biosensors.26 However, target- specific electrode surface modification is vital to achieving the selectivity of a functional biosensor. The most straightforward and widely used approach is to immobilize enzymes or proteins (antibodies) on the electrode with a polymer layer.27 Additionally, the sensor needs to bind with signalling or redox molecules to produce the target-specific responses. Even though a plethora of cutting-edge electrochemical biosensors have been developed, most of the approaches still suffer from low sensitivity, complex instrumentation, and multifaceted, tedious, and time-consuming chemistry. Furthermore, the clinical application of most of these has yet to be realized. With the advancement of nanotechnology, nanostructured magnetic materials have aroused immense interest in the field of analytical sensing and biomedicine due to their flexible and modular structure, easy synthesis, small size, low toxicity, intrinsic enzyme-mimicking activity, superparamagnetic behaviour, and biocompatibility.28-30 With their small volume, magnetic NPs (MNPs) smaller than the single domain limit (~20 nm for iron oxide) reveal superparamagnetism at room temperature; i.e., MNPs (ferromagnetic) lose their magnetism below their Curie temperature.30-32 In brief, the magnetic moment of an individual NP able to rotate randomly (in reference to the orientation of the MNP) by the influence of temperature. Due to the fact that, in the absence of an electromagnetic field, the net magnetic moment of NPs became zero at high enough temperatures. However, in the presence of a magnetic field, a net statistical alignment of magnetic moments occurs, similar to that of paramagnetic materials. This characteristic, obvious by the lack of remnant magnetization (i.e., the value of the magnetization at zero fields) after elimination of external magnetic fields. This enables MNPs to maintain their colloidal stability and avoid agglomeration, which is essential for magnetic manipulation of the sample in order to achieve nonspecific response free, highly specific and sensitive detection of target molecules.33 MNPs uniquely combine with more modestly sized or same-size molecular analytes and hence are often involved in isolation, purification, target-molecule-carrying, signal-generating, and signal-enhancing steps in biosensing, resulting in specific and highly sensitive diagnosis in clinics (Figure 2.1.1).34-36 Nanostructured materials also exhibit impressive advantages in molecular diagnostics, particularly in disease diagnosis applications. For instance, they break down the barrier to structural miniaturization of diagnostic platforms, enable direct contact with sensing environments (e.g., electrolyte, labeller), and provide reagent-less biosensing, biomimetic, in vivo detection, allowing them to be used as carrier or capture vehicles for loading a large number of specific biological probes.37-39 In addition, the plasmonic and electrochemical 18

properties of nanostructured materials can be exploited to adopt many novel transduction schemes, and the nanozyme activity could potentially replace natural enzymes in a wide range of uses in ELISA-like biosensing.36, 40 Over the past few decades, numerous superparamagnetic

NPs have been synthesized, including iron oxides (Fe3O4 and Fe2O3); different ferrites of cobalt, nickel, and manganese; gold-containing ferric oxide; graphene; and other functional nanostructured wrapped iron oxides.41-43 In recent years, porous nanomaterials have also attracted increasing interest, as they possess a large surface area and large pore volumes, narrow pore-size distribution, high loading capacity, and modifiable surface characteristics.42, 44-45 These intrinsic properties enabled them highly potential for the uptake and release of guest molecules. The porous structure has also provided enlarged catalytic volume which increases the mass transfer as compared to that of bulk materials of the same mass.46-47 Notably, for detecting biomarkers, each nanostructure is designed in such a way that it is biocompatible with the target (cells, proteins, exosomes, DNA, RNA, etc.) and can be integrated into target- isolation and purification, immobilization, signal-transduction, signal-generating, or signal- amplification steps. Thus, nanostructures need to be highly specific and selective of the biomolecule structure (shape, size, length, charge, interaction affinity, etc.). Based on the synthesis, intrinsic characteristics, and application of nanostructure materials, several reviews have also been authored in which their different synthetic strategies, functionalization, and biosensing were discussed.34, 43, 48-52 There is extensive literature based on the promise, facts, and challenges of the various nanostructures, e.g., graphene,53 carbon nanotubes,54 and quantum dots55 in biomedical applications, and the human healthcare field has already been expressly focusing on individual materials and their uses in biosensing. Recently, we authored “Nanoarchitecture Frameworks for Electrochemical miRNA Detection,”56 in which the pros and cons of previously reported nanostructures were discussed concerning microRNA detection. Another review also explored the structure-based relationship for sensing performance; however, no specific biomolecules were considered.57 No reviews have yet focused on the nanostructures that are specifically engineered for the detection of disease-specific biomarkers. Reports are needed in which the target-specific and unique engineering of nanostructures is comprehensively discussed to guide future science in detecting biomarkers, and hence disease diagnosis and prognosis. In summary, a review that focuses on biomolecule-specific magnetic nanostructure synthesis, further biofunctionalization, and intrinsic properties is required to achieve robust biosensing and magnetic nanostructure-based electrochemical and optical biosensors (nucleic acid sensor, immunosensor, cytosensor, etc.). 19

In this review, we have focused extensively on the biomolecule-specific magnetic nanostructure synthesis, further biofunctionalization, and intrinsic properties required to achieve robust biosensing and magnetic nanostructure-based, clinically relevant disease biomarker detection. Following the particular emphasis on nanostructure-based electrochemical and optical detection, the nanostructure-based point-of-care diagnostics are also discussed. With a comprehensive discussion in each section, this review also chronicles the needs and challenges involved in nanoarchitecture-based biomarker detection.

Figure 2.1.1: Schematic representation of the building blocks of nanoengineered superparamagnetic nanoparticles, their surface functionalization, and their functions for integration into biomolecular biosensors.

2.1.1. Synthesis of superparamagnetic nanoparticles To date, various approaches have been developed for the synthesis of superparamagnetic NPs and porous nanomaterials with controlled narrow size distribution for desired chemical and physical properties. Among the types of superparamagnetic NPs, ferrite colloids, magnetite, and maghemite are widely studied due to their biodegradability and biocompatibility. Broadly, the synthetic methodology can be categorized into three classes: general synthesis of MNPs, such as coprecipitation, thermal decomposition, microemulsion, hydrothermal reaction, and sol-gel synthesis; templating methods for the synthesis of porous nanomaterials; and doped metal synthesis for biofavorable hybrid metal NPs.

2.1.1.1. General synthesis The conventional and most commonly used wet chemical methods for the synthesis of superparamagnetic iron oxide nanoparticles (IONPs) are the coprecipitation method, wherein

Fe3O4 and γ-Fe2O3 are precipitated from a basic solution of ferric and ferrous salts or by oxidation of a ferrous hydroxide suspension using oxidizing agents.58-59 A mixture of ferric and ferrous ions in a 1:2 molar ratio is used to obtain iron oxide precipitation at room temperature or elevated temperature according to the following reaction;

. The size and shape of the IONPs depend on the type of iron salt (e.g., chloride, sulfide, or nitrite), ferric and ferrous ion ratio, pH of the solution, reaction temperature, and reaction conditions. One of the most pioneering examples of this method was reported by Sugimoto

(1980): iron oxides were obtained by the interaction of FeSO4 with KOH in the presence of a nitrate ion followed by aging of the resultant gelatinous suspension at 90 °C for several hours.60 Since then, several coprecipitation-based methods have been developed to synthesize IONPs.

For example, Fe3O4 magnetic powder with an average gain diameter of 15 nm was prepared from high-purity iron using ultrasonic-assisted chemical coprecipitation.61 Recently, Pereira et al. reported a one-step aqueous coprecipitation-based method for the synthesis of MFe2O4 using the alkanolamines isopropanolamine and di-iso-propanolamine as an alkaline agent.62 The synthesized MFe2O4 nanomaterials exhibited high colloidal stability, particle sizes in the range of 4−12 nm, and superparamagnetic properties. Moreover, this method generated smaller particle sizes (up to 6 times) and superior saturation magnetization (up to 1.3 times) than those prepared with NaOH or KOH. This method can be used to prepare IONPs on a large scale in a short time. NPs of different sizes and morphologies were also obtained by adjusting the pH,

oxidizing agents, ionic strength, and concentration of the growth solution. Moreover, the stirring rate and reaction time have a considerable effect on the structural properties, specifically the particle sizes and corresponding magnetic properties.63 However, the coprecipitation method is limited by the fact that the produced NPs tend to agglomerate in aqueous and physiological conditions. To overcome this limitation, different polymers, polyethylene or surfactants such as dextran and polyvinylalcohol (PVA), are used to immobilize the IONPs.64-65 The thermal decomposition approach can be used to obtain highly crystalline, monodisperse, and narrowly size-distributed IONPs from the high-temperature thermal decomposition of coordinated iron precursors or organometallics such as Fe (cup)3 (cup = N- 66 nitrosophenylhydroxylamine), Fe(acac)3 (acetylacetonate), or Fe(CO)5 in organic solvents. In this method, precursors are injected into either a hot reaction mixture (high temperature) or a room temperature reaction mixture, followed by heating in a closed or open reaction vessel. A size-controlled monodispersed IONPs synthesis was reported by Sun and Zeng, wherein

Fe(acac)3 in phenyl ether was heated to 265 °C in the presence of alcohol, oleic acid, and oleylamine. Larger monodispersed NPs with a size of up to 20 nm were obtained by using smaller MNP seeds.67 This procedure does not need a size selection process. The size of the IONPs is controlled by varying the aging temperature and other reaction parameters. This method is suitable for preparing NPs with a different shape (nanospheres or nanocubes). The size and shape can also be tailored by using varieties of precursors, solvents, or additives during the thermal decomposition process. One of the shortcomings of this approach is that the exact shape of the IONPs is not reproducible, as the nucleation of NPs involves boiling the solvents. The nanomaterial produced via this method is usually dissolved in nonpolar solvents. An alternative wet chemical method for obtaining crystalline NPs is the hydrothermal one, wherein a mixture of iron salts was dissolved in an aqueous medium and heated in a sealed Teflon container at a temperature (130 to 250 °C) higher than the boiling point of water and high vapour pressure (0.3 to 4 MPa).68-69 This method generates higher crystalline MNPs with a superior magnetic feature due to the synergistic effect of high temperature and pressure. The microemulsion method is also employed for the synthesis of shape- and size-controlled MNPs. The binary system (water/surfactants or oil/surfactants) of microemulsion, which can be formed by different types of self-assembled structures such as spherical and cylindrical micelles, enables the desired growth, nucleation, and agglomeration of NPs.70 In this method, an iron-containing nanoemulsion is mixed with sodium hydroxide, followed by lysis with acetone to remove the surfactants. The dynamics of MNPs and controlled size can be achieved 22

by varying the droplet size, reactant concentrations, and surfactant nature. Although these colloidal NPs show higher superparamagnetic properties, they need several washing procedures to remove surfactants and further stabilization before use in biomedical applications. Another two-phase method for the synthesis of NPs is the sol-gel method, wherein the hydroxylation and condensation of precursor molecules in solution generate the sol of nanometric particles. A three-dimensional metal oxide wet gel is obtained by further condensation and inorganic polymerization. Heat treatments are also required to obtain a crystalline state, as gel preparation is carried out at room temperature. The predetermined nanostructure with a pure amorphous phase and monodisperse, size-controlled NPs can be 71 synthesized using the sol-gel method. For instance, γ-Fe2O3 in silica with a size of 15 to 30 nm was synthesized by heating the gel to 400 °C. The gel was prepared by the hydrolysis of 72 Fe(NO3)3·9H2O and tetraethyl orthosilicate (TEOS) in ethanol. Some physical methods have also been developed to synthesize MNPs, such as electron beam lithography and gas-phase deposition.73 These methods required tedious and time-consuming procedures and were unable to generate size-controlled NPs. In comparison to physical processes, the wet chemical methods are more straightforward, more tractable, and more efficient for the synthesis of monodisperse, highly crystalline, superparamagnetic NPs with controlled size, shape, and composition.74

2.1.1.2. Template-based synthesis

The term “template synthesis” represents the direct preparation of nanomaterials with uniform morphology, such as size and shape, using a central structure as a template.69, 75 Within the template, a network structure is formed in such a manner that removal of the template may create a cavity with the designed morphological and stereochemical nanostructure. The template enables higher reproducibility of the structure and provides a skeleton for obtaining the desired function of a nanostructure.76 Template synthesis generally involves three main steps: first, selection or creation of the template; second, assembly or synthesis of the desired nanomaterials using common synthetic strategies; and third, removal of the template to generate a porous nanostructure. Broadly, there are three template-based syntheses: (i) soft- templating methods, (ii) hard-templating methods, and (iii) sacrificial-templating methods. The soft-templating methods use structurally flexible materials such as surfactants, polymers, micelles, and viruses, whereas in hard templating, the porous materials were prepared using a rigid structure such as colloidal silica, latex, or a carbon sphere.77 Elimination of the template

is relatively simple and easy in soft templating. Hard templating is advantageous for the synthesis of stable, controlled morphological NPs, but it requires tedious template removal procedures. Thus, proper template selection is a prerequisite for obtaining the desired porous nanostructure. Soft-templating approaches have been used for the synthesis of a variety of hollow nano- and microstructures, including superparamagnetic metal oxides78 and metals (e.g., Ni, Pd, and Ag).79-81 An emulsion, micelle, vesicle, or hydrophilic polymer is generally employed as a template. These soft templates are usually aggregated via inter- and intramolecular interactions such as hydrogen bonding, Van der Waals forces, and chemical bonding.82 The inorganic precursor is then deposited onto the surface of the interior of the template following conventional precipitation, electrochemical, or other synthetic approaches. Finally, removal of the soft template is usually carried out via a more straightforward process, such as washing or evaporation. This method offers excellent repeatability and simpler operation and avoids chemical or structural changes during template removal. As the biological system requires mild template removal, this technique has the potential for the synthesis of a hollow structure with bio-sensitive functionalities such as molecular catalysis, biocatalysis, and biosensing. Moreover, using an emulsion template, hollow shells with an additional functional component such as bioactive or catalytically active species can be synthesized by incorporating them into 83 the hollow materials. For example, Lu et al. reported achieving a magnetic Fe3O4@h-

C/noble-metal rattle-type NP by incorporating surface-functionalized, preexisting Fe3O4 nanocrystals into the droplets of oil/water emulsion, followed by interfacial polymerization to 84 generate a hollow polymer shell Fe3O4@h-P. The shell contains a carboxylate group and binds the noble metal via an ion exchange process. The metal cation was then converted to nanocrystals via pyrolysis under an inert atmosphere. Soft templates are sensitive to reaction parameters such as pH, temperature, and solvent polarity, which presents challenges to using this method to obtain a hollow structure with a particular structural feature. In hard-template synthesis, a rigid material provides the size, morphology, and surface properties of nanostructured materials. Colloidal SiO2, latex, and carbon spheres are the most frequently used materials as a hard template. In addition to the three steps of the soft-templating method, hard templating adopts one more step. The hard template inhibits the aggregation or crystallization of the precursor molecules and allows the generation of NPs with a structure opposite to that of the template. The surface of the as-synthesized template materials needs to be either modified or functionalized to fit its chemical compatibility with the precursor molecules. Different synthetic approaches to forming the shell on the template have been 24

applied to prepare the desired hollow nanostructure. The shells are formed by adsorption, layer- by-layer methods, chemical deposition, or nanocasting.69 In the adsorption methods, precursor molecules are adsorbed on the surface of the template materials via electrostatic attraction followed by thermal treatment to induce cross-linking and development of the continuous shell.

For example, Wang et al. demonstrated the synthesis of three nanoporous metal oxides (Fe2O3 nanorods, NiO nanosheets, and Co3O4 NPs) using sulfonated polystyrene (SP) microspheres as a hard template.85 The SP microsphere was achieved by gradual sulfonation of the outer sphere of polystyrene. When the sulfonated SP microsphere was placed in the solution of precursor metal salt, the ions from the precursor adsorbed within the outer surface of the sphere. A further heat treatment simultaneously cross-linked the metal precursors into the shell and removed the template polymer particles, thereby generating the hollow (hierarchical) metal oxide. Layer- by-layer methods also involve the electrostatic adsorption of metal precursor ions around the template materials to form alternating layers of oppositely charged building blocks.86 Unlike the adsorption or layer-by-layer shell formation, the chemical deposition method utilizes the chemical attachment of the shell precursors to the template surface. Here, cross-linking of the precursor occurs via condensation or polymerization reactions and does not require further heat treatment to develop a continuous shell around the template materials.87 Nanocasting synthesis offers superior structural features of hollow materials, such as shell thickness, porosity, and higher mass-transport properties.88 In the above-discussed methods, a porous scaffold around the shell is initially formed, followed by the infiltration of precursors into the porous rim of the template materials. Thermal or chemical treatment is required for appropriate cross-linking. The synthesis of nonspherical hollow materials using a hard template remains challenging due to the lack of templates and difficulties in achieving uniform surface (sharp edge and corners) coverage on the template.89 Another template-based method, called a sacrificial template, is used to synthesize various magnetic metal oxide NPs. The use of metal-organic frameworks (MOFs) or porous coordination polymers (PCPs) as a sacrificial template has drawn significant attention due to their structural diversity, large surface area, and different surface morphologies.90 Though thermal decomposition or calcination of iron oxide or iron hydroxide at a certain temperature generated porous NPs, the pore volumes and pore size obtained by this method are not very large. However, thermal treatment of MOFs or PCPs produces microporous metal/metal oxide NPs with a large surface area. Prussian blue (PB) coordination polymers and PB analogues are considered the most promising cyano-bridged MOFs for preparing porous metal oxide NPs. In such MOFs, iron ions are bridged by cyano groups [(-Fe-CN-Fe)-]. Cyano groups are labile 25

and easily removed (C and N are oxidized into gases and escape) by calcination, resulting in porosity in the metal nanostructure with super/quasisuperparamagnetism.91 Recently, some attempts to synthesize porous iron oxide through the thermal decomposition of PB have been reported. For instance, Hu et al. synthesized elongated PB nanocubes via selective etching followed by the conversion of PB crystals into iron(III) oxides with different crystalline phases.91-92 However, the obtained NPs had a low surface area, poor crystallinity, and several crystalline phases (α, , and γ phases). To overcome such limitations, Yamauchi et al. synthesized nanoporous IOs (crystalline α-Fe2O3 and γ-Fe2O3) with hollow interiors by calcinating previously prepared hollow PB nanocubes.91 The crystalline phases and their crystalline grain sizes were controlled by the volume of internal hollow cavities in PB nanocubes and by the calcination temperature (Figure 2.1.2). The cubic structure was preserved even at a higher calcination temperature (400 °C). Moreover, the crystalline phase of porous nanocubes became higher at an elevated temperature, but the particle size slightly decreased with increasing temperature. The reduction in particle size arises from the higher degree of decomposition of cyano groups. In addition to this method, later on, Yamauchi et al. controlled the particle size (from 20 nm to 500 nm) of nanoporous metal oxides by adjusting the amount of sodium citrate chelating agent for the synthesis of hollow PB nanocubes.93

Figure 2.1.2: Schematic illustration of the fate of PB nanoparticles at different applied calcination temperatures. Reproduced with permission.91

2.1.1.3. Multicomponent (hybrid) magnetic nanoparticle synthesis Hybrid nanomaterials combining two or more functional constituents and nanoscale functionalities have attracted increasing interest due to their numerous applications in electronics, catalysis, bioimaging, biotechnology, and nanotechnology.94 In hybrid (bimetallic) materials, one metal may confer the electrochemical or optical or both properties, whereas others may provide long-term stability, biocompatibility, and specific affinity to target biomolecules. During synthesis, the individual component may be combined and optimized independently for target analysis. Hybrid materials achieve the cooperative performance of individuals via interaction between or among constituents. To date, most biomolecular analyses involving hybrid materials have been reported with the combination of MNPs (mainly iron oxide) with gold (Au). However, other metals such as Pt, Zn, and Cu have also been explored. In addition to these, the synthesis of carbon and graphene, silica-containing hybrid materials, has been reported as a promising platform for biomedical applications. The general strategy for obtaining composite NPs is to prepare one NP, followed by coating or loading another constituent (metal or nonmetal) or using the first NP as a nucleation seed to deposit other materials.94-95 Au-containing MNPs (mostly iron oxide-based) (Au@MNPs) have attracted particular interest in the electroanalytical chemistry for bioseparation, the fabrication of immunoassays and the development of optical and electrochemical sensors due to the superparamagnetic properties of magnetic materials and the biofavorable (i.e., optical and electrical) behaviour of 96-97 Au. Different synthetic approaches have been reported for monodispersed Au-FexOy nanohybrids with diverse morphologies such as core/satellite, core/shell, multilayer, Au-coated iron oxide, and flower-like structures (Figure 2.1.3).98 In recent past decades, a significant number of Au-FexOy nanocomposites have been reported for the development of different biosensing tools such as glucose sensors and aptasensors.99 For instance, Chin et al. (2009) synthesized monodispersed Au and silver (Ag)–coated superparamagnetic Fe3O4 core-shell

NPs via seed-mediated growth. A thin layer of 2–3 nm Au and Ag NPs were attached to –NH2 3+ + 100 functionalized superparamagnetic Fe3O4 through the reduction of Au and Ag . Very recently, we developed an electrocatalytically active gold-loaded nanoporous superparamagnetic nanocube (Au@NPFe2O3NC) in which 2% Au NPs were loaded onto the porous cubic Fe2O3 nanocubes via the reduction of HAuCl4. A porous Fe2O3 nanocube was prepared by the calcination of Prussian blue (PB) nanocubes followed by the thermal decomposition of that porous PB powder.45

Figure 2.1.3: Schematic representation of the conversions of selected multilayer FexOy@Au composites. Reproduced with permission.98

Carbon nanomaterials such as graphene, graphene oxide (GO), and carbon nanotubes (CNTs) have also played a significant role in biomolecular analysis. For example, two- dimensional graphene (or GO) possesses excellent surface-bound properties such as a large surface area (up to 2630 m2/g), unique sp2/sp3 bonded structure, thermal conductivity, and high carrier mobility. They (carbon NMs) demonstrate a different binding affinity toward the double-stranded and single-stranded DNA and thereby are used in designing a sensor to detect DNA or a DNA-based marker by discriminating between different DNA sequences. Inspiring form their intrinsic properties, several graphene-containing superparamagnetic iron oxide nanocomposites have been synthesized. A one-step synthesis of Fe3O4 NPs decorated with reduced graphene oxide was reported by Teymourian et al. for the sensing of various analytes 101 (e.g., NADH, H2O2, uric acid, nitrite, ascorbic acid, and dopamine). A bifunctional Fe3O4- 102 Pt/rGO has also been reported, wherein Fe3O4 and Pt NPs were coated onto the rGO surface. This nanocomposite had been used for the catalytic reduction of methylene blue and the aerobic oxidation of benzyl alcohol. Very recently, a new composite of GO sheets and PB consisting of a different ratio of GO and PB has been reported by Tanaka et al.103 They synthesized nanoporous GO/iron oxide (IO) hybrid composites via thermal decomposition of GO- sheets/PB composites in the air at 400 °C. Among all ratios (GO:PB ratio = 25:75, 50:50, and 75:25), the 25:75 ratio resulted in a higher surface area (120 m2 g1) than those of the pure GO (34.9 m2 g1) and IO (93.1 m2 g1) samples. In addition to GO, several multiwalled CNT- containing IO nanohybrids also possessed promising applications for catalysis and biosensing

due to their combined functionalities, such as the high chemical stability and electrical conductivity of cylindrical graphene sheets and the superparamagnetic properties of IO.54, 104

2.1.2. Surface functionalization of superparamagnetic nanoparticles The ability to tune the surface in a controllable manner and at a particular molecular level makes possible the use of MNPs in biomedical and biotechnological applications. Generally, MNPs have a large surface-to-volume ratio (10–50 m2g-1) and thus possess surface energy. In order to minimize the surface energy, naked IONPs tend to aggregate via magnetic interactions, limiting their dispersion in solution and complex matrices.105-106 Moreover, naked IONPs have high chemical activity and are easily oxidized in the air; they are unstable in strongly acidic solutions and undergo leaching, which actively reduces their magnetism, reusability, and lifetime. Therefore, surface functionalization or covering of the surface with biocompatible compounds is crucial to (i) prevent degradation in an aqueous, acidic, or aggressive (blood) environment; (ii) suppress the magnetic interactions to avoid aggregation; and (iii) provide a functional group to bind or attach a wide range of biomolecules (e.g., protein and nucleic acids). These strategies include coating with biofavorable inorganic materials; immobilization with small organic molecules, surfactants, polymers, or biomolecules; and functionalization with metal and inorganic substances such as silica, graphene, gold, or platinum.

2.1.2.1. Small organic molecules, functional groups, and surfactants Organic molecule–functionalized superparamagnetic NPs have been used in various applications, especially in the field of biomedicine for targeted drug delivery, magnetic cell separation and isolation, designing immunoassays, etc.66 To prevent particle aggregation and preserve good biocompatibility, IONPs are generally functionalized using different organic materials (e.g., polyethylene glycol (PEG), dextran, and starch) and various functional groups 106 such as –OH, -COOH, -NH2, and –SH. These functionalized groups are also suitable for the further addition of different bioactive molecules for targeted bioapplication. For example, small-sized silane is used to modify the end groups of the IONP surfaces for conjugating NPs with polymer or other metal ions, biomolecules, or biological entities.107 p- Aminophenyltrimethoxysilane (APTS), mercaptopropyltriethoxysilane (MPTES) and 3- aminopropyltriethoxysilane (APTES) agents are the most commonly used silanes for fastening the -NH2 and –SH groups. For instance, Shen et al. reported a synthetic approach of APTS- coated magnetic IONPs (Fe3O4@APTS), where Fe3O4 NPs (mean diameter 6.5 nm) were synthesized in the presence of APTS via the hydrothermal synthetic route to obtain an amine 29

functional group on the IO surfaces.108 The organic compounds oleic acid and oleylamine, which consist of a C18 tail with a cis-double bond, play essential roles in the adequate stabilization of IONPs via high-temperature thermal decomposition, as they can form a stable, protective monolayer around the synthesized IO.59 For instance, oleic acid (OA) has been used to coat Fe3O4 during the synthesis via thermal decomposition of Fe(acac)3. The resulting OA- coated NPs had an OA coating 3 nm thick and possessed good superparamagnetic properties (magnetic saturation value 78.68 emu/g).109 Moreover, the IONPs obtained from the organic iron precursor are capped with nonpolar groups, and they become stable in organic solvents such as hexane. These hydrophobic NPs generally are not suitable for biological applications. To make them biocompatible, the hydrophobic organic phase needs to be replaced or transferred by the aqueous phase. To prepare water-soluble MNPs directly, small molecules such as amino acids, cyclodextrin, or citric acid need to be used in the reaction process.110-111 For example, Gao et al. synthesized hydrophilic superparamagnetic colloidal nanocrystals using an anionic polyelectrolyte PSSMA (4-styrenesulfonic acid-co-maleic acid) sodium salt containing both sulfonate and carboxylate groups as the stabilizer. The synthesis was achieved by a one-step solvothermal method. The nanocrystals were well dispersed in an aqueous solvent such as water, PBS buffer, or ethanol.112 In addition to these approaches, ligand-exchange transformation is used to convert oil-soluble functionalized iron oxide NPs to a water-soluble one. This method involves the addition of an excess amount of hydrophilic ligands into the nanoparticle solution, resulting in a displacement of the original ligands on the NP surface. A ligand-exchange method was reported for the synthesis of monodispersed water-soluble MNPs where oleic groups present (initially) on the NP surfaces were replaced by various capping agents bearing reactive hydroxyl moieties via ligand-exchange reactions.113 These hydroxyl groups could be exploited to initiate ring-opening polymerization of polylactic acid from the nanoparticle surfaces and esterified by acylation to permit the addition of alkyl halide moieties (Figure 2.1.4). Various surfactants, such as dodecyl amine, sodium oleate, and sodium- carboxymethyl cellulose, were also used to enhance the dispersibility of MNPs in aqueous media.114

Figure 2.1.4: Schematic representation of the functionalization of magnetic nanoparticles. Steps 1A and 1B: ligand-exchange reactions. Step 2: acylation of hydroxyl groups to prepare ATRP surface initiators. Step 3A: surface-initiated ring-opening polymerization of L-lactide. Step 3B: surface-initiated ATRP. Step 4: deprotection or additional reaction after polymerization. Step 5: grafting of end-functionalized PEG chains onto the nanoparticle surface using amidation chemistry. Reproduced with permission.113

2.1.2.2. Polymers Polymer-functionalized MNPs have drawn much attention due to several advantages of polymer coating, such as that polymer coating increases the pharmacokinetics and biodistribution of NPs and increases the Van der Waals attractive and repulsive forces to balance the magnetic forces on the NPs.59, 106 Several natural and biodegradable synthetic polymers, such as polysaccharides, polyaspartate, polyethylene glycol (PEG), poly(vinyl alcohol), and poly(vinyl-pyrrolidone), are currently used to produce MNPs with tailored and desired properties.59, 66, 68 There are two main purposes for the polymer coating of NPs: one is to broaden the application of NPs by introducing different functional groups, and the other is to manufacture monodisperse NPs with controlled composition and well-defined shape. However, after polymerization, the superparamagnetic properties of NPs have been decreased.

The conventional approaches to the polymer functionalization of MNPs are in-situ and post-synthesis coating. In the in-situ strategy, the NPs can be functionalized with polymers through mini- or microemulsion polymerization or the sol-gel process during the synthesis of NPs.115 Here, the organic precursor molecules capped the IONPs and formed an overlying layer through emulsion polymerization. The conventional structure obtained by this approach is mainly a matrix-dispersed structure or a core-shell structure.116 Unfortunately, this method failed to maintain the colloidal stability and thickness of the shell. The predominant method of the polymer coating is post-synthesis functionalization, where the polymer functionalization has been carried out on the prepared IONPs via the one-pot method, self-assembly, or heterogeneous polymerizations. The resulting polymer@NPs is prone to form a core-shell structure. Besides, a number of heterogeneous polymerizations with different water-soluble monomers have been used to prepare a well-defined monodisperse core-shell structure.117 An example of such polymerization was reported by Pimpha et al., where an all-in-one NP platform was developed using an oil-in-water emulsion system. The emulsion consisted of an iron oxide nanocrystal containing a hydrophobic oil core. The oil droplets were successively stabilized by a lipid mixture containing a near-infrared (NIR) fluorophore (Cy5.5), followed by further modification with PEGylated lipids to increase the stability of the particles. The lipid mixture favours the creation of small particles of 30–100 nm.118 Currently, MNPs are modified with smart polymers, which provide stimulus-responsive characteristics such as pH, temperature, and light for a wide variety of biomedical applications such as drug delivery, MRI, biomimetics, and biosensors.119-121

2.1.2.3. Bioactive molecules In recent years, biomolecule-functionalized MNPs have drawn attention in nanomedicine due to their biocompatibility and diverse application in the separation of biomolecules from complex biomatrix, detection, sensor-development, and other bioapplications. Numerous small biomolecules such as proteins, antibodies, enzymes, human/bovine serum albumin, avidin, and peptides have bound onto the surface of NPs.122-125 The bioactive molecules can be attached to the surface of NPs, mostly by activating the functional groups. In this strategy, small particles or polymer-functionalized NPs are synthesized, followed by the addition of biomolecules through physical adsorption or chemical bond. For example, Lee et al. reported an approach to conjugating the IONPs with single-strand oligonucleotides. In this report, water-soluble carboxyl group-containing magnetic γ-Fe2O3 was prepared, followed by modification of the

NP surface with streptavidin. Streptavidin-functionalized γ-Fe2O3 NPs were then used to bind 32

biotin-labelled oligonucleotides via the strong affinity interactions between avidin (streptavidin) and biotin.126 The reactivity of the NP surface–linked carboxyl group can be modified by the reaction with thionyl chloride (SOCl2) followed by coupling with the hydroxyl group-containing small molecules.127 As this approach has been carried out under anhydrous conditions (e.g., in DMSO or the presence of AlCl3), it is not suitable for immobilizing antibodies or enzymes. EDC (1-ethyl-3-(3-dimethylaminopropyl)carbodiimide) and NHS (N- hydroxysuccinimide) can be used instead of anhydrous reagents to make the carboxylated NPs suitable for amine group or enzyme immobilizations.128 In this modification, the aggregation of NPs via cross-linking of the particles also happened; therefore, it is not ideal for immobilizing proteins. Proteins can be immobilized on the surface of NPs with thiol functional groups. The protein–thiol bonds are highly selective, and thiol groups reduce the probability of NP aggregation.129 Another way to achieve chemical modification of NPs with proteins, antibodies, or enzymes is via the attachment of bifunctional aldehyde groups (e.g., glutaraldehyde) to their premodified amine groups through the Schiff-base condensation reactions.130

2.1.2.4. Inorganic materials Inorganic materials can possess numerous outstanding properties, such as high electron density and strong optical properties (e.g., Au and Ag), photoluminescence (e.g., CdSe, CdTe, and Y2O3), magnetic moment (e.g., cobalt and manganese oxides), and affinity interaction with biomolecules (e.g., Au, GO, and Pt).30, 131-135 Inorganic metal functionalization greatly enhances the antioxidation properties of unmodified NPs (e.g., iron oxides). Moreover, this coating extends the optoelectronic, storage, biocompatibility, catalytic, and sensing properties, and thereby, inorganic material–functionalized magnetic materials become very promising for application in catalysis, bioseparation, bio-labelling, biosensing (optical and electrochemical), and so on. Different inorganic materials such as silica, Au, carbon, GO, and metal oxides have been used to functionalize NPs, especially IONPs in nanomedicine.

Silica-coated MNPs such as IONPs (IONP@SiO2) are promising and widely used nanocomposite materials for biological applications. They possess several advantages: (i) silica coating enhances the dispersion of NPs in solution and complex biological samples, as silica coating can screen the interparticle interactions; (ii) they have good biocompatibility, stability, and hydrophilicity; (iii) the variation of silica shell thickness is relatively easy and straightforward; and (iv) silica coating enables the binding of biomolecules or other biofavorable ligands for biological applications.136 In general, three strategies have been used 33

to prepare IONP@SiO2 nanocomposites: the Stöber process, microemulsion synthesis, and aerosol pyrolysis. The Stöber process is the commonly used method for silica coating. In this method, IONPs are dispersed in alcohols, followed by the addition of silane. Water or ammonia solution is then added to form IONP@SiO2. The thickness of silica can easily be tuned from 5 to 200 nm by varying the concentration of ammonia and the ratio of silica precursors (tetraethoxysilane, TEOS).137 For example, Xuan et al. synthesized a monodisperse 138 Fe2O3@mesoSiO2 as a bifunctional agent for application in drug carriers and MRI. Micelles or inverse micelles have been used in the microemulsion process to obtain a confined and controlled coating of silica on core IONPs.139 This process requires the additional separation of core-shell NPs from a large number of surfactants. In aerosol pyrolysis, IONP@SiO2 is obtained by phase segregation of the iron precursors (bulk iron) and silica oxide or alkoxides in flame environments.140 Carbon-coated NPs (e.g., IONP@C) have also attracted enormous interest in bioapplications due to their elevated intrinsic electrical conductivity and excellent chemical and thermal stability. Carbon protects the NPs from oxidation and corrosion of the core materials. In general, a carbon coating can be applied using a simple three-step process. MNPs are first prepared using conventional synthetic approaches. MNPs generally act as seeds. Then the selective polymer is coated via a polymerization process. Finally, the IONP@C composite is generated by the annealing treatments.66 Recently, much attention has been drawn to the functionalization of magnetic materials by using highly conductive GOs. IONP/GO hybrid materials have been used in biological fields; for example, Chen et al. synthesized an amino- 141 dextran coated-Fe3O4/GO composite for cellular MRI. Different metallic NPs (e.g., Au, Ag, Pt, Cu, and Co) possess different properties, such as optical (surface plasmon resonance, light scattering, surface-enhanced Raman scattering), electrical (conductivity), and catalytic ones, and are widely used in contrast imaging, catalysis, sensing, and medicine.142-143 The combination of such metals with MNPs significantly enhances the functional properties of metal/NP composite via cooperative interaction of the intrinsic properties of metal and metal oxide NPs. Among all metal-coated NPs, Au-coated IONPs have been intensively studied and showed enormous potential in biotechnology and biomedicine.143 More specifically, the planar gold surfaces or gold nanoparticles (AuNPs) hold significant advantages for the electrochemical and optical detection of biomolecules, as they have a selective and specific affinity toward DNA, RNA, antibodies, and proteins.133 However, the Au coating on MNPs is not thick enough and may cause aggregation. Hence, ionic capping ligands must be attached to the system during NP synthesis. Moreover, Au-coated MNPs 34

(Au@MNPs) can be modified with a variety of molecular linkers to bind the target analyte for designing optical and electrochemical biosensors. For example, MNP@Au can be adjusted with 1,9-nonanedithiol and mercaptoundecanoic acid (MUA) through the ligand-exchange reactivity at the gold shells to form a thin film assembled core (iron oxide)–shell (gold) nanocomposite. The assembly of Fe3O4@Au nanoparticles involved an initial thiolate– oleylamine exchange reaction followed by cross-linking. The cross-linking was carried out by using the alkyl chain (dithiol as linker A) or hydrogen bonding (carboxylic acid-functionalized thiol as linker B) as shown in Figure 2.1.5.144

Figure 2.1.5: Schematic representation of the thiolate–oleylamine exchange reaction followed by cross-linking via different linkers (A and B) for the assembly of Fe3O4@Au NPs. Fe3O4 core: dark circle; Au shell: grey circle; oleylamine monolayer: thin grey zigzag lines. Reproduced with permission.144

Various metal oxides and metal sulphides, such as ZnO, TiO2, Al2O3, CuO, CdS, ZnS, and PbS, have also been used to modify the surface of MNPs for a wide variety of biological technologies, as they add unique and selective properties to nanocomposites.145-148 For example, Li et al. prepared air-stable Fe3O4@Al2O3 core-shell NPs through displacement reactions. The NPs possessed high magnetic responses and were used to capture a MALDI 149 target. In another example, thiol-modified superparamagnetic γ-Fe2O3 beads were modified by CdSe@ZnS quantum dot NPs to form luminescent/magnetic nanocomposite particles via thiol–metal conjugations. The composites showed a threefold higher emission quantum yield, and their emission peaks showed a slight blue shift as compared with individual luminescent QDs. This luminescent/magnetic nanocomposite was used for cell separation, magnetic separation, and luminescent detection.150

2.1.3. The function of superparamagnetic nanoparticles in biosensor development Superparamagnetic materials are traditionally used for the concentration, magnetic separation, isolation and purification, and identification of molecules and cells. Recently, the magnetic properties of superparamagnetic particles have also been used as detection and signal- amplifying tools in various biosensing platforms. The electrocatalytic activity is largely utilized to enhance detection sensitivity and specificity. Superparamagnetic materials are also used as nanocarriers to transfer different biomolecules or bring target biomolecules to a biosensing system. Additionally, it has been discovered that several superparamagnetic NPs, especially magnetic ferric oxide–containing NPs, exhibited intrinsic natural enzyme-like activity.

2.1.3.1. Magnetic capture and separations Superparamagnetic nanomaterials are becoming a promising tool for the capture, concentration, isolation, and separation of biomolecules such as in vitro cells, antibodies, proteins, DNA, enzymes, bacteria, and different pathogens from their complex biological matrixes.29, 66, 151 MNPs offer several significant advantages over conventional separation techniques such as chromatography: they possess a large surface-to-volume ratio, are readily dispersible in solution, can be quickly localized or retrieved using a typical external magnet, and provide high versatility and reusability of NPs after the magnetic separation.152 Generally, MNPs are functionalized with different surfactants, polymers, and ligands to introduce functional end groups such as –OH, -NH2, -COOH, and –SH for the selective capturing of target biomolecules. The porous structure of polymer beads containing a magnetic core is also employed to capture size-induced biomolecules. The separation of target biomolecules can be achieved via hydrophobic interactions, antibody-antigen interactions, or direct affinity adsorptions. Two factors greatly influence the separation efficiency of NPs: (i) the amount of active functional group per mass of magnetic materials and (ii) the saturation magnetization value.153 A large amount of functional group and high saturation magnetisation value favours the efficient separation of biomolecules. Magnetic microsphere or composite materials fulfil these two requirements, as they contain a higher amount of magnetic materials, a porous polymer structure, and selective metal structure for affinity interaction with target biomolecules. NPs with a porous structure can also be good candidates, as they have an enormous surface area and the ability to incorporate a larger number of functional groups or biomolecules. A schematic representation of the magnetic separation of biomolecules is shown in Figure 2.1.6.

Figure 2.1.6: Schematic representation of superparamagnetic NP–based bioseparation of biomolecules. Reproduced with permission.154

Recently, Min et al. isolated and purified genomic DNA from human blood using monodisperse biofunctionalized superparamagnetic nanoparticles.155 The NPs were functionalized with meso-2,3-dimercaptosuccinic acid (DMSA), which contains both carboxylate and thiol groups for DNA absorption. Small quantities of resulting DMSA-MNPs can isolate DNA from blood samples with a maximum yield of 86.16%, which was shown to be much better than that with a commercial microbead (NucliSENS-easyMAG, BioMérieux).

Colloidal IO-based non-porous magnetic NPs was also reported to be used in the isolation of DNA from chicken erythrocyte and Bifidobacterium longum bacterial strain. The isolated DNA was tested and quantified by PCR amplification and used for the identification of Bifidobacterium bacterial strains.156 In the case of proteins and peptides, the magnetic particles can couple with an appropriate affinity ligand or biopolymers to exhibit the affinity interaction toward the target. The particles can then be added to the sample matrix to bind with the target. In another approach, the respective antibody has been used as a free affinity ligand and added to the samples to unite with the antibody-specific target proteins or peptides. Magnetic particles are

then functionalized with a secondary antibody (e.g., protein A or protein G) to facilitate interaction with the target–antibody complex.157 For instance, Wittrup et al. performed an antibody-conjugated MNP–based isolation of heparan sulphate proteoglycan (HSPG)–induced endocytic vesicles to understand the role of the GRP75 protein in the HSPG-mediated endocytosis of macromolecules.158 The extracellular vesicles and exosome can also be isolated using an antibody–MNP complex following a method similar to that for proteins. Superparamagnetic nanoparticles are also widely used in the isolation of mammalian and bacterial cells from complex biomatrixes. Specific cell-recognition moiety-functionalized NPs are incubated with the cells, followed by separation using an external magnet.29 For example, anti-CD4 antibody–functionalized IONPs were used to isolate and separate the CD4+ T cells from human blood samples. The anti-CD4 antibody has a specific affinity to CD4+ T cells. Thus, anti-CD4 IONPs precisely capture the T cells. The cells were then captured with a magnet, and cell-purification efficiency was tested by using mass spectrometry through assessment of the T cell peptide mass fingerprints.159 Cell-specific ligands (e.g., oligonucleotides or peptides) such as aptamers are also used in the recognition and isolation of cancer cells using MNPs. Aptamer (specific sequence for acute leukemia cells)-modified MNPs were reported to develop a rapid collection and detection platform for acute leukemia cells from whole blood samples. In this strategy, the aptamer was modified with iron oxide– doped NPs and used to extract the target cells, while aptamer-tagged fluorescent NPs were used to detect the target cells.160 The separation of apoptotic cells can also be achieved by using streptavidin-modified MNPs. The apoptotic cells express the phosphatidylserine receptors (not revealed by a healthy cell), which can specifically bind to annexin V.161 Thus biotinylated- annexin V-labeled apoptotic cells can bind to streptavidin-modified MNPs via the accessible avidin-biotin interaction. This MNP–cell complex can be easily separated and collected using a magnet before visualization of the cells.162 Though MNP-based isolation and preparation has many advantages (e.g., being inexpensive, sensitive, and reusable), the method is relatively time-consuming due to the multiple steps involved in the isolation and separation process. However, the processing time can be reduced by integrating the isolation and separation method into a microfluidic device using free-flow magnetophoresis.163 Chen et al. have developed a POC viral assay to purify and concentrate whole-particle HIV-1 (human immunodeficiency virus) from plasma samples by using a microfluidic magnetic separator chip. In this method, the anti-CD44 antibody was conjugated with a superparamagnetic NPs. Virus-containing plasma was then mixed with antibody MNPs to obtain an HIV–MNP conjugate. The conjugate was then passed through a packed bed of IO particles, and an external 38

magnet was applied to magnetize the bed, which trapped the HIV–MNP conjugates, thereby separating the viral protein with a 62% extraction efficiency and 80-fold concentration.164 In addition to biomolecules, MNPs can be used to absorb heavy metals such as arsenic from the aqueous environment, and even bare MNPs (without surface coverage) can be used to separate pollutants from wastewater via nonspecific interactions.165 Table 2.1 summarizes the potential magnetic nanoparticles used for magnetic isolation of target biomolecules from complex biomatrix.

2.1.3.2. Development of a detection platform (sensor and biosensor) The application of superparamagnetic NPs to the development of a sensing and detection platform is highly advantageous, as most of the biological species are nonmagnetic (they possess inherited low background noise), and their size is analogous to that of the biomolecular targets (e.g., DNA, protein, cells, and exosomes).28 Therefore, MNPs can be easily integrated with biomolecules that have a lower steric hindrance, and such an interaction can be easily controlled by controlling the NP morphology and sensor fabrication steps. Several magnetic metal NPs, metal oxide NPs, nanowires, and composite materials have been reported to have the potential for use in a wide variety of biosensors for a broad range of bioanalytical applications.41, 166 MNPs with a different size, nature, morphology, and composition can play significant roles in diverse sensing platforms. Figure 2.1.7 schematically represents the critical function of MNPs that can be achieved when they are integrated into a sensor platform, and the details of each feature will be discussed in the following sections.

Figure 2.1.7: The function of superparamagnetic nanomaterials in a biomolecule detection platform

Table 2.1. Magnetic nanostructures used for magnetic isolation of target biomolecules.

Magnetic NPs Biofunctional Group Function Target Ref.

Magnetic isolation of CTCs using MNP- Fe3O4 EpCAM MCF-7 and MDA-MB-231 cells 351 EpCAM-CTC interaction Selective capture and magnetic enrichment of Fe3O4NPs AMP E. coli, S. aureus and P. aeruginosa 352 bacteria

Anti-S. agalactiae Immunomagnetic optical probe witkwah both MNPs antibody through EDC and magnetic and fluorescent properties S. agalactiae 353 sulfo-NHS coupling for separation and enrichment of bacteria.

Antibody (Ab2) through Isolation of biomarker protein and peroxidase- Fe3O4@GO PSA and PSMA 354 EDC-NHS coupling mimetic for electrochemical detection.

Gold-coated magnetic DNA aptamers Isolating bacterial strain from waste water Escherichia coli cells 355 microdiscs Polymeric magnetic (iron CSA-1-Ab antigen through Isolation of target bacteria and enrichment of Salmonella 356 oxide) NPs diketonic functionalities SERS probe Direct interaction of Ti and Fe3O4@TiO2 Exosome enrichment and separation Circling exosomal PD-L1 357 Phospholipids of antibody. Dynabeads MyOne Streptavidin Magnetic separation and purification DNA, RNA, Protein 57,81,238,301

AuNP-Fe2O3NC Au-protein interaction Isolation of exosome Exosome 310 Isolation and magnetic enrichment of AuNP-Fe2O3NC Au-protein interaction p53 autoantibody 207 autoantibody

2.1.3.2.1. Immobilization of biomolecules Immobilization of biomolecules has been considered a critical step in biosensor design, as the direct adsorption or attachment of biomolecules onto bulk materials results in the denaturation or decrease of their activity.167 Therefore, traditional approaches to the immobilization of biomolecules in sensing platforms have been dramatically improved in recent years by the inclusion of magnetic and superparamagnetic NPs. Nanomaterials offer a larger surface area for enhanced biomolecule loading, have a greater ability to adsorb biomolecules and reduce the diffusion limit of biomolecules on the transducer surface.168 The prerequisite requirement for biomolecule (e.g., protein or enzyme) immobilization is that the particles should provide an inert and biocompatible environment so that NPs cannot interfere with the native structure and function of target species.169 Therefore, in recent decades, we have witnessed a significant number of studies on designing bioactive nanomaterials and their application as an immobilization platform in biosensing.170-172 Among all materials, superparamagnetic NPs have been used extensively to immobilize and capture biomolecules or analytes (direct attachment) on a transducer surface.173-175 In addition to the high surface area, high binding capacity, and high catalytic specificity of superparamagnetic materials, MNPs provide for the easy capture of biomolecule-modified NPs on the transducer surface. There are four principal methodologies to link biomolecules (proteins or enzymes) with NPs. As shown in Figure 2.1.8, these are electrostatic adsorption, covalent attachment, affinity-based immobilization, and direct conjugation to the NP surface.176

Figure 2.1.8: General strategies for the immobilization of MNPs with biomolecules.

The electrostatic interaction (through the intrinsic charge of NPs and biomolecules) approach is widely used, and such an interaction between NPs and proteins can be easily modulated by the ionic strength and pH of the medium. Covalent attachment (NPs modified by a primary amine or carboxylic groups) is another frequently used method of NP– protein/enzyme conjugation, and the conjugation is greatly improved by controlling the surface chemistry of the NPs and reaction media. For instance, a ferric oxide nanomaterial can be coated with a polymer (e.g., PEKY), which facilitates the covalent attachment to a monoclonal antibody.177 It can also be coated with dextran to bind peptide sequences.178 In addition to NP surface coating, the NP surface can be modified with thiol chemistry followed by the covalent addition of proteins, enzymes, or antibodies.179 Wang and Lee proposed a method for the direct binding of proteins onto superparamagnetic Fe3O4 through carbodiimide chemistry. They activated the Fe3O4 surface with two kinds of carbodiimide, cyanamide and N-ethyl-Nˊ-(3- dimethylamino-propyl)carbodiimide (EDC), followed by immobilization by two model proteins, trypsin and avidin. The immobilization was increased by increasing the molar ratio of the EDC/NPs, and NP activation with EDC gave a higher yield of immobilization than cyanamide did. The avidin-bound NPs were bound to a biotinylated ssDNA and hybridized with a DNA probe to form a DNA sensor.180 The avidin (streptavidin)–biotin interactions are

the most robust noncovalent affinity interactions and using this specific affinity; avidin coated NPs can selectively bind with biotinylated proteins or disease-specific antibody modified NPs can bind with the selective target proteins.181 Protein can also be directly conjugated with the NPs using a linker, and such conjugation is desirable when NPs are used in the preparation of sensors involving electron transfer. Composite nanomaterials containing Au or Ag are usually used to bind biomolecules in this approach. For instance, Au or Ag can be bound to a protein using a cysteine residue of proteins through Au–thiol/Ag–thiol chemistry. Lately, biomolecule (e.g., proteins, DNA, RNA)–gold direct adsorption (the affinity interaction of DNA/RNA/proteins with gold) has also been considered a more promising method to adsorb biomolecules over unmodified gold NPs or gold surfaces.20, 133 Though a number of significant studies have been carried out and streptavidin-modified MNPs (e.g., magnetic beads or protein- G magnetic beads) are commercially available, the application of magnetic beads in a sensor platform suffers from a lack of reproducibility and renewal of transducer surfaces. In this regard, nanoporous superparamagnetic materials containing different bioactive materials could be a possible solution, as they will not require complex chemistry and would have a large surface-to-volume ratio, strong magnetic affinity, and reusable surface morphology.

2.1.3.2.2. Electrocatalytic signal amplification With the growing demand for the trace amount of biomolecule detection, several signal- amplification strategies have been developed. These strategies include the polymerase chain reaction, mass spectrophotometry, label-amplified signal enhancement (electroactive molecules, different redox molecules and metal ions) and the integration of enzyme-mediated signal-amplification strategies.182-183 Although these methods offer adequate sensitive detection, they are time-consuming, destructive, depend on costly detection approaches, and require highly skilled professionals. Integration of the signal-amplification properties of NMs into the sensing platform opens the door to ultrasensitive and selective detection due to their inherent catalytic and conductive properties. The NMs generally act as a catalyst to trigger enhancement of the detectable signal. NP electrocatalysis is highly adaptable and can be readily scaled for the ultralow amount of biomolecule sensing. The biocompatible NMs can produce a synergistic effect between the conductivity and catalytic activity to accelerate the signal transduction events, resulting in the lowering of detection limits even at zeptomolar levels.184- 185 Moreover, the accelerated signal-transduction events broaden the gap between the two successive concentrations in a range of 5–6 orders of magnitude. For instance, superparamagnetic NPs based on a highly sensitive, ultralow protein detection approach has 43

been reported by Krishnan et al. In this method, antibody-labelled superparamagnetic NMs were used for signal amplification in a surface plasmon resonance (SPR) immunoassay for the detection of a cancer biomarker, a prostate-specific antigen (PSA) in the serum. A monoclonal antibody, ab2, was first conjugated with superparamagnetic particles (MP) (Dynabeads, Invitrogen), followed by the formation of MP–Ab2–PSA particles. As can be seen in Figure 2.1.9, the MP–Ab2–PSA bioconjugates were then applied to a PSA-specific antibody (ab2) immobilized Au-SPR chips. The SPR signal was monitored constantly after the addition of MP–Ab2–PSA bioconjugates and the SPR signal quantify a and ultralow amount of PSA with a detection limit of 10 fg mL-1 (ca. 300 aM).186

Figure 2.1.9: Schematic representation of an SPR immunosensor based on clustered magnetic NMs for signal amplification. Reproduced with permission.186

In constructing electrochemical sensors, one of the crucial requirements is electrical contact between the electrode surface and redox biomolecules. Generally, the active sites of biomolecules are surrounded by a thick protein shell, which blocks the direct electron transfer between biomolecules and the electrode surface. Metal NPs can resolve the barrier and enhance the rate of electron transfer due to their high conductive properties. In those circumstances, MNPs could be used for loading a higher amount of signalling tracer and controlling them magnetically for achieving an ultrasensitive detection. For example, iron oxide-based NPs were employed as a signal enhancing element for the electrochemical immuno-sensing of carcinoembryonic antigen (CEA). The three-layer MNP Au–PB–Fe3O4 (Fe3O4 magnetic core, PB as an interlayer, and gold as a shell) was functionalized with a bioenzyme (HRP and glucose oxidase), and it exhibited excellent redox electrochemical activity and superior enzyme catalysis activity toward glucose in an immunoassay of CEA and α-fetoprotein (AFP). The

sensor can detect 4 pgmL-1 and 7 pgmL-1 of CEA and AFP, respectively. An external magnet could be used to isolate NPs, with subsequent magnetic purification to make the sensor reproducible and reusable.187 A similar sandwich-type immunoassay has been reported for ultrasensitive detection of the carbohydrate antigen (CA-125), and AFP based on the magnetic NMs as a signal amplifier.188 Silica NPs and silica-coated magnetic materials can also be used as carriers for signal amplification in the development of an ultrasensitive immunoassay due to the small size, high surface-to-volume ratio, and excellent biocompatibility of silica.189-190

2.1.3.2.3. Signal-generating probes The labelling of biomolecules in the construction of biosensor is a common strategy for obtaining output signals. In the most conventional approaches, different enzymes, redox molecules, electroactive molecules, and metal ions have been used as a signal generation label.191 The natural enzymes possessed poor stability under environmental stress, and sensor design had a high cost. The use of redox molecules as a label is a popular approach, but it has been limiting their application, as it can only transfer a few electrons (or even just one), which directly affects the sensitivity of the biosensor. The integration of biofunctionalized NMs as a label to obtain a signal, especially an electrochemical signal, has opened new prospects in biosensing. The NM label generated an enormous signal, which is associated with the ultrasensitive detection of an ultralow number of biomolecules. The biomolecule generally attached to or modified to the biocompatible NMs, and hence retained their activity to interact with its counterpart. There are two strategies by which NMs generate signals: (i) they load an increasingly large number of target biomolecules, and (ii) they act as an ultramicroelectrode array for the electrolysis of a bulk amount of substrates.166 NMs generally contain a great many electrochemically active elements (or atoms), thereby increasing the loading of electroactive species onto the transducer surface and enhancing the sensitivity of the readout system. For instance, a flow injection chemiluminescence (FI-CL) assay for the detection of DNA hybridization was developed based on a biobarcode-functionalized MNP label. As can be seen from Figure 2.1.10, MNPs are functionalized with amino-modified probe DNA (pDNA) and biobarcode DNA (bbcDNA) to form bbc-p-DNA-MNPs. The target DNA (tDNA) was captured on a capture DNA– assembled (cDNA) gold electrode surface. The bbc-p-DNA-MNPs were then allowed to hybridize with the overhang region of pDNA through pDNA–tDNA hybridization. The bbc-p-DNA-MNPs were dissolved in nitric acid solution, which releases a large number of ferric ions. Thus 3+ ultrasensitive detection of a DNA hybridization event was achieved by the luminol–H2O2–Fe CL free-radical reaction system.192 45

Figure 2.1.10: Schematic representation of a biobarcode-functionalized MNP label–based chemiluminescence detection of DNA hybridization (A); the flow injection chemiluminescence readout system for the quantification of Fe3+ (B). Reproduced with permission.192

Iron oxide and some other promising MNPs catalyze common chromogenic substances and efficiently enhance the detection signal in immunosorbent assays. The cascade reactions are highly sensitive to the pH of the detection buffer, temperature, and time, resulting in a time-consuming, dose-dependent reaction and poor reproducibility. The integration of MNPs as a signal-generation and amplification label in a colorimetric assay could give better and more detection, as the assay is independent of the cascade reaction as well as such reaction conditions. The released ions from MNPs generated an enhanced signal intensity, which is directly linked to the level of target biomolecules. Recently, Zhang et al. developed a colorimetric immunoassay for the sensitive detection of a cancer biomarker carcinoembryonic antigen (CEA) based on a ferric oxide MNP as a signal generator and amplifier label. Herein, an antibody (ab1) was loaded into a high-binding 96- well microplate, followed by the addition of a target CEA. An antirabbit IgG (Ab2)–functionalized MNP was then added to form a sandwich-like immunocomplex. The MNP, acting as an iron pool, released a large amount of Fe2+ when it was dissolved in a mixture of bathophenanthrolinedisulfonic acid disodium salt (BPT) and ascorbic acid (AA). The Fe2+ generated a stable red solution, which gave a naked-eye and quantitative evaluation through colorimetry results in the sensitive detection of CEA with a limit of detection (LOD) of 3.6 pg mL−1.193. Another exciting application of MNPs is to design novel magneto-switchable electrodes with unique properties. Various MNPs, nanorods, and nanosheets have been used to turn on or off the electrochemical reactions, depending on the

physical translocation and orientation of the MNPs in response to the direction of an external magnetic field.194-195 The electrochemical reaction is triggered by the reposition of an external magnet, resulting in a change in the magnetic field strength of the electrode interface. Willner et al. presented an alkyl-chain-functionalized MNP–based magneto-switchable hydrophilicity/hydrophobicity-controlled electrode surface. As can be seen from Figure 2.1.11, the two-phase platform consists of a gold electrode with an aqueous buffer solution and an organic (toluene) phase located on the electrode surface. Hydrophobic superparamagnetic

Fe3O4 NPs were used to regulate the interfacial properties of the electrode surfaces. When a magnet was placed below the electrode, the hydrophobic MNPs were pulled down from the upper toluene layer into the aqueous layer, resulting in the formation of a membrane-like film on the surface, thereby inhibiting the interfacial electron transfer to create an “off” state. The “on” state was achieved by placing the external magnet above the electrode surface, which pulled the MNPs up from the aqueous to the organic phase, allowing contact of the redox probe (e.g., ferrocene or ferrocyanide dicarboxylic acid) with the electrode surface and facilitating the oxidation process.196

Figure 2.1.11: Magneto-controlled reversible translocation of the functionalized MNPs between the organic phase above the aqueous electrolyte and the electrode surface. (A) A magnet below the electrode surface pulls the hydrophobic MNPs, forming a membrane-like layer on the electrode surface (“off” state); (B) a magnet positioned above the electrode returns the MNPs to the organic phase, resulting in the generation of an electrochemical reaction (“on” state). Reproduced with permission.196

A photoelectrochemical current can also be generated and magnetically controlled via magnet-bound quantum dots. With DNA hybridization, the biocatalytic reaction can be

controlled using capture probe–functionalized MNPs directed to and from the electrode surface.197-198 In the most refined magneto-switchable electrode systems, the activity of electrodes was regulated (activation or inactivation) by complex multi-enzyme systems or different biochemicals added to solutions, thereby mimicking the biological event and properties.199-200

2.1.3.3. Nanocarriers Due to their tremendous magnetic nature, biocompatibility, and high surface area, magnetic NMs are becoming excellent candidates for carrying agents in numerous biomedical applications, such as magnetic guided drug delivery, the separation of magnetically tagged DNA, the identification of biological species, magnetic resonance imaging, and cancer therapies.32, 201-202 NMs can also be used to support the isolation system for concentrating the target biomolecules or high-level loading of signal molecules. Generally, single-wall carbon nanotubes (SWNTs) have been used to anchor probe DNA in the transducer surface when constructing a hybridization-based sensor. For a DNA sensor, the target DNA was hybridized with the probe DNA. Based on the decreasing DPV response to the guanine base of DNA, the target DNA can be quantified.203 Magnetic NMs can easily concentrate and purify the target and hence dramatically enhance detection sensitivity. For instance, Wanunu et al. reported the nanopore-based electronic detection of microRNAs based on a protein-functionalized magnetic bead. In this assay, the target miRNA was first hybridized with a capture probe to form a probe- microRNA duplex. The duplex was allowed to bind with protein p19–functionalized magnetic beads. The magnetic beads authorized the ultrahigh enrichment of the duplex and purification of the duplex from the RNA pool. The suplex was then eluted and electronically detected by using a nanopore system. The magnetic bead produced 100-fold enrichment of the duplex and enabled pictogram-level detection of liver-specific miRNA from rat liver miRNA.204 Au- containing NMs have also been used as carriers for target or signalling molecules. For instance,

AuNPs-coated Fe3O4 NPs can serve as a reusable carrier for the immobilization of biomolecules (enzymes).205 Introducing pores into magnetic NMs can tremendously enhance the surface area and significantly improve the functionalization with biomolecules, drug loading, concentrating target molecules.206 Therefore, porous magnetic NMs containing AuNPs could provide a great platform as carriers of different biomolecules. Recently, Sharda et al. developed a new class of gold-loaded superparamagnetic NMs (Au@NPFe2O3) as a dispersible capture agent for the naked-eye and electrochemical detection of cancer-specific autoantibodies. As can be seen in Figure 2.1.12, the synthesized Au@NPFe2O3 nanocubes were 48

functionalized with the p53 protein, followed by the dispersion of p53 protein–functionalized NMs into the serum samples containing target p53–specific autoantibodies. The higher surface area of superparamagnetic NMs facilitates the bindings of a higher amount of target, and purification and isolation of target as well. The horseradish peroxidase (HRP)–modified IgG (secondary antibody) then bound with the target and was used to catalyze the oxidation of the

3,3,5,5’-tetramethylbenzidine (TMB)/H2O2 system. The generated color indicates the presence of a p53 autoantibody, and subsequently, the amount was quantified by colourimetry and amperometry.207 Similarly, different magnetic NMs are also utilized in microfluidics and lab- on-chip systems as a carrier of target molecules, target-binding proteins, or signalling molecules.

Figure 2.1.12: Schematic representation of the detection of p53 autoantibodies based on a superparamagnetic Au@NPFe2O3 nanocube as a dispersible nanocarrier. Reproduced with permission.207

2.1.3.4. Natural enzyme mimetics: nanozymes Nanozymes is a term referring to the NMs that exhibit natural enzyme-like activity to catalyze the oxidation reaction of various organic substances (e.g., chromogenic substances).208-209 Nanozymes have been considered an excellent alternative to natural enzymes in the fields of biosensor development, nanomedicine, and environmental remediation due to their much higher stability, easy storage, and lower cost.210-212 With the tremendous advancement in nano research and distinctive properties of NMs, a wide range of NMs (e.g., ferromagnetic NPs, gold NPs, rare earth NPs, metal complexes, and polymers) have been developed to exhibit

peroxidase-, oxidase-, catalase-, superoxide dismutase-, and lactase-like activities.213-215 Among these NMs, iron oxide-based MNPs are highly advantageous in bioanalysis due to their chemical and biological inertness, easy surface modification, and magnetic field–based separation efficiency. In 2007, Yan et al. discovered that, surprisingly, Fe3O4 NPs possess an intrinsic peroxidase-mimicking activity that catalyzes the oxidation of three chromogenic substances named TMB, DAB (di-azo-aminobenzene), and OPD (o-phenylenediamine).208 The experiment showed that the Michaelis–Menten constant (Km) value of MNPs with TMB was about four times lower than that of those with HRP, indicating that NPs have higher catalyzing activity for the oxidation of TMB. This is because HRP contains only one iron ion, in contrast 2+ 3+ to a large amount of Fe and Fe on the surface of NPs. However, the Km value of NPs with

H2O2 (as a substrate) was significantly higher than that of those with HRP, demonstrating that a higher amount of H2O2 was required to achieve the maximum mimicking activity. Like natural HRP, the activity is highly dependent on the pH and temperature of the reaction buffer. Moreover, the NP size influences the activity: the smaller the particle size, the more activity was observed, as smaller NPs possess a higher surface-to-volume ratio to interact with the substrates. The mimetic activities are generally performed by the generation of a hydroxyl free radical from H2O2 following the Fenton reaction (as shown in equations 1-3), and subsequent oxidation of TMB (equation 4) is carried out by the generated hydroxyl free radical.216-218

3 2  Fe  H 2O2  FeOOH  H (1)

2 2 FeOOH  Fe  HO2  (2)

2 3  Fe  H 2O2  Fe  OH OH (3)

OH TMB TMBoxBlue (4)

The reaction mechanism is considered to be the ping-pong mechanism, as no tertiary intermediate is formed between the NPs and the two substrates (the product from one substrate is dissociated before the second one binds). Moreover, the steady-state kinetic experiment showed that the substrate concentration-dependent double-reciprocal (Lineweaver–Burk) plot was parallel, which is characteristic of the ping-pong mechanism. Inspired by Yan’s discovery, many researchers have worked to develop a colorimetric sensing platform of H2O2, glucose, cysteine, etc., based on the MNP peroxidase mimetic activity using TMB and ABTS (2,2'-and-bis(3-ethylbenzothiazoline-6-sulphonic acid) as substrates. For instance, Wei et al. developed a colorimetric platform for the detection of glucose using Fe3O4 as a nanozyme and ABTS as a substrate. The oxidized ABTS generated a

green product, which enabled naked-eye detection. The colorimetric readout can detect H2O2 -5 219 as low as 3 mM and glucose as low as 3×10 mol/L. Later, H2O2 detection was used to detect several ions, glucose, etc. In addition to ABTS, other chromogenic substances, such as TMB, 4-AAA-phenol, OPD, and DPD, have also been used, and it has been shown that TMB gives more selective and sensitive detection than ABTS, as H2O2 can oxidize ABTS even in the 220 absence of peroxidases. Doped ferrites such as Prussian blue/Fe2O3, CoFe2O4, graphene/Fe2O3, ZnFe2O4, and CoF2O4 had also been explored as peroxidase mimetics for the 221-224 detection of H2O2, glucose, L-cysteine, etc. For instance, CL detection of H2O2 and glucose was performed using CoFe2O4 MNPs. The CoFe2O4 MNPs efficiently catalyzed the decomposition of H2O2 into hydroxyl free radicals (·OH), which subsequently catalyzed the oxidation of luminol. Coupling the NP-catalyzed CL reaction of luminol–H2O2 with the glucose oxidase–catalyzed glucose oxidation reaction enabled CL detection of glucose from blood samples with a detection limit of 0.024 µM.224 Apart from iron oxide-based NMs, other nanomaterials, such as AuNPs, CuONPs, Eu2O2SNPs, AgX (X= Cl, Br, I), polyoxometalates, PtNPs, carbon nanotubes, and nanodots, have been exhibited peroxidase-like activity.225-227 One of the major challenges of MNPs is the specificity of nanozymes, as NPs do not have a binding pocket (active sites). The catalytic reaction of nanozymes takes place at the surface; thus there is a possibility of substrate diffusion to the surface that can react with all substrates, irrespective of substrate shape and size. Many approaches have been utilized to introduce substrate specificity or a molecular-recognition function. These approaches include attaching aptamers, antibodies, peptides, or molecular imprinting. Among them, molecular imprinting has been considered superior to biological ligands, as biological substances are less stable, easily denatured, and expensive and require sophisticated operations. Molecular imprinting refers to the polymerization of a monomer (complement to the template) around template molecules to form a prepolymer binding complex. 228 Very recently, Zhang et al. grew molecularly imprinted polymers on Fe3O4 to form substrate binding pockets and measured peroxidase-like activity toward the oxidation of TMB and ABTS. Moderate enhancement of the specificity and catalytic activity was achieved by imprinting with a neutral monomer. However, when imprinting with a charged monomer, the catalytic activity increased 229 dramatically, 100-fold, compared to that of the bare Fe3O4. Another limitation of MNPs for biosensing is associated with the fact that most of these materials demonstrate their highest peroxidase-like activity at a high temperature (40–45 °C), which limits their application in disease-specific biomolecule detection at room temperature. Therefore, a highly porous

framework of MNPs with high specificity could offer enhanced peroxidase-mimetic activity even at room temperature.

2.1.4. Application of superparamagnetic nanoparticles in biomolecular detection

2.1.4.1. Electrochemical biosensors Electrochemical techniques have aroused great interest in constructing biosensors for the detection of biomolecules, as the electrochemical biosensors offer sensitive, highly selective, portable, easy-to-operate, and stable operation as well as fast detection. In electrochemical sensors, electrode materials play a critical role in obtaining high-performance sensing platforms via various analytical principles, such as voltammetry (CV, DPV, LSV, etc.) and amperometry (i-t). Incorporating superparamagnetic nanomaterials into common electrode materials not only can produce a synergistic effect among biocompatibility, magnetism, catalysis, and conductivity but also can provide accelerated signal transduction and amplified biorecognition events, resulting in an ultrasensitive biosensing platform. In recent decades, significant research has been conducted on the construction of various magnetic materials, such as the construction of functional electrode surfaces as signalling tags or as electrocatalysts, giving rise to advanced electrochemical biosensors. For example, Zhu et al. highlighted recent advances in nanomaterial-based electrochemical sensors and biosensors. Electrochemical, nonelectrochemical, magnetic, and chemical optical sensors and biosensors based on nanocrystalline iron oxides and their composites have also been highlighted by Urvanoba et al.41, 49 In this section, we mainly focus on the advances of nanostructured superparamagnetic materials-based electrochemical nucleic acid assays, immunoassays, and cytosensing platforms.

2.1.4.1.1. Nucleic acid assay The detection of target sequences of nucleic acids (DNA or RNA) has aroused considerable attention due to their wide application in gene therapy, molecular diagnostics, epigenetics, pathogen detection, and early screening of malignant diseases, like cancer. Highly sensitive and specific detection is desirable, as the DNA levels are scant in most of the diagnostic specimens, such as cancer, infectious diseases, or pathogens. In recent decades, several electrochemical DNA/RNA detection platforms have been developed with high selectivity, excellent sensitivity (attomolar detection limit), and on-site measurements.230 Most of the standard DNA biosensors consist of a single-stranded (ss) capture probe immobilized onto the

sensing surface for the hybridization of target sequences with electrochemical tags for signal generation.6 Several amplification strategies have been utilized to achieve ultrasensitive detection. The natural enzyme (e.g., alkaline phosphatase (ALP) and HRP)–assisted amplification (enzymatic catalytic reactions) has been widely used for this purpose.231-232 The integration of magnetic materials with such amplification not only can enhance sensitivity and selectivity but also offers a low-cost miniaturization platform. For example, an HRP-modified

Fe3O4 NP (as a signal amplification source)–based sequence-specific DNA sensor has been proposed by Dong et al.233 In this sensor, HRP was adsorbed onto the NP surface through layer- by-layer (LbL) techniques, followed by the loading of Au-NPs to obtain Au-HRP-Fe3O4 conjugates. A signal probe and diluting probe sequences were then added to form DNA–Au–

HRP–Fe3O4 (DAHF) bioconjugates. As shown in Figure 2.1.13, the capture probe was immobilized on a freshly prepared gold nanofilm electrode surface to hybridize target DNA (two-base mismatched DNA). In the presence of target DNA, the DAHF bioconjugates attached to the GNF surface via the hybridization of target DNA with a diluting probe sequence. An HRP-catalyzed TMB oxidation reaction was then carried out to amplify the chronoamperometric signal. In conjunction with gold-coated ferric-oxide NPs and streptavidin peroxidase–induced signal amplification, Loaiza et al. have developed similar electrochemical DNA sensors for the detection of specific DNA hybridization events.234 High sensitivity in such hybridization-based DNA detection can be accomplished by the utilization of optimum hybridization conditions, the design of the capture and signalling probe, and the elimination of nonspecific bindings in the transducer surface. A peptide nucleic acid (PNA) probe has been known to possess excellent sequence-specific affinity and stability in the recognition of target sequences as they maintain equidistance between nucleobases, rigid amido bonding, flexible amino-ethyl linkers and intramolecular hydrogen bonding, MNPs have also been immobilized with a PNA probe for the construction of DNA sensors based on PNA-DNA hybridization.235

Figure 2.1.13: A magnetic Fe3O4–based sandwich-type detection strategy for two-base mismatched DNA detection. Reproduced with permission.233

miRNA is a small noncoding RNA that post-transcriptionally regulates gene expression by modulating downstream proteins, thus acting as a potential diagnostic and prognostic biomarker of various diseases, including cancer. Several conventional approaches, such as northern blotting, in situ hybridization, microarrays, and quantitative real-time PCR (qRT- PCR), and several biosensors have been applied successfully. Though each of these methods has superior detection sensitivity, the conventional approaches rely on either PCR-based amplification or fluorescent labelling and expensive, complicated procedures.236 In recent decades, several electrochemical and optical biosensors have been developed that may give a superior analytical performance; yet they still rely on complex and tedious amplification processes, expensive biomaterials, and time-consuming and difficult procedures. Recently, nanomaterials have been merged with conventional biosensors to create an easy and inexpensive miRNA detection platform. For example, carbon nanotube (CNT) and single-wall carbon nanotube (SWCNT) nanowires have been widely used in electrode design.237 Recently, we described the synthesis of specially engineered superparamagnetic gold-loaded nanoporous iron oxide nanocubes (Au@NPFe2O3NC) and an amplification-free, nonenzymatic, and sensitive miRNA detection platform45 based on the magnetic, electrocatalytic, and miRNA- adsorbing properties of these nanocubes. The nanocubes showed excellent electrocatalytic activity toward the reduction of redox molecules Ru(NH3)6Cl3. As can be seen from Figure 2.1.14, the nanocubes have been attached to a screen-printed carbon electrode (SPCE) with the help of an external magnet without any complex electrode modification steps. Target miRNAs

were physically adsorbed onto the gold NPs of nanocubes via RNA–gold affinity interactions. The amount of miRNA adsorbed onto the nanocubes was quantified by chronocoulometric 3+ (CC) charge interrogation in the presence of a signalling redox molecule [Ru(NH3)6] . The assay can successfully detect a 1 pM level of miRNA (miR-21) in tissue samples derived from patients with esophageal squamous cell carcinoma (ESCC). To achieve more sensitive detection of miRNA, we recently reported another biosensor design, wherein a redox molecule 3+ 3- [Ru(NH3)6] coupled with another redox molecule [Fe(CN)6] to create an electrocatalytic 238 3+ redox cycle. The CC signals for the electrocatalytic reduction of [Ru(NH3)6] were 3- 3- amplified by further reduction of the solution-phase [Fe(CN)6] . Here, as the [Fe(CN)6] in solution is a stronger oxidant, it initiates the electrocatalytic cycle via the oxidation of reduced 2+ 3+ [Ru(NH3)6] for the regeneration of [Ru(NH3)6] . The redox cycle–induced signal- enhancement steps to enhance the detection sensitivity up to 100 aM for the detection of miRNA (miR-107) from cancer cell lines and a panel of tissue samples derived from patients with ESCC. This design could be integrated into a microfluidic device to achieve a low-cost, simple, ultrasensitive miRNA sensor.

Figure 2.1.14: Superparamagnetic nanoparticle-based miRNA sensor. The target miRNA (miR-21) was initially extracted from target cell lines or tissue samples, followed by magnetic bead-based isolation and purification. The target miR-21 was then directly adsorbed onto the nanocube-attached SPCE. An enhanced electrochemical signal was generated by the CC 3+/2+ interrogation of stoichiometrically bound [Ru(NH3)6] complexes with nanocube surface- bound miRNA. Reproduced with permission.45

2.1.4.1.2. Immuno assay Electrochemical immunosensors are a specific antibody-antigen interaction (noncovalent)– based detection platform in which the immunochemical reaction event is coupled to a signal transducer. Generally, an enzyme-labelled antibody or antigen is immobilized at the electrode surface to perform a sandwich-mode operation. Voltammetric or amperometric detection is used to accomplish the conversion of biorecognition events into an analytically significant electrochemical response.239 In MNP-based electrochemical immunosensors, the recognition element is immobilized with the NPs, which are later used for either electrode modification or as signal-generating probes. MNPs have generally been polymerized to obtain a thin polymer shell so they can be easily functionalized with antibodies, redox molecules, or any linking groups.240 For example, an ultrasensitive sandwich-type electrochemical immunosensors for the detection of a cancer biomarker; prostate-specific antigen (PSA) have been reported based on ferrocene-modified magnetite NPs.241 Ferrocene-functionalized MNPs and graphene sheets were used as the electroactive label and sensing platform, respectively. Graphene sheets were chosen, as they offer an enhanced surface area for capturing a larger amount of primary antibodies (Ab1) and for the sensitive detection of ferrocene. In this assay, dopamine (DA) was first anchored to the magnetite surface, followed by the conjugation of ferrocene monocarboxylic acid (FC) and a secondary antibody (Ab2) onto Fe3O4 through the amino groups of DA (DA-Fe3O4−FC-Ab2). The high number of DA molecules anchored to the Fe3O4 surface increased the immobilization of ferrocene and Ab2 onto the magnetite NPs, which in turn increased the sensitivity of the immunosensor. In another report, functional nanocomposites of Au and Ag core-shell magnetic graphene loaded with a cadmium ion 2+ 242 (Au@Ag/GS-Fe3O4/Cd )–based immunosensor were used for the detection of IgG. The amino-functionalized magnetic graphene nanocomposites (NH2-GS-Fe3O4) were used to bond Au and Ag core-shell NPs (Au@Ag NPs) followed by the adsorption of a cadmium ion (Cd2+). 2+ The Au@Ag/GS-Fe3O4/Cd increases the electrocatalytic activity toward hydrogen peroxide

(H2O2) and improves the effective immobilization of antibodies. The sensor can detect the IgG with a detection limit of 2 fg/mL.

2.1.4.1.3. Cytosensors In electrochemical cytosensors, a biocompatible recognition unit is fabricated onto the electrode surface to specifically recognize the target cells, followed by coupling with a sensitive electrical readout. Varieties of specially engineered nanomaterials have been shown 56

to anchor target-cell recognition units, such as aptamers, antibodies, or receptors, as well as to report enhanced cell-recognition events. For example, Au NP–decorated magnetic Fe3O4 nanoprobes were reported for the construction of an electrochemical cytosensor for the detection of leukemia cells and the quantitative estimation of death-receptor expression on leukemia cell surfaces.243 The nanoprobes were assembled through the co-immobilization of both rhTRAIL (recombinant human TRAIL) and HRP on Au NP–Fe3O4 beads. The rhTRAIL had been used for specific and selective recognition of DR4/DR5 on leukemia cell surfaces, and an amplified electrochemical signal was achieved via HRP catalyzation of the oxidation of thionine by H2O2, resulting in the sensitive detection of leukemia cells with LOD ∼40 cells. In a later year, nonenzymatic nanoelectrocatalysts, Fe3O4@nanocage (Ag-Pd) core-satellite hybrid NPs, were engineered by the same research group as signal-amplifying nanoprobes for the ultrasensitive detection of low abundant circulating tumour cells (CTCs).244 CTCs have been considered a valuable biomarker for early diagnosis of cancer, as they play critical roles in metastasis. As can be seen from Figure 2.1.15(A), the nanoelectrocatalysts were prepared via the electrostatic interaction between positively charged (as-synthesized) Fe3O4/PDDA (di- allyl-dimethyl ammonium chloride) and negatively charged Ag−Pd nanocages. Fe3O4@Ag−Pd hybrid particles were shown to be highly catalytically active toward the electrochemical reduction of a redox dye; thionines and the nanocage were used as signal-amplifying nanoprobes for cytosensor design. The inherent electroactive properties of Fe3O4 and the large surface area, high conductivity, and distinctive porous hollow structure of the Ag−Pd nanocages contribute synergistically toward amplification of the catalytic signals. Figure 2.1.15(B) illustrates the steps involved in the sensor design. A GCE surface was modified with the Au NPs, followed by the conjugation of SYL3C−SH (thiolated cell–targeting aptamers) via

Au−thiol interactions. An SYL3C-functionalized hybrid nanoprobe, SYL3C−Fe3O4@Ag−Pd, was formed by the conjugation of thiolated SYL3C with the Ag−Pd nanocages through metal−thiol interactions. After capturing the target cells (CTCs) onto the SYL3C-Au modified

GCE, the SYL3C−Fe3O4@Ag−Pd nanoprobes were attached to form a sandwich-resembling superstructure. The DPV responses were then measured in the presence of thionines to quantify the two model cells (MCF-7 and T47D) and the sensors able to detect ∼4 MCF-7 and ∼5 T47D cells. Thiolated sgc8c aptamer–immobilized Au NP–coated magnetic Fe3O4 NPs were also reported for the detection of leukemia cancer cells.245

Figure 2.1.15: Schematic illustration of (A) the fabrication of Fe3O4@Ag-Pd hybrid NPs and (B) the steps involved in sensor design for the detection of CTCs. Reproduced with permission.244 Sun and co-workers have engineered a hybrid nanoelectrocatalyst

(Fe3O4/MnO2/Au@Pd) as both an electrochemical signal amplifier and a carrier of nanoprobes for the sensitive detection of human hepatocellular cancer cells (HepG2).246 The nanocatalyst was designed by the conjugation of an Au@Pd core-shell nanosphere and Fe3O4/MnO2 nanocomposite and modified with a thiolated capture probe, HRP, and hemin to form a

Fe3O4/MnO2/Au@Pd–HRP–aptamer/hemin/G-quadruplex nanoprobe. After the capturing target cells on a thiolated aptamer–attached AuNP/GCE surface, a sandwich-like structure was formed with the as-prepared nanoprobes. After that, an amplified DPV signal was obtained from benzoquinone (BQ) generated from Fe3O4/MnO2/Au@Pd–HRP–aptamer/hemin/G- quadruplex nanoprobes that catalyzed the oxidation of hydroquinone (HQ) with H2O2. The generated DPV response was directly related to the number of nanoprobes present on the AuNP/GCE surface, thus reflecting the number of cells present in the system. Very recently,

Fe3O4NP nanozymes with reduced graphene oxide/molybdenum disulphide (rGO/MoS2) composites have also been reported for the detection of CTCs.247

2.1.4.2. Optical biosensors Novel metal nanostructures with a dimension below 100 nm possess numerous unique optical properties, such as absorption, emission, surface plasmon polariton propagation, and localized and surface plasmon resonance, in the extended visible and near-infrared (IR) regions.34, 248 Their distinct properties, including magnetism, engineered dimension, opto-electronics, and surface functionalization with antibodies, ligands, and recognition elements capacity, make superparamagnetic NPs superior candidates for the rational design of a sensing platform for biomolecules with low concentration. Based on such distinctive properties and plasmonic behaviours, numerous MNPs-based optical detection has been reported.

2.1.4.2.1. Colourimetry and fluorescence detection Colourimetric sensors have been considered for emerging spot-chemical detection techniques, as they offer detection with minimal facilities due to the replacement of complicated steps and expensive instrumentation. They can be easily integrated into the solid strips, papers, smartphone, thus allowing portability and affordability of the detection of target samples. Integration of MNPs with colourimetric sensors offers more sensitive, accurate, miniaturized, cost-effective, instantaneous in situ detection of target analytes. The detection mechanism generally relies on the functionalization of MNPs with the recognition elements (e.g., antibodies or aptamers), followed by the capture and magnetic isolation–purification of targets.249,250 The purified MNP-bound target is then used to interact with a chromogenic substance or to undergo NP enrichment to generate analyte-defined colours. For instance, anti- Staphylococcus aureus (S. aureus) antibody–functionalized AuNP/MNPs have been used to detect a food pathogen, S. aureus, in milk.251 The immunomagnetic composite was inoculated into the milk sample to capture the target, followed by magnetic isolation and purification. The resuspension of captured bacteria-antibody/AuNP/MNP complexes was passed through a selective filtration system, while the target unbounded NP composite filtered through the membrane and the target bound NP composite did not penetrate, resulting in corresponding red spot through a gold enhancement solution. The assay can detect 1.5 × 105 CFU of S. aureus in milk, but it required a predesigned filtration system, signal (colour intensity) enhancement, and solution treatment. Suaifan et al. reported a lab-on-a-chip (LOC) strip-based biosensor for detecting cancer biomarker PSA protease without using any chromophore labelling or colour- enhancement solution.252 In this sensor design, a PSA substrate peptide was covalently bound to a magnetic bead and linked to the gold surface of the paper strip. The PSA protease was then applied to cleave the strip-bound PSA substrates, resulting in the release of a black magnetic 59

carrier complex and thereby exposing the target corresponding gold-coloured sensor surface to naked-eye detection. Recently, several other paper colourimetric biosensors based on the proteolytic activity of the target have also been reported to detect food contaminants in complex foods and milk.253-254 However, in recent decades, many MNP-colorimetric sensors have been reported based on the intrinsic peroxidase mimetic activity of MNPs, which is described separately in section 2.1.5.3.

2.1.4.2.2. SPR biosensors Surface plasmon resonance (SPR) sensors possess a unique capacity for real-time monitoring of the interaction of biological analytes by evaluating the refractive index changes during complex (usually sandwich-type) formation or dissociation at the SPR sensing surfaces.255 SPR sensors have generally suffered from the low signal intensity and nonspecific bindings for the detection of trace amounts of proteins, nucleic acids, and other small biomolecules.256 To obtain 257 258 259 260 signal enhancement, several metal nanoparticles, including Au, Pt, Pd, and SiO2, have been investigated; however, these NPs (due to their low molecular weight and slow diffusion to the sensor surface) are unable to realize the promise of achieving highly selective and sensitive detection. Target-specific enrichment, isolation of the target from complex biometrics, and simultaneous signal enhancement could be a way to perform targeted and sensitive SPR-based biomolecule or biomarker detection. Magnetic nanostructures hold promise for providing a high surface-to-volume ratio, minimum disturbance to surface functionalization, easy and faster binding, better miscibility, magnet-based isolation and purifications, high molecular weight, and necessarily quick delivery of target analytes to the sensor surface through a magnetic field gradient to choke the 261 slow, diffusion-driven mass transfer. By utilising Fe3O4 MNPs as amplification reagent, a sandwich SPR sensor has been reported for detecting thrombin by utilizing Fe3O4 MNPs as amplification reagents; the MNP–aptamer conjugates were used to accomplish the antithrombin aptamer–thrombin–MNP sandwich on an SPR sensor, and a LOD of 0.017 nM was achieved.262 However, though the greater mass and superior refractive index of the MNPs located in the evanescent field give rise to signal enhancement, the MNPs tend to aggregate and possess bad biocompatibility if it is not well-functionalized with biomolecules or antibody or any metallic coating through cumbersome chemistries.96, 263 One of the prominent approaches to overcome this is to employ the well-known surface chemistry of Au. The loading (or coating) of gold onto MNPs stabilizes the particles in solution, enabling the straightforward binding of capture probe molecules or target biomolecules and improving the signal enhancement by combining the refractive indexes from both Au and MNPs (GMNPs - gold MNPs).264 60

Besides Au, graphene oxide (GO)–modified MNPs are also promising in SPR-based biosensing. GO provides multiple sites for the deposition of other metal oxides, whereas the reduced GO (rGO) provides an sp2-hybridized lamellar scaffold for carrying different proteins and probe 265 molecules. By combining both the Au and GO with Fe3O4 MNPs, an SPR sensor was developed for detecting the human IgG protein.266 As shown in Figure 2.116, the carboxyl group– functionalized GO was employed as a sensing surface for immobilizing the capture antibody (Ab1). A magnetic nanohybrid was formed by combining the MNPs with hollow Au NPs (HGNPs), followed by modification with a detection antibody (Ab2). An Ab2 nanohybrid was then utilized to magnetically collect target IgG from the sample and to rapidly deliver the target sample to the sensor surface using an external magnet. Taking the advantages of magnetic field–driven mass-transfer, inner and outer surfaces derived notable plasmonic fields, and the significant signal amplification of magnetic nanohybrid the developed SPR sensor achieved a LOD of 1.88 ngmL-1 toward the detection of IgG. Notable signal enhancements are observed when the plasmonic effects spreading over the inner and outer surface of HGNPS are combined with the magnetic field-driven mass transfer effects of Fe3O4 NPs. Based on this amplification strategy, improved SPR was reported, in which a detection antibody (Ab2)–modified polydopamine-

Ag@Fe3O4/reduced graphene oxide (PDA-Ag@Fe3O4/rGO) was utilized to attach the target and form a sandwich structure with the target IgG and Ab1 immobilized onto an Au surface.267

Fe3O4 facilitates the magnet-guided sample and Ab2 collection; the PDA permitted efficient immobilization of the antibody and prevented agglomeration. Ag NPs were excited to generate SPR, and their hot electrons were doped on thin graphene films, which improved the response of the target IgG.

Figure 2.1.16: Schematic representation of a GO-sensing film and Fe3O4-HGNPS- hybrid probe- based magnetic field-assisted SPR biosensor for IgG detection Reproduced with permission.266

2.1.4.2.3. LSPR biosensor Unlike SPR, a localized surface plasmon resonance (LSPR) biosensor depends on the light- matter interactions of noble metal NPs, where the sensing is realized by monitoring the changes in LSPR peak position arising from the refractive index alteration of the environment or changes in nanoparticle coupling strength.255, 268 LSPR can be designed either in solution-phase or surface-bound mode. Though solution-phase LSPR follows a simpler operation, the colloidal stability and dilution of NPs limit their in-field application. Surface-bound LSPR (in which NPs are immobilized on a solid surface), in contrast, alleviates the problem of NP colloidal stability in solution and is easy to integrate into the multiplex and array-based assays. Oligo-functionalized Au NPs are primarily used for aggregation-based DNA or RNA sensors, where the hybridization of the target or complementary sequences cause aggregation, resulting in the redshift of the LSPR peak.268-269 The integration of MNPs with plasmonic NPs such as Au, Ag, or Pt facilitates the dual benefit of MNP-based easy and various modes of conjugation, dispersibility, and magnetic separation of preferred biomolecules, as well as plasmonic NP- 270-272 based SPR signal amplification. For instance, citrate-stabilized Fe3O4@Au core-shell NPs were employed for the multiplex detection of serum proteins and miRNAs as the plasmon signal amplification label. Both the number of analytes bound onto the sensor (i.e., antigen for an antibody or miRNA for a capture nucleotide) and the level of interaction of the detection

probe–loaded core/shell NP labels stimulates the SPR signal change.270 Interestingly, the changes are larger for small oligonucleotide hybridization than for the large sandwich protein immuno-mode. A more recent example of a magnet-assisted LSPR immunoassay in which

Fe3O4/Au core-shell NPs (FACSNPs) were utilized to fabricate sensing spots of a microarray system.273 By coupling the unique superparamagnetic property of the iron oxide nanocore with the strong plasmonic strength of AuNPs on the FACSNPs demonstrated superior sensitivity to the local refractive index change upon cytokine binding. In this microarray system, magnet- assisted self-assembly was performed to obtain pattern uniform antibody functioned microarray on a glass substrate. The FACSNP microarray can conduct 384 tests on four different cytokines for each sample, with 16 replicates per cytokine test. The integration of FACSNP microarray sensors into a simple opto-fluidic device enables real-time, parallel detection of multiple cytokines with a LOD of ≈20 pg mL-1 using 1 µL of clinical samples.

2.1.4.2.4. SERS biosensors The surface-enhanced Raman spectroscopy (SERS) technique shows vast superiority for use in biological systems aiding from the technology itself using Raman scattering enhancement since it can provide orders of magnitude of signal intensity upon employment of proper substrates and enhancers. Recently, SERS has been integrated with biosensing for the detection of sensitive biomolecules such as proteins,274 DNA,275 RNA,276 and cellular components277 due to their unique Raman responses. Generally, Au and Ag NPs are the predominantly utilized ones, as their surface plasmons lie in the visible region of the electromagnetic spectrum that overlaps the laser excitation wavelengths frequently used for Raman.278 The integration of MNPs has accelerated the detection by incorporating magnetic capture of the target from clinical samples as well as carrying reporter molecules (Au, Ag) or dyes. For instance, Au@MNPs, in which both plasmonic properties from Au and magnetic properties from MNPs are integrated, thereby enhancing the SERS detection sensitivity by increasing the active concentration of both the tag and reporter within the stationary focus beam of a laser following magnet-based pull-down.96, 279-280 Yang et al. explored a SERS- based immunoassay to detect an influenza virus, H3N2, through the formation of a sandwich structure consisting of 4-mercaptobenzoic acid (4-MBA)–labelled Au NPs as SERS tags, Fe3O4/Au NPs magnetic supporting substrates, and target influenza viruses (Figure 2.1.17).281 In this immunosensor, Fe3O4/Au enriches and separates the viruses from the biological matrix, thereby simplifying the sample pretreatment and the coupling agent 4-MBA loaded on Au NPs bind Influenza A IgG. The 4-MBA itself acts as a Raman reporter due to its intrinsically strong Raman scattering. The advantages of utilizing 4-MBA rely on the fact that, unlike thioglycolic 63

acid or α-lipoic acid, it does not require the addition of further Raman reporter molecules.282 2 The developed immunoassay could detect down to H3N2 10 TCID50/mL. Like Au, Fe3O4@Ag MNPs was utilized as both a SERS tag and a target capturing agent to design a sensitive miRNA 279 detection platform. In this sensor design, the DNA probe–modified Fe3O4@Ag MNPs were first allowed to capture target miRNA from cancer cells followed by DSN-based signal amplification for SERS detection. Upon the hydrolysis of the DNA probes of the DNA/RNA duplex, the Raman tags could diffuse away from the Ag surface and induce a Raman intensity attenuation. Though SERS tags are highly versatile and tremendously useful, the precise fabrication of identical nanogaps distribution using the metal (single) is highly challenging, which consequently limits the sensor platform from high sensitivity and reproducibility.

Figure 2.1.17: Schematic illustration of a magnetic SERS immunosensor for sensitive detection of avian influenza virus. Reproduced with permission.281

The SERS platform should have both the chemical enhancement abilities and strong electromagnetic effect to provide drastically higher sensitivity than Raman spectroscopy for detecting trace amounts of disease-specific biomarkers (e.g., neurodegenerative disease. This can be achieved by incorporating GO into the SERS platform, due to its ability to improve the Raman signal through chemical enhancement.283 The unique structure of sp2-carbon nanosheets is promising for π−π stacking, and the highly electronegative oxygen species present on the GO surface can enhance the local electric field on the adsorbed molecules. Additionally, GO has a strong ability to quench the molecule’s fluorescence, which adds further advantages as a SERS substrate via immense decreasing of the background signal. Moreover, the platform with multiple hotspots to enrich the target analytes facilitates amplified SERS

signals. The numerous hotspots can be attained by creating subtle nanogaps using a core-shell– satellite structure. Several researchers have reported taking advantage of both the core-shell arrangement and the GO-induced enhancement and magnetic effect of MNPs to produce several SERS platforms.284-285 For instance, magnetic core–plasmonic shell NP-attached hybrid GO-based multifunctional nanoplatforms have been reported for detecting Alzheimer’s disease (AD) biomarkers tau-protein and β-amyloid.285 In this platform, magnetic core-shell NPs were assembled on 2D GO surface and utilized to isolate the biomarkers from complex patient samples. However, though MNPS are widely used in SERS, still they need to integrate with different functional particles and required cumbersome conjugation chemistries. A single nanostructure with magnetic properties, high plasmonic and electronic properties, and high analyte adsorption capabilities with an intense layer could provide a single-particle SERS platform and overcome the existing limitation of using multiple materials and complex chemistries.

2.1.4.3. Biosensors based on nanozymes Peroxidases (mostly HRP) play significant roles in bioanalytical chemistry, as they are widely used (conjugated with antibodies or signalling molecules) as signalling agents to catalyze the oxidation of various chromogenic substances in sensor design. After Yan reported the peroxidase mimetics of iron oxide NPs and their application in the development of an immunoassay, a vast number of studies have been conducted on the design of biosensors based on nanozymes.208, 214, 286 Yan and co-author designed two immunoassays for detecting the hepatitis B virus surface antigen (preS1) and cardiac troponin 1 (Tn1) using ferric oxide– immobilized protein A and an antibody, respectively. In the first immunoassay, an ELISA plate was coated with Pres1, followed by the addition of an anti-HBV Pres1 antibody. Ferric oxide– immobilized protein A was then added to complete the immunorecognition events and catalyze the TMB substrate solution to generate antigen-corresponding colour signals. In the second assay, Tn1–immobilized ferric oxide was mixed with a serum to bind the target Tn1 and magnetically separate it from the complex serum matrix, followed by assay design onto an ELISA plate. Based on such peroxidase mimetics, polyacrylic acid-coated iron oxide MNPs were designed and used to detect carcinoma cells (A549 cells). The assay was able to differentiate A549 cells from non-carcinoma cells (H9c2).287 Since then, several immunoassays have been reported using magnetic nanoparticles. However, the nanozyme activity usually decreases when they are modified with proteins, antibodies, or nucleotides. Recently, Yang et al. reported that phosphate-containing adenosine analogues such as adenosine 5′-

monophosphate (AMP) enhance the oxidation reaction of hydrogen peroxide (H2O2) as well as 288 TMB oxidation. They have found that the nanozyme activity in Fe3O4 NPs gradually increases with increasing AMP concentration and polyadenosine length, and it follows a trend: AMP > adenosine 5′-diphosphate > adenosine 5′-triphosphate. Based on these AMP-enhanced peroxidase mimetics, they have designed a selective fluorescent turnoff system for the quantification of human serum albumin (HSA) in the urine. Very recently, we reported on electrocatalytic and colourimetric (naked eye) detection of p53 autoantibodies using peroxidase mimetics of gold-loaded nanoporous ferric oxide nanocubes.289 In this sensor design, the biotinylated p53 antigen was immobilized on a neutravidin-modified screen-printed carbon electrode (SPCE), followed by the addition of serum or plasma samples containing the target p53 autoantibody (Figure 2.1.18). α-Human IgG–functionalized Au−NPFe2O3NCs were then added to the electrode surface to form an immunocomplex with the target p53 autoantibody.

The electrode surface was then covered and incubated with the freshly prepared TMB/H2O2 solution to facilitate the nanocube-catalyzed oxidation of TMB. The colour change was observed and quantified by using colourimetry and chronoamperometry.

Figure 2.1.18 Schematic representation of the assay for p53 autoantibody detection using the peroxidase mimetics of gold-loaded nanoporous ferric oxide nanocubes. Reproduced with permission.289

The peroxidase-mimicking activity of nanozymes has also been used to detect nucleotides. For instance, a colourimetric DNA-sensing platform has been demonstrated using

dsDNA shielding activity against the peroxidase mimetics of NPs.290 The target was subjected to PCR amplification, followed by mixing with unmodified MNPs. The PCR-amplified target DNA present in the solution results in a decrease in access of the positively charged substrate o-phenylenediamine (OPD) to the MNPs through its electrostatic interactions with the negatively charged phosphate backbone of DNA and direct adsorption of DNA molecules on the surface of MNPs. This leads to significant inhibition of substrate (OPD and H2O2) binding to MNPs. Table 2.12 summarizes the recently employed (in the past three years) magnetic nanostructures and their role in the detection of disease-specific biomarkers. In addition to the above-discussed strategies, other techniques, such as giant magnetoresistive (GMR),291-293 quartz crystal microbalance (QCM),294-295 microcantilevers,296-297 and resonance Rayleigh scattering (RRS),298 play critical roles in the development of a sandwich assay for protein- based biomarker detection.

Table 2.2: Engineered MNPs for disease-specific biomolecular (electrochemical and optical) sensing Magnetic nanostructures (MNSs) Function of MNSs Target biomolecule LOD Ref. Electrochemical detection

45 Au@NPFe2O3NC Immobilization of miRNA miRNA (miR-21) 100 fM Carrying a large number of redox signalling molecules, 299 Fe3O4 NPs miRNA (miR-141) 0.28 fM thionine and Fc, for target-specific DPV readout

300 CoFe2O4 MNPs Nanoelectrocatalyst for toluidine blue catalysis miRNA (miR-21) 0.3 fM GO-loaded–iron oxide hybrid Electrocatalytic signal enhancement miRNA (miR-21) 100 aM 301 materials Immobilization of miRNA DNA–Au@MNPs as dispersible DNA–Au@MNPs as dispersible electrodes miRNA (miR-21) 10 aM 302 electrodes Nanocarriers for signal probe PdNPs@Fe-MOFs Immobilization of redox probes and electrocatalysts for miRNA (miR-122) 0.003 fM 303 signal enhancement Immobilization of ssDNA ctDNA (FGFR2 : FAM76A GO-loaded MNPs 1 fM 304 Electrocatalytic signal amplification fusion gene) Immobilization of probe DNA and redox probe of 305 Porous Fe3O4 miRNA (miR-141) 1.4 aM 3– [Fe(CN)6] Immobilization of target biomarker and magnetic field– 306 Fe3O4@GO@MIP Interleukin-8 (IL-8) 0.04 pM based assembly Magnetic actuator Au-coated iron oxide NPs PSA 0.085 ng/mL 307 Immunoprobe 4.5 pgmL−1 Aptamer immobilization Thrombin, PDGF-BB −1 308 Fe3O4@3D–rGO@PP4VP 29.4 pgmL Electrode fabrication LYS 14 pgmL−1

309 Au@NPFe2O3NC Nanocarriers for target p53 from serum p53 autoantibody 0.02 U mL−1 Direct isolation of target protein from serum; acts as a p53 autoantibody 0.08 U/mL 40 Au−NPFe2O3NC nanozyme Exosome 103 exosomes/mL 310 DNA probe labelled with NPS to offer an electrochemical −15 311 Fe3O4/TMC/Au RASSF1A methylation 2×10 M readout

312 Fe3O4/TMC/Au Nanocomposite utilized as a tracing tag to label antibodies EGFR fg/mL Influenza virus DNA 8.4 pM Au/MNP-CNT Magnetic alignment of sensing platform on a Pt electrode 313 Norovirus DNA 8.8 pM Colourimetric and fluorescence detection

314 ZnFe2O4/rGO Aptamer conjugation to act as a signal probe S. typhimurium 11 CFU/mL MNPs Immobilization of target-specific DNA probe 2 ETA gene 1.2 ng/mL 315 Immobilization of detection antibody and HRP; DL–MBs EV71 0.1 ng/mL 316 colorimetric readout AuMNPs Immobilization with complementary DNA probe HPV-16 E6 100 pM 317 Primary antibody conjugation and magnetic separation 318 SiO2-coated iron oxide NPs CEA 3.7 pg/mL and purification E. coli pathogens Immunomagnetic beads Carrier for purification/separation and magnetic focus AFP 104 CFU mL−1 319 CEA Functionalizing with detection antibody and iron oxide- Iron oxide NPs to-Prussian blue (PB) NP transformation for readout PSA 1.0 ng mL−1 320 signals Recognition and concentration elements and magnetic 321 Fe3O4 Aβ oligomer 36 pM separation

Functionalizing with target-specific DNA probe 1 Vibrio cholera O1 OmpW gene 2.34 ng/mL 322 MNPs

−13 323 Fe3O4@SiO2–Au nanocomposites Fluorescence quenching and magnetic separation PSA 3.0×10 g/mL SPR and LSPR

262 Fe3O4 Signal enhancement thrombin 0.017 nM Loading of capture antibody -1 264 Fe3O4@Au human-interleukin-17A 0.05 ngmL Target delivery to the sensor surface Binding of selective antibody Iron oxide MNPs Signal enhancement due to the large refractive index and MCF-7 cancer cell 500 cells/mL 324 larger mass Capturing target IgG Magnet-derived faster sample delivery -1 266 HGNP–Fe3O4 IgG 1.88 ngmL Signal enhancement by the combined effect of HGNPs

and Fe3O4 MNPs Capturing target CTnI and LSPR signal enhancement Cardiac troponin I (cTnI) 30 pM 325 Magnet pattern sensing array and LSPR signal -1 Fe3O4/Au Core-shell Cytokines ≈20 pgmL 273 enhancement IL-6, 28 pM; IL-8, 18 Interleukin (IL-6 and IL-8) & Fe3O4@Au core/shell Signal amplification label pM; miR-21, 502 fM; 270 miRNA (miR-21 and miR-155) and miR-155, 483 fM Water-dispersed MNPs Target capture and immuno-recognition cTnI 15 ngmL-1 326 3-Mode signal enhancement by the combined effect of −1 267 Ag@Fe3O4/rGO IgG 0.019–40.00 μgmL HGNPs, Ag, and Fe3O4 Sc –4 327 Aptamer–Fe3O4 NPs Signal amplification PrP 1×10 ng/mL SERS Au-spiked silica-coated iron Separation of target DNA DNA oligonucleotides 328 oxide spheres Reduced signal based on the hairpin structure

329 Fe3O4@Ag Magnetic capture, isolation, and purification of miRNA miRNA (let-7b) 0.3 fM Conjugation of S. aureus antibody to form sandwich- Staphylococcus aureus (S. 330 Au-coated MnFe2O4 10 cells/mL structured immunoassay aureus) Supporting substrates; enrichment and separation of 2 281 Fe3O4/Au NPs AIV 10 TCID50/mL viruses from a complex matrix Iron magnetic core−gold Target-specific antibody conjugation, magnetic β Amyloid 0.312 ng/mL 285 plasmonic-shell NPs separation, and electromagnetic effect on SERS hotspot Tau protein 0.15 ng/mL Abbreviation: TMC: N-trimethyl chitosan; EGFR: epidermal growth factor receptor; PSA: prostate-specific antigens; PrPSc: prion disease-associated isoform; AFP: alpha fetal protein; CEA: carcinoembryonic antigen; ETA: Exotoxin A; PP4VP: plasma-polymerized 4-vinyl pyridine; EV 71: human enterovirus 71; HPV: human papillomavirus;

IgG: Immunoglobulin G; TCID50: tissue culture infection dose at 50% endpoint; AIV: avian influenza viruses.

2.1.5. Point-of-care testing: The impact of superparamagnetic particles With the advancement of nanotechnology, miniaturization, and microprocessors, it is now possible to analyze samples (urine, blood, saliva, etc.) in real-time while discussing with patients. The combination of the microfluidics, paper fluidics, engineered NPs, and magnetic impact represents the automated, integrated POC system that is relatively simple to use and offers rapid and accurate results with moderately complex equipment. In recent decades, there have been numerous reports on various approaches to integrating MNPs into optical, electrochemical, and piezoelectric biosensing platforms as labels or integrating them into the transducer.331 For instance, the use of micro-urine nanoparticles (μUNPDs) for the detection of trace amounts of molecular markers KIM-1 and Cystatin C in urine has been reported.332 This technique utilizes an automated on-chip assay followed by screening with a handheld device for the readout. Currently, a lateral-flow immunoassay (LFIA) is considered to be a potent, low-cost detection technique for biomolecules. It is highly advantageous over conventional ELISA, as it offers one-step rapid detection by eliminating multiple additions and washing steps.333 The commonly used detecting agents in LFIAs are gold NPs, carbon NPs, and monodisperse latex. Though they give good detection, they lack sensitivity, as the signal generated beneath the surface is missed by conventional optical detectors. MNPs, due to their magnetism and brown colour, can give sensitive detection that can easily be read with an optical sensor and quantified by magnetic flow reader.334 Recently, Pt-decorated magnetic core-shell nanoparticles (MPt/CS NPs) were demonstrated to integrate into lateral-flow POC immunoassay strips for human chorionic gonadotropin (hCG) using the intrinsic magnetic and enzyme-like properties of MPt/CS NPs.335 In this POC device design, MPt/CS NPs conjugated with antibodies (Ab-MPt/CS NPs), followed by the magnetic enrichment of target analytes using Ab-MPt/CS NPs. Ab-MPt/CS NPs capturing target analytes were then applied to the sample pad of lateral-flow immunoassay (LFIA) strips. Ab-MPt/CS NPs containing sufficient amounts of target analytes were captured on the T line, whereas the remaining Ab-MPt/CS NPs were captured on the C line. Finally, a substrate solution was applied to the test pad to obtain amplified signals in T and C lines by peroxidase-like reactions mediated by MPt/CS NPs, as shown in Figure 2.1.19. A similar LFIA system has been reported for the detection of EVs isolated from human plasma.336 In this report, different materials (colloidal gold, carbon black, and MNPs) as LFIA labels were employed and compared. In all such MNP LFIAs, an external magnetic field is required to magnetize the MNPs to detect them in a corresponding applied field. Lago-Cachón et al. reported a superparamagnetic NP–based radio-frequency lateral-flow assay (RF-LFA) method over the external magnetic field–based MLFA, where the continuous 72

flow of MNPs is measured.337 The main benefit of this RF-LFA system over the MNP-based LFA is that it does not require an external magnet-based signal readout. Although the MNP- based LFA offers rapid analysis and a simplified detection system, it is not usually well suited to the multiplex analysis of several biomarkers from a particular disease or to discriminate among multiple diseases presenting with similar symptoms.

Figure 2.1.19: Schematic demonstration for the detection of human hCG using magnetic nanozyme–based LFIA strips. Reproduced with permission.335

An easy-to-use, rapid POC system that could detect multiple biomarkers simultaneously and enable the differentiation of disease states would be highly beneficial for in-field diagnostics of outbreaks of infectious diseases. Recently, an immunoassay technology was reported in which the benefit of MNPs is integrated with SERS readout system to enable the detection of three disease-specific antigens from Ebola, Lassa, and malaria from a single blood sample.338 In this immunoassay system, disease-specific antibodies were conjugated with both the SERS nanotags and a magnetic microparticle and stored in a tube (either liquid or dried stage) (Figure 2.1.20). Upon the addition of a blood sample, a sandwich was formed containing a magnetic particle–target antigen–SERS tag followed by a magnetic pull to separate the sandwich and bring them to the side of the reaction tube. A laser was exposed outside the tube to illuminate the sidewall corresponding to the SERS tag in the sample. As different Raman reporters were integrated, different SERS spectra (Figure 2.120(B)), each with a unique optical “fingerprint,” represent each nanotag (proportional to the level of an antigen in the blood sample). This POC diagnostic is highly advantageous because it does not require

any washing steps, the antibody-modified magnetic particles and SERS tags can be prepackaged, and it is highly suitable for in-field testing. Though several POC systems integrating MNPs (either carrier, magnetic separation, isolation, magnetic focus, or signalling label), LFAs, or an immunoassay system and/or PCR amplification with different readouts (SERS, NMR, or colourimetry) have been reported (Table 2.3), most of them require antibody functionalization, three-step sandwich formation (using capture and detection antibodies), and a complicated readout system.339-346 Table 2.3 summarizes the recently reported (since 2016) magnetic nanostructure–based POC systems for clinics. Therefore, magnetic nanostructures that would not need any further antibody functionalization or a complex readout strategy yet to be performed and highly desirable for future diagnostics in clinics.

Figure 2.1.20: Schematic representation of the detection of Ebola, Lassa, and malaria using magnetic particles and SERS nanotags. (A) Depiction of SERS nanotag technology. (B) Characteristic spectra from three SERS nanotags specific to the Ebola, Lassa, and malaria histidine-rich protein 2 (HRP2) antigen. (C) Representation of the homogeneous no-wash (HNW) sandwich immunoassay using magnetic particles and SERS nanotags. Reproduced with permission.338

Table 2.3: Magnetic nanostructure-based POC diagnostic system for rapid analysis of disease biomarkers Magnetic Analyte Assay Functions Target biomolecule Detection limit Ref. nanostructure volume duration

Fe3O4–QD magnetic Magnetic separations Antigens from S. typhi 1 mL 35 min 3.75×103 CFU/mL 347 nanosphere Magnetic signal (readout) label Signalling probe carrier kim-1 0.1 ng/mL MNPs - - 332 NMR readout Cystain C 20 ng/mL

Sandwich structure formation Antigens from EBOV, LASV, 105 to MNPs 45 µL 30 min 338 Magnetic support for SERS tag and malaria 106 PFU/ml

Magnetic Magnet-drive fluid mixing Hendra virus - 15 min 0.48 ng/mL 339 beads Support of hybridization between capture Magnetic beads probe and DNA Foodborne pathogens - - 10 CFU/mL 340 Magnetic capture and purification

Fe3O4 core–Au shell Magnetic capture of analyte Valosin-containing proteins 100 µL 1 hr 40 min 25 fg/mL 341 nanoprobe Magnetic focus zone (VCPs)

Sandwich structure formation Acute myocardial infraction MNPs 80 µL 20 min 0.014 miu/mL 342 (AMI)

Support for PCR amplification 343 Fe3O4/Au/Fe3O4 Genomic DNA 5 min 5 ng

Magnetic field-based protein preconcentration Magnetic beads Troponin 100 µL 15 min 1 ng/mL 344 The magnetic focus of the target

Magnetic separation of target 345 Fe3O4/Sio2/QDs Clenbuterol 0.3 mL - 0.16 ng/mL Fluorescent label

PAA–Fe3O4 Signalling label NT–proBNP protein 70 µL 40 min 100 pg/mL 346 Nanocrystal clusters

348 SMNPs Signalling label for magnetic signals Unconjugated estriol (µE3) 100 µL 15 min 0.86 nmol/L ~23 CFU Modification of pathogen-specific antibodies Fe3O4/Au core-shell E. coli O157:H7 per ml to control the movement of the captured 100 μl 30 min 349 nanoprobes S. Typhimurium ~17 CFU bacteria at the detection zone per ml 3 µL Antibody functionalization and magnetic Single nucleotide 0.04 pg/μL with Au MNPs (GMNPs) genomic 5 min 350 readout polymorphisms (SNPs) plasmid DNA

2.1.6. Future perspectives and conclusion The requirements of ultrasensitive detection of biomolecules with low concentration and miniaturized biosensing have made the engineering of MNPs in the synthesis, biofunctionalization, and direct application onto a sensor platform the hottest direction in current research. As discussed in this review, many nanotechnological biosensing applications have been reported in both academic research and commercial products over the last two decades. Magnetism properties have been used to capture and isolate the target analytes from a complex biomatrix, therefore reducing the presence of nonspecific biomolecules in the sensors. The application of MNPs in biosensing has broadened when it has been capable of functionalizing with different biomolecules through covalent and noncovalent interactions. The biofunctional MNPs can be used as capturing agents, tracers, carriers, signal generators, signal enhancers, catalysts, and optical emitters. The intrinsic peroxidase-mimicking properties of MNPs have also been widely used in sensor design, and this has opened a broader window of catalysis and replace natural enzymes. These excellent and unique physicochemical properties make MNPs a superior choice for electronic, optical, and optoelectronic biosensor design for the detection of cells, nucleic acid, proteins, extracellular vesicles, and pathogens. Additionally, the nano-dimension and favourable size of biomolecules may facilitate progress toward advanced uses in point-of-care diagnostics and clinics. In this review, we have summarized recently reported (2015 to 2019) remarkable achievements in the application of magnetic nanostructures for the development of electrochemical and optical biosensors and POC diagnostics. Moreover, with a brief focus on MNPs and their synthesis, biofunctionalization, and intrinsic properties required for advanced biosensing, this review has described the challenges involved in current MNP-based detection approaches and has offered an outlook on where imminent developments may be focused. The advances in (superpara-) magnetic, nanostructure-based, disease-specific biomarker quantification reported in recent scientific literature are a sign of flourishing research; nevertheless, a lot of issues need to be considered and addressed to make MNPs for a point-of-care platform; (i) The stability of SMNPs in the aqueous system (e.g., different buffer) still needs to be considered, as many biosensors are based on an aggregation of MNPs; (ii) biomolecules are radially highly sensitive to environmental stress, and they require certain physiological conditions; thus a higher degree of MNP biocompatibility is required, (iii) the biofunctionalization of MNP using a variety of ligands, polymers, enzymes, and inorganic materials enable them to be used for biosensing, however, the half-life of biomolecule–MNP complexes is still low; (iv) the functional molecules cover the core magnetic molecules of 77

biomolecule–MNP complexes, thereby interfering with the magnetic susceptibility of MNPs; and (v) the combination of two or more nanostructures are still considered to achieve functional magnetic nanostructures with large surface areas, high loading, magnetic, and signal transduction and enhancement capacity as well as disease-specific biomolecule binding sites. Single-step magnetic nanostructure synthesis with in situ probe functionalization and easy signal transduction and readout (electrochemical or optical) could also be considered for future magnetic nanostructure-based diagnostics for clinics.

2.1.7. References 1. Heyn, H.; Esteller, M., DNA methylation profiling in the clinic: applications and challenges. Nature Reviews Genetics 2012, 13 (10), 679. 2. Sayed, D.; Abdellatif, M., MicroRNAs in development and disease. Physiological Reviews 2011, 91 (3), 827-887. 3. Jones, P. A.; Baylin, S. B., The fundamental role of epigenetic events in cancer. Nature Reviews Genetics 2002, 3 (6), 415. 4. Mitchell, P. S.; Parkin, R. K.; Kroh, E. M.; Fritz, B. R.; Wyman, S. K.; Pogosova- Agadjanyan, E. L.; Peterson, A.; Noteboom, J.; O'Briant, K. C.; Allen, A., Circulating microRNAs as stable blood-based markers for cancer detection. Proceedings of the National Academy of Sciences 2008, 105 (30), 10513-10518. 5. Van der Pol, E.; Böing, A.; Gool, E.; Nieuwland, R., Recent developments in the nomenclature, presence, isolation, detection and clinical impact of extracellular vesicles. Journal of Thrombosis and Haemostasis 2016, 14 (1), 48-56. 6. Labib, M.; Sargent, E. H.; Kelley, S. O., Electrochemical methods for the analysis of clinically relevant biomolecules. Chemical Reviews 2016, 116 (16), 9001-9090. 7. Mattick, J. S., The central role of RNA in human development and cognition. FEBS Letter 2011, 585 (11), 1600-1616. 8. Sinden, R. R., DNA structure and function. Elsevier, 2012. 9. Zhang, J.; Li, S.; Li, L.; Li, M.; Guo, C.; Yao, J.; Mi, S., Exosome and exosomal microRNA: trafficking, sorting, and function. Genomics, Proteomics & Bioinformatics 2015, 13 (1), 17-24. 10. Wojdacz, T. K.; Møller, T. H.; Thestrup, B. B.; Kristensen, L. S.; Hansen, L. L., Limitations and advantages of MS-HRM and bisulfite sequencing for single locus methylation studies. Expert Review of Molecular Diagnostics 2010, 10 (5), 575-580.

11. Srinivasan, K.; Shiue, L.; Hayes, J. D.; Centers, R.; Fitzwater, S.; Loewen, R.; Edmondson, L. R.; Bryant, J.; Smith, M.; Rommelfanger, C., Detection and measurement of alternative splicing using splicing-sensitive microarrays. Methods 2005, 37 (4), 345- 359. 12. Fleige, S.; Pfaffl, M. W., RNA integrity and the effect on the real-time qRT-PCR performance. Molecular Aspects of Medicine 2006, 27 (2-3), 126-139. 13. Ozsolak, F.; Milos, P. M., RNA sequencing: advances, challenges and opportunities. Nature Reviews Genetics 2011, 12 (2), 87. 14. Ebert, M. P.; Korc, M.; Malfertheiner, P.; Röcken, C., Advances, challenges, and limitations in serum-proteome-based cancer diagnosis. Journal of Proteome Research 2006, 5 (1), 19-25. 15. Law, W.-C.; Yong, K.-T.; Baev, A.; Prasad, P. N., Sensitivity improved surface plasmon resonance biosensor for cancer biomarker detection based on plasmonic enhancement. ACS Nano 2011, 5 (6), 4858-4864. 16. Culha, M.; Stokes, D.; Allain, L. R.; Vo-Dinh, T., Surface-enhanced Raman scattering substrate based on a self-assembled monolayer for use in gene diagnostics. Analytical Chemistry 2003, 75 (22), 6196-6201. 17. Yang, J.; Xu, G.; Kong, H.; Zheng, Y.; Pang, T.; Yang, Q., Artificial neural network classification based on high-performance liquid chromatography of urinary and serum nucleosides for the clinical diagnosis of cancer. Journal of Chromatography B 2002, 780 (1), 27-33. 18. Kolch, W.; Neusüß, C.; Pelzing, M.; Mischak, H., Capillary electrophoresis–mass spectrometry as a powerful tool in clinical diagnosis and biomarker discovery. Mass Spectrometry Reviews 2005, 24 (6), 959-977. 19. Wang, J., Electrochemical biosensors: towards point-of-care cancer diagnostics. Biosensors and Bioelectronics . 2006, 21 (10), 1887-1892. 20. Hossain, T.; Mahmudunnabi, G.; Masud, M. K.; Islam, M. N.; Ooi, L.; Konstantinov, K.; Al Hossain, M. S.; Martinac, B.; Alici, G.; Nguyen, N.-T., Electrochemical biosensing strategies for DNA methylation analysis. Biosensors and Bioelectronics 2017. 21. Islam, M. N.; Masud, M. K.; Haque, M. H.; Hossain, M. S. A.; Yamauchi, Y.; Nguyen, N. T.; Shiddiky, M. J., RNA biomarkers: diagnostic and prognostic potentials and recent developments of electrochemical biosensors. Small Methods 2017, 1 (7), 1700131. 22. Maurer, J. J., Rapid detection and limitations of molecular techniques. Annual Review of Food Science and Technology 2011, 2, 259-279. 79

23. Cooper, G. R., Standard Methods of Clinical Chemistry: By the American Association of Clinical Chemists. Elsevier, 2013. 24. Chen, G.; Roy, I.; Yang, C.; Prasad, P. N., Nanochemistry and nanomedicine for nanoparticle-based diagnostics and therapy. Chemical Reviews 2016, 116 (5), 2826- 2885. 25. Steel, A. B.; Herne, T. M.; Tarlov, M. J., Electrochemical quantitation of DNA immobilized on gold. Analytical chemistry 1998, 70 (22), 4670-4677. 26. Shi, J.; Porterfield, D. M., Surface modification approaches for electrochemical biosensors. In Biosensors-Emerging Materials and Applications, InTech, 2011. 27. Li, J.; Tan, S. N.; Ge, H., Silica sol-gel immobilized amperometric biosensor for hydrogen peroxide. Analytica Chimica Acta 1996, 335 (1), 137-145. 28. Beveridge, J. S.; Stephens, J. R.; Williams, M. E., The use of magnetic nanoparticles in analytical chemistry. Annual Review of Analytical Chemistry 2011, 4, 251-273. 29. Reddy, L. H.; Arias, J. L.; Nicolas, J.; Couvreur, P., Magnetic nanoparticles: design and characterization, toxicity and biocompatibility, pharmaceutical and biomedical applications. Chemical Reviews 2012, 112 (11), 5818-5878. 30. Masud, M. K.; Kim, J.; Billah, M. M.; Wood, K.; Shiddiky, M. J.; Nguyen, N.-T.; Parsapur, R. K.; Kaneti, Y. V.; Alshehri, A. A.; Alghamidi, Y. G., Nanoarchitectured peroxidase-mimetic nanozymes: mesoporous nanocrystalline α-or γ-iron oxide? Journal of Materials Chemistry B 2019, 7 (35), 5412-5422. 31. Colombo, M.; Carregal-Romero, S.; Casula, M. F.; Gutiérrez, L.; Morales, M. P.; Böhm, I. B.; Heverhagen, J. T.; Prosperi, D.; Parak, W. J., Biological applications of magnetic nanoparticles. Chemical Society Reviews 2012, 41 (11), 4306-4334. 32. Neuberger, T.; Schöpf, B.; Hofmann, H.; Hofmann, M.; Von Rechenberg, B., Superparamagnetic nanoparticles for biomedical applications: possibilities and limitations of a new drug delivery system. Journal of Magnetism and Magnetic materials 2005, 293 (1), 483-496. 33. Banerjee, R.; Katsenovich, Y.; Lagos, L.; McIintosh, M.; Zhang, X.; Li, C.-Z., Nanomedicine: magnetic nanoparticles and their biomedical applications. Current Medicinal Chemistry 2010, 17 (27), 3120-3141. 34. Zhang, S.; Geryak, R.; Geldmeier, J.; Kim, S.; Tsukruk, V. V., Synthesis, assembly, and applications of hybrid nanostructures for biosensing. Chemical Reviews. 2017, 117 (20), 12942-13038.

35. Chen, A.; Chatterjee, S., Nanomaterials based electrochemical sensors for biomedical applications. Chemical Society Reviews 2013, 42 (12), 5425-5438. 36. Quesada-González, D.; Merkoçi, A., Nanomaterial-based devices for point-of-care diagnostic applications. Chemical Society Reviews 2018, 47 (13), 4697-4709. 37. Mishra, S.; Sharma, S.; Javed, M.; Pottoo, F.; Abul, M. B.; Amir, M.; Sarfaroz, M., Bioinspired Nanocomposites: Applications In Disease Diagnosis And Treatment. Pharmaceutical Nanotechnology 2019. 38. Zhu, X.; Li, J.; He, H.; Huang, M.; Zhang, X.; Wang, S., Application of nanomaterials in the bioanalytical detection of disease-related genes. Biosensors and Bioelectronics 2015, 74, 113-133. 39. Xia, N.; Zhang, L., Nanomaterials-based sensing strategies for electrochemical detection of microRNAs. Materials 2014, 7 (7), 5366-5384. 40. Masud, M. K.; Yadav, S.; Islam, M. N.; Nguyen, N. T.; Salomon, C.; Kline, R.; Alamri, H. R.; Alothman, Z. A.; Yamauchi, Y.; Hossain, M. S. A.; Shiddiky, M. J. A., Gold- Loaded Nanoporous Ferric Oxide Nanocubes with Peroxidase-Mimicking Activity for Electrocatalytic and Colorimetric Detection of Autoantibody. Analytical Chemistry 2017, 89 (20), 11005-11013. 41. Urbanova, V.; Magro, M.; Gedanken, A.; Baratella, D.; Vianello, F.; Zboril, R., Nanocrystalline iron oxides, composites, and related materials as a platform for electrochemical, magnetic, and chemical biosensors. Chemistry of Materials 2014, 26 (23), 6653-6673. 42. Azhar, A.; Li, Y.; Cai, Z.; Zakaria, M. B.; Masud, M. K.; Hossain, M. S. A.; Kim, J.; Zhang, W.; Na, J.; Yamauchi, Y., Nanoarchitectonics: A New Materials Horizon for Prussian Blue and Its Analogues. Bulletin of the Chemical Society of Japan 2019, 92 (4), 875-904. 43. Hao, R.; Xing, R.; Xu, Z.; Hou, Y.; Gao, S.; Sun, S., Synthesis, functionalization, and biomedical applications of multifunctional magnetic nanoparticles. Advanced Materials 2010, 22 (25), 2729-2742. 44. Ryckman, J. D.; Liscidini, M.; Sipe, J.; Weiss, S., Direct imprinting of porous substrates: a rapid and low-cost approach for patterning porous nanomaterials. Nano Letter 2010, 11 (5), 1857-1862. 45. Masud, M. K.; Islam, M. N.; Haque, M. H.; Tanaka, S.; Gopalan, V.; Alici, G.; Nguyen, N.-T.; Lam, A. K.; Hossain, M. S. A.; Yamauchi, Y., Gold-loaded nanoporous

superparamagnetic nanocubes for catalytic signal amplification in detecting miRNA. Chemical Communication 2017, 53 (58), 8231-8234. 46. Wei, Q.; Xiong, F.; Tan, S.; Huang, L.; Lan, E. H.; Dunn, B.; Mai, L., Porous one‐ dimensional nanomaterials: design, fabrication and applications in electrochemical energy storage. Advanced Materials 2017, 29 (20), 1602300. 47. Liu, J.; Qiao, S. Z.; Hu, Q. H.; Lu, G. Q., Magnetic nanocomposites with mesoporous structures: synthesis and applications. Small 2011, 7 (4), 425-443. 48. Lu, A. H.; Salabas, E. e. L.; Schüth, F., Magnetic nanoparticles: synthesis, protection, functionalization, and application. Angewandte Chemie International Edition 2007, 46 (8), 1222-1244. 49. Zhu, C.; Yang, G.; Li, H.; Du, D.; Lin, Y., Electrochemical sensors and biosensors based on nanomaterials and nanostructures. Analytical Chemistry2014, 87 (1), 230-249. 50. Xi, Z.; Ye, H.; Xia, X., Engineered noble-metal nanostructures for in vitro diagnostics. Chemistry of Materials 2018, 30 (23), 8391-8414. 51. Devi, S.; Tharmaraj, V., Nanomaterials for Advanced Analytical Applications in Chemo- and Biosensors. In Advanced Nanostructured Materials for Environmental Remediation, Springer: 2019, 91-110. 52. Yadav, H. M.; Otari, S. V., Hybrid Nanostructures in a Diagnostic and Comprehensive Approach to Combat Cancer. In Hybrid Nanostructures for Cancer Theranostics, Elsevier: 2019, 159-172. 53. Reina, G.; González-Domínguez, J. M.; Criado, A.; Vázquez, E.; Bianco, A.; Prato, M., Promises, facts and challenges for graphene in biomedical applications. Chemical Society Reviews 2017, 46 (15), 4400-4416. 54. Kumar, S.; Rani, R.; Dilbaghi, N.; Tankeshwar, K.; Kim, K.-H., Carbon nanotubes: a novel material for multifaceted applications in human healthcare. Chemical Society Reviews 2017, 46 (1), 158-196. 55. Xu, Y.; Wang, X.; Zhang, W. L.; Lv, F.; Guo, S., Recent progress in two-dimensional inorganic quantum dots. Chemical Society Reviews 2018, 47 (2), 586-625. 56. Masud, M. K.; Umer, M.; Hossain, M. S. A.; Yamauchi, Y.; Nguyen, N.-T.; Shiddiky, M. J., Nanoarchitecture frameworks for electrochemical miRNA detection. Trends in Biochemical Sciences 2019. 57. Wang, J.; Ma, Q.; Wang, Y.; Li, Z.; Li, Z.; Yuan, Q., New insights into the structure– performance relationships of mesoporous materials in analytical science. Chemical Society Reviews 2018, 47 (23), 8766-8803. 82

58. Massart, R., Preparation of aqueous magnetic liquids in alkaline and acidic media. IEEE Transactions on Magnetics 1981, 17 (2), 1247-1248. 59. Wu, W.; He, Q.; Jiang, C., Magnetic iron oxide nanoparticles: synthesis and surface functionalization strategies. Nanoscale Research Letters 2008, 3 (11), 397. 60. Sugimoto, T.; Matijević, E., Formation of uniform spherical magnetite particles by crystallization from ferrous hydroxide gels. Journal of Colloid and Interface Science 1980, 74 (1), 227-243.

61. Wu, S.; Sun, A.; Zhai, F.; Wang, J.; Xu, W.; Zhang, Q.; Volinsky, A. A., Fe3O4 magnetic nanoparticles synthesis from tailings by ultrasonic chemical co-precipitation. Materials Letters 2011, 65 (12), 1882-1884. 62. Pereira, C.; Pereira, A. M.; Fernandes, C.; Rocha, M.; Mendes, R.; Fernández-García, M. P.; Guedes, A.; Tavares, P. B.; Grenèche, J.-M.; Araújo, J. o. P., Superparamagnetic

MFe2O4 (M= Fe, Co, Mn) nanoparticles: tuning the particle size and magnetic properties through a novel one-step coprecipitation route. Chemistry of Materials 2012, 24 (8), 1496-1504. 63. Karaagac, O.; Yildiz, B. B.; Köçkar, H., The influence of synthesis parameters on one- step synthesized superparamagnetic cobalt ferrite nanoparticles with high saturation magnetization. Journal of Magnetism and Magnetic Materials 2019, 473, 262-267. 64. Goon, I. Y.; Zhang, C.; Lim, M.; Gooding, J. J.; Amal, R., Controlled fabrication of polyethylenimine-functionalized magnetic nanoparticles for the sequestration and quantification of free Cu2+. Langmuir 2010, 26 (14), 12247-12252. 65. Suh, S. K.; Yuet, K.; Hwang, D. K.; Bong, K. W.; Doyle, P. S.; Hatton, T. A., Synthesis of nonspherical superparamagnetic particles: in situ coprecipitation of magnetic nanoparticles in microgels prepared by stop-flow lithography. J. Am. Chem. Soc. 2012, 134 (17), 7337-7343. 66. Wu, W.; Wu, Z.; Yu, T.; Jiang, C.; Kim, W.-S., Recent progress on magnetic iron oxide nanoparticles: synthesis, surface functional strategies and biomedical applications. Science and Technology of Advanced Materials 2015, 16 (2), 023501. 67. Park, J.; An, K.; Hwang, Y.; Park, J.-G.; Noh, H.-J.; Kim, J.-Y.; Park, J.-H.; Hwang, N.- M.; Hyeon, T., Ultra-large-scale syntheses of monodisperse nanocrystals. Nature Materials 2004, 3 (12), 891. 68. Ali, A.; Hira Zafar, M. Z.; ul Haq, I.; Phull, A. R.; Ali, J. S.; Hussain, A., Synthesis, characterization, applications, and challenges of iron oxide nanoparticles. Nanotechnology, Science and Applications 2016, 9, 49. 83

69. Xie, W.; Guo, Z.; Gao, F.; Gao, Q.; Wang, D.; Liaw, B.-s.; Cai, Q.; Sun, X.; Wang, X.; Zhao, L., Shape-, size-and structure-controlled synthesis and biocompatibility of iron oxide nanoparticles for magnetic theranostics. Theranostics 2018, 8 (12), 3284. 70. Majidi, S.; Zeinali Sehrig, F.; Farkhani, S. M.; Soleymani Goloujeh, M.; Akbarzadeh, A., Current methods for synthesis of magnetic nanoparticles. Artificial Cells, Nanomedicine, and Biotechnology 2016, 44 (2), 722-734. 71. Stojanovic, B. D.; Dzunuzovic, A. S.; Ilic, N. I., Review of methods for the preparation of magnetic metal oxides. In Magnetic, Ferroelectric, and Multiferroic Metal Oxides, Elsevier, 2018, 333-359. 72. Del Monte, F.; Morales, M.; Levy, D.; Fernandez, A.; Ocana, M.; Roig, A.; Molins, E.;

O'Grady, K.; Serna, C., Formation of γ-Fe2O3 isolated nanoparticles in a silica matrix. Langmuir 1997, 13 (14), 3627-3634. 73. Khan, I.; Saeed, K.; Khan, I., Nanoparticles: Properties, applications and toxicities. Arabian Journal of Chemistry 2017. 74. Gupta, A. K.; Gupta, M., Synthesis and surface engineering of iron oxide nanoparticles for biomedical applications. Biomaterials 2005, 26 (18), 3995-4021. 75. Huczko, A., Template-based synthesis of nanomaterials. Applied Physics A 2000, 70 (4), 365-376. 76. Wade, T.; Wegrowe, J.-E., Template synthesis of nanomaterials. The European Physical Journal-Applied Physics 2005, 29 (1), 3-22. 77. Thomas, A.; Goettmann, F.; Antonietti, M., Hard templates for soft materials: creating nanostructured organic materials. Chemistry of Materials 2008, 20 (3), 738-755. 78. Wu, H.; Wu, G.; Ren, Y.; Li, X.; Wang, L., Multishelled Metal Oxide Hollow Spheres: Easy Synthesis and Formation Mechanism. Chemistry – A European Journal 2016, 22 (26), 8864-8871. 79. Sasidharan, M.; Liu, D.; Gunawardhana, N.; Yoshio, M.; Nakashima, K., Synthesis, characterization and application for lithium-ion rechargeable batteries of hollow silica nanospheres. J. Mater. Chem. 2011, 21 (36), 13881-13888. 80. Tang, Y.; Chen, Y.; Zhou, P.; Zhou, Y.; Lu, L.; Bao, J.; Lu, T., Electro-catalytic performance of PdCo bimetallic hollow nano-spheres for the oxidation of formic acid. Journal of Solid State Electrochemistry 2010, 14 (11), 2077-2082. 81. Liu, Q.; Liu, H.; Han, M.; Zhu, J.; Liang, Y.; Xu, Z.; Song, Y., Nanometer-Sized Nickel Hollow Spheres. Advanced Materials 2005, 17 (16), 1995-1999.

82. Wang, G.-H.; Sun, Q.; Zhang, R.; Li, W.-C.; Zhang, X.-Q.; Lu, A.-H., Weak acid–base interaction induced assembly for the synthesis of diverse hollow nanospheres. Chemistry of Materials 2011, 23 (20), 4537-4542. 83. Purbia, R.; Paria, S., Yolk/shell nanoparticles: classifications, synthesis, properties, and applications. Nanoscale 2015, 7 (47), 19789-19873. 84. Sun, Q.; Guo, C. Z.; Wang, G. H.; Li, W. C.; Bongard, H. J.; Lu, A. H., Fabrication of magnetic yolk–shell nanocatalysts with spatially resolved functionalities and high activity for nitrobenzene hydrogenation. Chemistry–A European Journal 2013, 19 (20), 6217-6220. 85. Wang, B.; Wang, G.; Wang, H., Synthesis and electrochemical investigation of hollow hierarchical metal oxide microspheres for high performance lithium-ion batteries. Electrochimica Acta 2015, 156, 1-10. 86. Xiang, Y.; Lu, S.; Jiang, S. P., Layer-by-layer self-assembly in the development of electrochemical energy conversion and storage devices from fuel cells to supercapacitors. Chemical Society Reviews 2012, 41 (21), 7291-7321. 87. Kamata, H.; Li, X.; Chung, U.-i.; Sakai, T., Design of Hydrogels for Biomedical Applications. Advanced Healthcare Materials 2015, 4 (16), 2360-2374. 88. Prieto, G.; Tüysüz, H.; Duyckaerts, N.; Knossalla, J.; Wang, G.-H.; Schüth, F., Hollow Nano- and Microstructures as Catalysts. Chemical Reviews. 2016, 116 (22), 14056- 14119. 89. Liu, Y.; Goebl, J.; Yin, Y., Templated synthesis of nanostructured materials. Chemical Society Reviews 2013, 42 (7), 2610-2653. 90. Zhou, H.-C.; Long, J. R.; Yaghi, O. M., Introduction to metal–organic frameworks. ACS Publications, 2012. 91. Hu, M.; Belik, A. A.; Imura, M.; Mibu, K.; Tsujimoto, Y.; Yamauchi, Y., Synthesis of superparamagnetic nanoporous iron oxide particles with hollow interiors by using prussian blue coordination polymers. Chemistry of Materials 2012, 24 (14), 2698-2707. 92. Hu, M.; Jiang, J.-S.; Zeng, Y., Prussian blue microcrystals prepared by selective etching and their conversion to mesoporous magnetic iron (III) oxides. Chemical Communication 2010, 46 (7), 1133-1135. 93. Chiang, Y. D.; Hu, M.; Kamachi, Y.; Ishihara, S.; Takai, K.; Tsujimoto, Y.; Ariga, K.; Wu, K. C. W.; Yamauchi, Y., Rational Design and Synthesis of Cyano‐ Bridged Coordination Polymers with Precise Control of Particle Size from 20 to 500 nm. European Journal of Inorganic Chemistry 2013, 2013 (18), 3141-3145. 85

94. Zeng, H.; Sun, S., Syntheses, Properties, and Potential Applications of Multicomponent Magnetic Nanoparticles. Advanced Functional Materials 2008, 18 (3), 391-400. 95. Vallabani, N. V. S.; Singh, S., Recent advances and future prospects of iron oxide nanoparticles in biomedicine and diagnostics. 3 Biotech 2018, 8 (6), 279-279. 96. Silva, S. M.; Tavallaie, R.; Sandiford, L.; Tilley, R. D.; Gooding, J. J., Gold coated magnetic nanoparticles: from preparation to surface modification for analytical and biomedical applications. Chemical communications 2016, 52 (48), 7528-7540. 97. Hu, P.; Zhang, S.; Wu, T.; Ni, D.; Fan, W.; Zhu, Y.; Qian, R.; Shi, J., Fe–Au Nanoparticle‐Coupling for Ultrasensitive Detections of Circulating Tumor DNA. Advanced Materials 2018, 30 (31), 1801690. 98. Leung, K. C.-F.; Xuan, S.; Zhu, X.; Wang, D.; Chak, C.-P.; Lee, S.-F.; Ho, W. K.-W.; Chung, B. C.-T., Gold and iron oxide hybrid nanocomposite materials. Chemical Society Reviews 2012, 41 (5), 1911-1928. 99. Erathodiyil, N.; Ying, J. Y., Functionalization of Inorganic Nanoparticles for Bioimaging Applications. Accounts of Chemical Research 2011, 44 (10), 925-935. 100. Chin, S. F.; Iyer, K. S.; Raston, C. L., Facile and Green Approach To Fabricate Gold and Silver Coated Superparamagnetic Nanoparticles. Crystal Growth & Design 2009, 9 (6), 2685-2689.

101. Teymourian, H.; Salimi, A.; Khezrian, S., Fe3O4 magnetic nanoparticles/reduced graphene oxide nanosheets as a novel electrochemical and bioeletrochemical sensing platform. Biosensors and Bioelectronics 2013, 49, 1-8. 102. Wu, S.; He, Q.; Zhou, C.; Qi, X.; Huang, X.; Yin, Z.; Yang, Y.; Zhang, H., Synthesis of Fe 3 O 4 and Pt nanoparticles on reduced graphene oxide and their use as a recyclable catalyst. Nanoscale 2012, 4 (7), 2478-2483. 103. Tanaka, S.; Salunkhe, R. R.; Kaneti, Y. V.; Malgras, V.; Alshehri, S. M.; Ahamad, T.; Zakaria, M. B.; Dou, S. X.; Yamauchi, Y.; Hossain, M. S. A., Prussian blue derived iron oxide nanoparticles wrapped in graphene oxide sheets for electrochemical supercapacitors. RSC Advances 2017, 7 (54), 33994-33999. 104. Cao, Y., Preparation and magnetic properties of a multi-walled carbon nanotube-iron oxide nanoparticle composite. Fullerenes, Nanotubes and Carbon Nanostructures 2015, 23 (7), 623-626. 105. Xu, L.; Liang, H.-W.; Yang, Y.; Yu, S.-H., Stability and reactivity: positive and negative aspects for nanoparticle processing. Chemical Reviews 2018, 118 (7), 3209-3250. 86

106. Cardoso, V. F.; Francesko, A.; Ribeiro, C.; Bañobre‐López, M.; Martins, P.; Lanceros‐ Mendez, S., Advances in magnetic nanoparticles for biomedical applications. Advanced Healthcare Materials 2018, 7 (5), 1700845. 107. Huang, X.; Schmucker, A.; Dyke, J.; Hall, S. M.; Retrum, J.; Stein, B.; Remmes, N.; Baxter, D. V.; Dragnea, B.; Bronstein, L. M., Magnetic nanoparticles with functional silanes: evolution of well-defined shells from anhydride containing silane. Journal of Materials Chemistry 2009, 19 (24), 4231-4239. 108. Shen, M.; Cai, H.; Wang, X.; Cao, X.; Li, K.; Wang, S. H.; Guo, R.; Zheng, L.; Zhang, G.; Shi, X., Facile one-pot preparation, surface functionalization, and toxicity assay of APTS-coated iron oxide nanoparticles. Nanotechnology 2012, 23 (10), 105601. 109. Li, D.; Jiang, D.; Chen, M.; Xie, J.; Wu, Y.; Dang, S.; Zhang, J., An easy fabrication of monodisperse oleic acid-coated Fe 3 O 4 nanoparticles. Materials Letters 2010, 64 (22), 2462-2464. 110. Lartigue, L.; Oumzil, K.; Guari, Y.; Larionova, J.; Guérin, C.; Montero, J.-L.; Barragan- Montero, V.; Sangregorio, C.; Caneschi, A.; Innocenti, C., Water-soluble rhamnose- coated Fe3O4 nanoparticles. Organic letters 2009, 11 (14), 2992-2995. 111. Chalasani, R.; Vasudevan, S., Cyclodextrin functionalized magnetic iron oxide nanocrystals: a host-carrier for magnetic separation of non-polar molecules and arsenic from aqueous media. Journal of Materials Chemistry 2012, 22 (30), 14925-14931. 112. Gao, J.; Ran, X.; Shi, C.; Cheng, H.; Cheng, T.; Su, Y., One-step solvothermal synthesis

of highly water-soluble, negatively charged superparamagnetic Fe3O4 colloidal nanocrystal clusters. Nanoscale 2013, 5 (15), 7026-7033. 113. Lattuada, M.; Hatton, T. A., Functionalization of monodisperse magnetic nanoparticles. Langmuir 2007, 23 (4), 2158-2168. 114. Ghosh Chaudhuri, R.; Paria, S., Core/shell nanoparticles: classes, properties, synthesis mechanisms, characterization, and applications. Chemical Reviews 2011, 112 (4), 2373- 2433. 115. Oh, J. K.; Park, J. M., Iron oxide-based superparamagnetic polymeric nanomaterials: design, preparation, and biomedical application. Progress in Polymer Science 2011, 36 (1), 168-189. 116. El-Sherif, H.; El-Masry, M.; Emira, H. S., Magnetic polymer composite particles via in situ inverse miniemulsion polymerization process. Journal of Macromolecular Science, Part A: Pure and Applied Chemistry 2010, 47 (11), 1096-1103.

117. Pimpha, N.; Chaleawlert-umpon, S.; Sunintaboon, P., Core/shell polymethyl methacrylate/polyethyleneimine particles incorporating large amounts of iron oxide nanoparticles prepared by emulsifier-free emulsion polymerization. Polymer 2012, 53 (10), 2015-2022. 118. Jarzyna, P. A.; Skajaa, T.; Gianella, A.; Cormode, D. P.; Samber, D. D.; Dickson, S. D.; Chen, W.; Griffioen, A. W.; Fayad, Z. A.; Mulder, W. J., Iron oxide core oil-in-water emulsions as a multifunctional nanoparticle platform for tumor targeting and imaging. Biomaterials 2009, 30 (36), 6947-6954. 119. Lungu, I. I.; Radulescu, M.; Mogosanu, G. D.; Grumezescu, A. M., pH sensitive core-shell magnetic nanoparticles for targeted drug delivery in cancer therapy. Romanian Journal of Morphology and Embryology 2016, 57 (1), 23-32. 120. Ghorbani, M.; Hamishehkar, H.; Arsalani, N.; Entezami, A. A., Preparation of thermo

and pH-responsive polymer@ Au/Fe3O4 core/shell nanoparticles as a carrier for delivery of anticancer agent. Journal of Nanoparticle Research 2015, 17 (7), 305. 121. Dai, S.; Ravi, P.; Tam, K. C., pH-Responsive polymers: synthesis, properties and applications. Soft Matter 2008, 4 (3), 435-449. 122. Grancharov, S. G.; Zeng, H.; Sun, S.; Wang, S. X.; O'Brien, S.; Murray, C.; Kirtley, J.; Held, G., Bio-functionalization of monodisperse magnetic nanoparticles and their use as biomolecular labels in a magnetic tunnel junction based sensor. The Journal of Physical Chemistry B 2005, 109 (26), 13030-13035. 123. You, C.-C.; De, M.; Rotello, V. M., Monolayer-protected nanoparticle–protein interactions. Current Opinion in Chemical Biology 2005, 9 (6), 639-646. 124. Katz, E.; Willner, I., Integrated nanoparticle–biomolecule hybrid systems: synthesis, properties, and applications. Angewandte Chemie International Edition 2004, 43 (45), 6042-6108. 125. Song, X.-R.; Goswami, N.; Yang, H.-H.; Xie, J., Functionalization of metal nanoclusters for biomedical applications. Analyst 2016, 141 (11), 3126-3140.

126. Lee, C.; Huang, K.; Wei, P.; Yao, Y., Conjugation of γ-Fe2O3 nanoparticles with single strand oligonucleotides. Journal of Magnetism and Magnetic Materials 2006, 304 (1), e412-e414. 127. McCarthy, J. R.; Weissleder, R., Multifunctional magnetic nanoparticles for targeted imaging and therapy. Advanced drug delivery Reviews 2008, 60 (11), 1241-1251. 128. Veiseh, O.; Sun, C.; Fang, C.; Bhattarai, N.; Gunn, J.; Kievit, F.; Du, K.; Pullar, B.; Lee, D.; Ellenbogen, R. G., Specific targeting of brain tumors with an optical/magnetic 88

resonance imaging nanoprobe across the blood-brain barrier. Cancer Research 2009, 69 (15), 6200-6207. 129. Laurent, S.; Forge, D.; Port, M.; Roch, A.; Robic, C.; Vander Elst, L.; Muller, R. N., Magnetic iron oxide nanoparticles: synthesis, stabilization, vectorization, physicochemical characterizations, and biological applications. Chemical Reviews 2008, 108 (6), 2064-2110. 130. Kluchova, K.; Zboril, R.; Tucek, J.; Pecova, M.; Zajoncova, L.; Safarik, I.; Mashlan, M.; Markova, I.; Jancik, D.; Sebela, M., Superparamagnetic maghemite nanoparticles from solid-state synthesis–their functionalization towards peroral MRI contrast agent and magnetic carrier for trypsin immobilization. Biomaterials 2009, 30 (15), 2855-2863.

131. Wang, G.; Jin, L.; Dong, Y.; Niu, L.; Liu, Y.; Ren, F.; Su, X., Multifunctional Fe3O4–

CdTe@SiO2–carboxymethyl chitosan drug nanocarriers: synergistic effect towards magnetic targeted drug delivery and cell imaging. New Journal of Chemistry 2014, 38 (2), 700-708. 132. Gowd, G. S.; Patra, M. K.; Mathew, M.; Shukla, A.; Songara, S.; Vadera, S. R.; Kumar, 3+ N., Synthesis of Fe3O4@Y2O3: Eu core–shell multifunctional nanoparticles and their magnetic and luminescence properties. Optical Materials 2013, 35 (9), 1685-1692. 133. Koo, K. M.; Sina, A. A.; Carrascosa, L. G.; Shiddiky, M. J.; Trau, M., DNA–bare gold affinity interactions: mechanism and applications in biosensing. Analytical Methods 2015, 7 (17), 7042-7054. 134. Haque, M. H.; Gopalan, V.; Islam, M. N.; Masud, M. K.; Bhattacharjee, R.; Al Hossain, M. S.; Nguyen, N.-T.; Lam, A. K.; Shiddiky, M. J., Quantification of gene-specific DNA methylation in oesophageal cancer via electrochemistry. Analytica Chimica Acta 2017, 976, 84-93. 135. Hosu, O.; Florea, A.; Cristea, C.; Sandulescu, R., Functionalized Advanced Hybrid Materials for Biosensing Applications. In Advanced Biosensors for Health Care Applications, Elsevier, 2019, 171-207. 136. Yi, D. K.; Selvan, S. T.; Lee, S. S.; Papaefthymiou, G. C.; Kundaliya, D.; Ying, J. Y., Silica-coated nanocomposites of magnetic nanoparticles and quantum dots. Journal of American Chemical Society 2005, 127 (14), 4990-4991. 137. Stöber, W.; Fink, A.; Bohn, E., Controlled growth of monodisperse silica spheres in the micron size range. Journal of Colloid and Interface Science 1968, 26 (1), 62-69. 138. Xuan, S.-h.; Lee, S.-F.; Lau, J. T.-F.; Zhu, X.; Wang, Y.-X. J.; Wang, F.; Lai, J. M.; Sham, K. W.; Lo, P.-C.; Yu, J. C., Photocytotoxicity and magnetic relaxivity responses 89

of dual-porous γ-Fe2O3@ meso-SiO2 microspheres. ACS Applied Materials & Interfaces 2012, 4 (4), 2033-2040. 139. Park, J.-N.; Zhang, P.; Hu, Y.-S.; McFarland, E. W., Synthesis and characterization of

sintering-resistant silica-encapsulated Fe3O4 magnetic nanoparticles active for oxidation and chemical looping combustion. Nanotechnology 2010, 21 (22), 225708.

140. Li, Y.; Hu, Y.; Jiang, H.; Li, C., Double-faced γ-Fe2O3||SiO2 nanohybrids: flame synthesis, in situ selective modification and highly interfacial activity. Nanoscale 2013, 5 (12), 5360-5367. 141. Cong, H. P.; He, J. J.; Lu, Y.; Yu, S. H., Water‐Soluble Magnetic‐Functionalized Reduced Graphene Oxide Sheets: In situ Synthesis and Magnetic Resonance Imaging Applications. Small 2010, 6 (2), 169-173. 142. Dreaden, E. C.; Alkilany, A. M.; Huang, X.; Murphy, C. J.; El-Sayed, M. A., The golden age: gold nanoparticles for biomedicine. Chemical Society Reviews 2012, 41 (7), 2740- 2779. 143. Zhang, J.; Thurber, A.; Hanna, C.; Punnoose, A., Highly shape-selective synthesis, silica coating, self-assembly, and magnetic hydrogen sensing of hematite nanoparticles. Langmuir 2009, 26 (7), 5273-5278. 144. Wang, L.; Luo, J.; Maye, M. M.; Fan, Q.; Rendeng, Q.; Engelhard, M. H.; Wang, C.; Lin, Y.; Zhong, C.-J., Iron oxide–gold core–shell nanoparticles and thin film assembly. Journal of Materials Chemistry 2005, 15 (18), 1821-1832. 145. Wu, W.; Zhang, S.; Xiao, X.; Zhou, J.; Ren, F.; Sun, L.; Jiang, C., Controllable synthesis, magnetic properties, and enhanced photocatalytic activity of spindlelike mesoporous α- Fe2O3/ZnO core–shell heterostructures. ACS Applied Materials & Interfaces 2012, 4 (7), 3602-3609. 146. Sun, L.; Wu, W.; Zhang, S.; Liu, Y.; Xiao, X.; Ren, F.; Cai, G.; Jiang, C., Spindle-Like

α-Fe2O3 Embedded with TiO2 Nanocrystalline: Ion Implantation Preparation and Enhanced Magnetic Properties. Journal of Nanoscience and Nanotechnology 2013, 13 (8), 5428-5433. 147. Li, S.-K.; Huang, F.-Z.; Wang, Y.; Shen, Y.-H.; Qiu, L.-G.; Xie, A.-J.; Xu, S.-J.,

Magnetic Fe3O4@C@Cu2O composites with bean-like core/shell nanostructures: synthesis, properties and application in recyclable photocatalytic degradation of dye pollutants. Journal of Materials Chemistry 2011, 21 (20), 7459-7466.

148. Liu, L.; Xiao, L.; Zhu, H.-Y.; Shi, X.-W., Studies on interaction and illumination damage of CS-Fe3O4@ ZnS: Mn to bovine serum albumin. Journal of Nanoparticle Research 2013, 15 (1), 1394. 149. Li, Y.; Liu, Y.; Tang, J.; Lin, H.; Yao, N.; Shen, X.; Deng, C.; Yang, P.; Zhang, X.,

Fe3O4@Al2O3 magnetic core–shell microspheres for rapid and highly specific capture of phosphopeptides with mass spectrometry analysis. Journal of Chromatography A 2007, 1172 (1), 57-71.

150. Wang, D.; He, J.; Rosenzweig, N.; Rosenzweig, Z., Superparamagnetic Fe2O3 beads− CdSe/ZnS quantum dots core− shell nanocomposite particles for cell separation. Nano Letters 2004, 4 (3), 409-413. 151. Chen, F.; Shi, R.; Xue, Y.; Chen, L.; Wan, Q.-H., Templated synthesis of monodisperse mesoporous maghemite/silica microspheres for magnetic separation of genomic DNA. Journal of Magnetism and Magnetic Materials 2010, 322 (16), 2439-2445. 152. Yildiz, I., Applications of magnetic nanoparticles in biomedical separation and purification. Nanotechnology Reviews 2016, 5 (3), 331-340. 153. Franzreb, M.; Siemann-Herzberg, M.; Hobley, T. J.; Thomas, O. R., Protein purification using magnetic adsorbent particles. Applied Microbiology and Biotechnology 2006, 70 (5), 505-516. 154. Fatima, H.; Kim, K.-S., Magnetic nanoparticles for bioseparation. Korean Journal of Chemical Engineering 2017, 34 (3), 589-599. 155. Min, J. H.; Woo, M.-K.; Yoon, H. Y.; Jang, J. W.; Wu, J. H.; Lim, C.-S.; Kim, Y. K., Isolation of DNA using magnetic nanoparticles coated with dimercaptosuccinic acid. Analytical Biochemistry 2014, 447, 114-118. 156. Křížová, J.; Španová, A.; Rittich, B.; Horák, D., Magnetic hydrophilic methacrylate- based polymer microspheres for genomic DNA isolation. Journal of Chromatography A 2005, 1064 (2), 247-253. 157. Safarik, I.; Safarikova, M., Magnetic techniques for the isolation and purification of proteins and peptides. BioMagnetic Research and Technology 2004, 2 (1), 7. 158. Wittrup, A.; Zhang, S.-H.; Svensson, K. J.; Kucharzewska, P.; Johansson, M. C.; Mörgelin, M.; Belting, M., Magnetic nanoparticle-based isolation of endocytic vesicles reveals a role of the heat shock protein GRP75 in macromolecular delivery. Proceedings of the National Academy of Sciences 2010, 107 (30), 13342-13347.

159. Mandal, S. M.; Ghosh, A. K.; Mandal, M., Iron oxide nanoparticle assisted purification and mass spectrometry based proteolytic mapping of intact CD4+ T cells from human blood. Preparative Biochemistry and Biotechnology 2008, 39 (1), 20-31. 160. Herr, J. K.; Smith, J. E.; Medley, C. D.; Shangguan, D.; Tan, W., Aptamer-conjugated nanoparticles for selective collection and detection of cancer cells. Analytical Chemistry 2006, 78 (9), 2918-2924. 161. Song, E.-Q.; Wang, G.-P.; Xie, H.-Y.; Zhang, Z.-L.; Hu, J.; Peng, J.; Wu, D.-C.; Shi, Y.- B.; Pang, D.-W., Visual recognition and efficient isolation of apoptotic cells with fluorescent-magnetic-biotargeting multifunctional nanospheres. Clinical Chemistry 2007, 53 (12), 2177-2185. 162. Van Engeland, M.; Nieland, L. J.; Ramaekers, F. C.; Schutte, B.; Reutelingsperger, C. P., Annexin V-affinity assay: a Review on an apoptosis detection system based on phosphatidylserine exposure. Cytometry 1998, 31 (1), 1-9. 163. Pamme, N.; Eijkel, J. C.; Manz, A., On-chip free-flow magnetophoresis: Separation and detection of mixtures of magnetic particles in continuous flow. Journal of Magnetism and Magnetic Materials 2006, 307 (2), 237-244. 164. Chen, G. D.; Alberts, C. J.; Rodriguez, W.; Toner, M., Concentration and purification of human immunodeficiency virus type 1 virions by microfluidic separation of superparamagnetic nanoparticles. Analytical Chemistry 2009, 82 (2), 723-728. 165. Kilianová, M.; Prucek, R.; Filip, J.; Kolařík, J.; Kvítek, L.; Panáček, A.; Tuček, J.; Zbořil, R., Remarkable efficiency of ultrafine superparamagnetic iron (III) oxide nanoparticles toward arsenate removal from aqueous environment. Chemosphere 2013, 93 (11), 2690- 2697. 166. Hayat, A.; Catanante, G.; Marty, J. L., Current trends in nanomaterial-based amperometric biosensors. Sensors 2014, 14 (12), 23439-23461. 167. Ansari, S. A.; Husain, Q., Potential applications of enzymes immobilized on/in nano materials: a Review. Biotechnology Advances 2012, 30 (3), 512-523. 168. Johnson, A. K.; Zawadzka, A. M.; Deobald, L. A.; Crawford, R. L.; Paszczynski, A. J., Novel method for immobilization of enzymes to magnetic nanoparticles. Journal of Nanoparticle Research 2008, 10 (6), 1009-1025. 169. Mitchell, D. T.; Lee, S. B.; Trofin, L.; Li, N.; Nevanen, T. K.; Söderlund, H.; Martin, C. R., Smart nanotubes for bioseparations and biocatalysis. Journal of the American Chemical Society 2002, 124 (40), 11864-11865.

170. Cipolatti, E. P.; Silva, M. J. A.; Klein, M.; Feddern, V.; Feltes, M. M. C.; Oliveira, J. V.; Ninow, J. L.; de Oliveira, D., Current status and trends in enzymatic nanoimmobilization. Journal of Molecular Catalysis B: Enzymatic 2014, 99, 56-67. 171. Choi, J.; Lee, J. I.; Lee, Y. B.; Hong, J. H.; Kim, I. S.; Park, Y. K.; Hur, N. H., Immobilization of biomolecules on biotinylated magnetic ferrite nanoparticles. Chemical Physics Letters 2006, 428 (1), 125-129. 172. Xing, Z.-C.; Chang, Y.; Kang, I.-K., Immobilization of biomolecules on the surface of inorganic nanoparticles for biomedical applications. Science and Technology of Advanced Materials 2010, 11 (1), 014101. 173. Stanciu, L.; Won, Y.-H.; Ganesana, M.; Andreescu, S., Magnetic particle-based hybrid platforms for bioanalytical sensors. Sensors 2009, 9 (4), 2976-2999. 174. Kouassi, G. K.; Irudayaraj, J., Magnetic and gold-coated magnetic nanoparticles as a DNA sensor. Analytical chemistry 2006, 78 (10), 3234-3241. 175. Kudr, J.; Haddad, Y.; Richtera, L.; Heger, Z.; Cernak, M.; Adam, V.; Zitka, O., Magnetic Nanoparticles: From Design and Synthesis to Real World Applications. Nanomaterials 2017, 7 (9), 243. 176. Ahmad, R.; Sardar, M., Enzyme immobilization: an overview on nanoparticles as immobilization matrix. Biochemistry and Analytical Biochemistry 2015, 4 (2), 1. 177. Tiefenauer, L. X.; Kuehne, G.; Andres, R. Y., Antibody-magnetite nanoparticles: in vitro characterization of a potential tumor-specific contrast agent for magnetic resonance imaging. Bioconjugate Chemistry 1993, 4 (5), 347-352. 178. Josephson, L.; Tung, C.-H.; Moore, A.; Weissleder, R., High-efficiency intracellular magnetic labeling with novel superparamagnetic-Tat peptide conjugates. Bioconjugate Chemistry 1999, 10 (2), 186-191. 179. Sestier, C.; Da‐Silva, M. F.; Sabolovic, D.; Roger, J.; Pons, J. N., Surface modification of superparamagnetic nanoparticles (ferrofluid) studied with particle electrophoresis: Application to the specific targeting of cells. Electrophoresis 1998, 19 (7), 1220-1226. 180. Wang, T.-H.; Lee, W.-C., Immobilization of proteins on magnetic nanoparticles. Biotechnology and Bioprocess Engineering 2003, 8 (4), 263-267. 181. Di Marco, M.; Shamsuddin, S.; Razak, K. A.; Aziz, A. A.; Devaux, C.; Borghi, E.; Levy, L.; Sadun, C., Overview of the main methods used to combine proteins with nanosystems: absorption, bioconjugation, and encapsulation. International Journal of Nanomedicine 2010, 5, 37.

182. Qing, T.; He, D.; He, X.; Wang, K.; Xu, F.; Wen, L.; Shangguan, J.; Mao, Z.; Lei, Y., Nucleic acid tool enzymes-aided signal amplification strategy for biochemical analysis: status and challenges. Analytical and Bioanalytical chemistry 2016, 408 (11), 2793- 2811. 183. Lei, J.; Ju, H., Signal amplification using functional nanomaterials for biosensing. Chemical Society Reviews 2012, 41 (6), 2122-2134. 184. Cannon, B.; Campos, A. R.; Lewitz, Z.; Willets, K. A.; Russell, R., Zeptomole detection of DNA nanoparticles by single-molecule fluorescence with magnetic field-directed localization. Analytical Biochemistry 2012, 431 (1), 40-47. 185. Tel-Vered, R.; Bard, A. J., Generation and detection of single metal nanoparticles using scanning electrochemical microscopy techniques. The Journal of Physical Chemistry B 2006, 110 (50), 25279-25287. 186. Krishnan, S.; Mani, V.; Wasalathanthri, D.; Kumar, C. V.; Rusling, J. F., Attomolar detection of a cancer biomarker protein in serum by surface plasmon resonance using superparamagnetic particle labels. Angewandte Chemie International Edition 2011, 50 (5), 1175-1178. 187. Zhuo, Y.; Yuan, P.-X.; Yuan, R.; Chai, Y.-Q.; Hong, C.-L., Bienzyme functionalized three-layer composite magnetic nanoparticles for electrochemical immunosensors. Biomaterials 2009, 30 (12), 2284-2290. 188. Bi, S.; Yan, Y.; Yang, X.; Zhang, S., Gold Nanolabels for New Enhanced Chemiluminescence Immunoassay of Alpha-Fetoprotein Based on Magnetic Beads. Chemistry – A European Journal 2009, 15 (18), 4704-4709. 189. Tang, D.; Su, B.; Tang, J.; Ren, J.; Chen, G., Nanoparticle-based sandwich electrochemical immunoassay for carbohydrate antigen 125 with signal enhancement using enzyme-coated nanometer-sized enzyme-doped silica beads. Analytical Chemistry 2010, 82 (4), 1527-1534. 190. Pan, J.; Yang, Q., Antibody-functionalized magnetic nanoparticles for the detection of carcinoembryonic antigen using a flow-injection electrochemical device. Analytical and Bioanalytical Chemistry 2007, 388 (1), 279-286. 191. Ahmed, A.; Rushworth, J. V.; Hirst, N. A.; Millner, P. A., Biosensors for whole-cell bacterial detection. Clinical microbiology Reviews 2014, 27 (3), 631-646. 192. Bi, S.; Zhou, H.; Zhang, S., Bio-bar-code functionalized magnetic nanoparticle label for ultrasensitive flow injection chemiluminescence detection of DNA hybridization. Chemical Communications 2009, (37), 5567-5569. 94

193. Zhang, Z.; Xie, J.; Yu, J.; Lu, Z.; Liu, Y., A novel colorimetric immunoassay strategy using iron (iii) oxide magnetic nanoparticles as a label for signal generation and amplification. Journal of Materials Chemistry B 2017, 5 (7), 1454-1460. 194. Pacakova, B.; Kubickova, S.; Salas, G.; Mantlikova, A.; Marciello, M.; Morales, M.; Niznansky, D.; Vejpravova, J., The internal structure of magnetic nanoparticles determines the magnetic response. Nanoscale 2017, 9 (16), 5129-5140. 195. Katz, E., Magneto‐switchable Electrodes and Electrochemical Systems. Electroanalysis 2016, 28 (5), 904-919. 196. Katz, E.; Sheeney-Haj-Ichia, L.; Basnar, B.; Felner, I.; Willner, I., Magnetoswitchable controlled hydrophilicity/hydrophobicity of electrode surfaces using alkyl-chain- functionalized magnetic particles: Application for switchable electrochemistry. Langmuir 2004, 20 (22), 9714-9719. 197. Katz, E.; Willner, I., Switching of directions of bioelectrocatalytic currents and photocurrents at electrode surfaces by using hydrophobic magnetic nanoparticles. Angewandte Chemie 2005, 117 (30), 4869-4872. 198. Zhu, X.; Han, K.; Li, G., Magnetic nanoparticles applied in electrochemical detection of controllable DNA hybridization. Analytical Chemistry 2006, 78 (7), 2447-2449. 199. Privman, M.; Tam, T. K.; Pita, M.; Katz, E., Switchable electrode controlled by enzyme logic network system: approaching physiologically regulated bioelectronics. Journal of the American Chemical Society 2008, 131 (3), 1314-1321. 200. Wang, X.; Zhou, J.; Tam, T. K.; Katz, E.; Pita, M., Switchable electrode controlled by Boolean logic gates using enzymes as input signals. Bioelectrochemistry 2009, 77 (1), 69-73. 201. Zablotskaya, A.; Segal, I.; Popelis, Y.; Mishnev, A.; Maiorov, M.; Zablotsky, D.; Blums, E.; Nikolajeva, V.; Eze, D., Iron oxide superparamagnetic nanocarriers bearing amphiphilic N‐heterocyclic choline analogues as potential 40agents. Applied Organometallic Chemistry 2015, 29 (6), 376-383. 202. Thorat, N. D.; Bohara, R. A.; Malgras, V.; Tofail, S. A.; Ahamad, T.; Alshehri, S. M.; Wu, K. C.-W.; Yamauchi, Y., Multimodal superparamagnetic nanoparticles with unusually enhanced specific absorption rate for synergetic cancer therapeutics and magnetic resonance imaging. ACS Applied Materials & Interfaces 2016, 8 (23), 14656- 14664.

203. Zhang, X.; Jiao, K.; Liu, S.; Hu, Y., Readily reusable electrochemical DNA hybridization biosensor based on the interaction of DNA with single-walled carbon nanotubes. Analytical Chemistry 2009, 81 (15), 6006-6012. 204. Wanunu, M.; Dadosh, T.; Ray, V.; Jin, J.; McReynolds, L.; Drndić, M., Rapid electronic detection of probe-specific microRNAs using thin nanopore sensors. Nature Nanotechnology 2010, 5 (11), 807-814.

205. Cao, Y.; Wen, L.; Svec, F.; Tan, T.; Lv, Y., Magnetic AuNP@Fe3O4 nanoparticles as reusable carriers for reversible enzyme immobilization. Chemical Engineering Journal 2016, 286, 272-281. 206. Zakaria, M. B.; Belik, A. A.; Liu, C. H.; Hsieh, H. Y.; Liao, Y. T.; Malgras, V.; Yamauchi, Y.; Wu, K. C. W., Prussian Blue Derived Nanoporous Iron Oxides as Anticancer Drug Carriers for Magnetic‐Guided Chemotherapy. Chemistry–An Asian Journal 2015, 10 (7), 1457-1462. 207. Yadav, S.; Masud, M. K.; Islam, M. N.; Gopalan, V.; Lam, A. K.-y.; Tanaka, S.; Nguyen, N.-T.; Hossain, M. S. A.; Li, C.; Yamauchi, M. Y.; Shiddiky, M. J. A., Gold-loaded nanoporous iron oxide nanocubes: a novel dispersible capture agent for tumor- associated autoantibody analysis in serum. Nanoscale 2017, 9 (25), 8805-8814. 208. Gao, L.; Zhuang, J.; Nie, L.; Zhang, J.; Zhang, Y.; Gu, N.; Wang, T.; Feng, J.; Yang, D.; Perrett, S., Intrinsic peroxidase-like activity of ferromagnetic nanoparticles. Nature Nanotechnology 2007, 2 (9), 577-583. 209. Wang, X.; Hu, Y.; Wei, H., Nanozymes in bionanotechnology: from sensing to therapeutics and beyond. Inorganic Chemistry Frontiers 2016, 3 (1), 41-60. 210. Luo, W.; Zhu, C.; Su, S.; Li, D.; He, Y.; Huang, Q.; Fan, C., Self-catalyzed, self-limiting growth of glucose oxidase-mimicking gold nanoparticles. ACS Nano 2010, 4 (12), 7451- 7458. 211. Gao, N.; Dong, K.; Zhao, A.; Sun, H.; Wang, Y.; Ren, J.; Qu, X., Polyoxometalate-based nanozyme: Design of a multifunctional enzyme for multi-faceted treatment of Alzheimer’s disease. Nano Research 2016, 9 (4), 1079-1090. 212. Shen, X.; Zhu, L.; Liu, G.; Tang, H.; Liu, S.; Li, W., Photocatalytic removal of pentachlorophenol by means of an enzyme-like molecular imprinted photocatalyst and inhibition of the generation of highly toxic intermediates. New Journal of Chemistry 2009, 33 (11), 2278-2285. 213. Breslow, R., Biomimetic chemistry: biology as an inspiration. Journal of Biological Chemistry 2009, 284 (3), 1337-1342. 96

214. Wei, H.; Wang, E., Nanomaterials with enzyme-like characteristics (nanozymes): next- generation artificial enzymes. Chemical Society Reviews 2013, 42 (14), 6060-6093. 215. Shin, H. Y.; Park, T. J.; Kim, M. I., Recent research trends and future prospects in nanozymes. Journal of Nanomaterials 2015, 2015. 216. Chen, Z.; Yin, J.-J.; Zhou, Y.-T.; Zhang, Y.; Song, L.; Song, M.; Hu, S.; Gu, N., Dual enzyme-like activities of iron oxide nanoparticles and their implication for diminishing cytotoxicity. ACS Nano 2012, 6 (5), 4001-4012. 217. Dalui, A.; Pradhan, B.; Thupakula, U.; Khan, A. H.; Kumar, G. S.; Ghosh, T.; Satpati, B.; Acharya, S., Insight into the mechanism revealing the peroxidase mimetic catalytic activity of quaternary CuZnFeS nanocrystals: colorimetric biosensing of hydrogen peroxide and glucose. Nanoscale 2015, 7 (19), 9062-9074. 218. Tanaka, S.; Masud, M. K.; Kaneti, Y. V.; Shiddiky, M. J.; Fatehmulla, A.; Aldhafiri, A. M.; Farooq, W. A.; Bando, Y.; Hossain, M. S. A.; Yamauchi, Y., Enhanced Peroxidase Mimetic Activity of Porous Iron Oxide Nanoflakes. ChemNanoMat 2019, 5 (4), 506-513.

219. Wei, H.; Wang, E., Fe3O4 magnetic nanoparticles as peroxidase mimetics and their

applications in H2O2 and glucose detection. Analytical Chemistry 2008, 80 (6), 2250- 2254. 220. Zuo, X.; Peng, C.; Huang, Q.; Song, S.; Wang, L.; Li, D.; Fan, C., Design of a carbon nanotube/magnetic nanoparticle-based peroxidase-like nanocomplex and its application for highly efficient catalytic oxidation of phenols. Nano Research 2009, 2 (8), 617-623. 221. Liu, Y.; Yu, F., Substrate-specific modifications on magnetic iron oxide nanoparticles as an artificial peroxidase for improving sensitivity in glucose detection. Nanotechnology 2011, 22 (14), 145704. 222. Dutta, A. K.; Maji, S. K.; Srivastava, D. N.; Mondal, A.; Biswas, P.; Paul, P.; Adhikary, B., Peroxidase-like activity and amperometric sensing of hydrogen peroxide by Fe 2 O

3 and Prussian Blue-modified Fe2O3 nanoparticles. Journal of Molecular Catalysis A: Chemical 2012, 360, 71-77. 223. Xing, Z.; Tian, J.; Asiri, A. M.; Qusti, A. H.; Al-Youbi, A. O.; Sun, X., Two-dimensional hybrid mesoporous Fe 2 O 3–graphene nanostructures: a highly active and reusable peroxidase mimetic toward rapid, highly sensitive optical detection of glucose. Biosensors and Bioelectronics 2014, 52, 452-457.

224. Shi, W.; Zhang, X.; He, S.; Huang, Y., CoFe2O4 magnetic nanoparticles as a peroxidase mimic mediated chemiluminescence for hydrogen peroxide and glucose. Chemical Communications 2011, 47 (38), 10785-10787. 97

225. Ghosh, A. B.; Saha, N.; Sarkar, A.; Dutta, A. K.; Biswas, P.; Nag, K.; Adhikary, B.,

Morphological tuning of Eu2O2S nanoparticles, manifestation of peroxidase-like activity and glucose assay use. New Journal of Chemistry 2016, 40 (2), 1595-1604. 226. Wang, G.-L.; Xu, X.-F.; Qiu, L.; Dong, Y.-M.; Li, Z.-J.; Zhang, C., Dual responsive enzyme mimicking activity of AgX (X= Cl, Br, I) nanoparticles and its application for cancer cell detection. ACS Applied Materials & Interfaces 2014, 6 (9), 6434-6442. 227. Kim, M.; Kim, M. S.; Kweon, S. H.; Jeong, S.; Kang, M. H.; Kim, M. I.; Lee, J.; Doh, J., Simple and Sensitive Point‐of‐Care Bioassay System Based on Hierarchically Structured Enzyme‐Mimetic Nanoparticles. Advanced Healthcare Materials 2015, 4 (9), 1311-1316. 228. Schirhagl, R., Bioapplications for molecularly imprinted polymers. Analytical Chemistry 2014, 86 (1), 250-261. 229. Zhang, Z.; Zhang, X.; Liu, B.; Liu, J., Molecular imprinting on inorganic nanozymes for hundred-fold enzyme specificity. Journal of the American Chemical Society 2017, 139 (15), 5412-5419. 230. Abu-Salah, K. M.; Zourob, M. M.; Mouffouk, F.; Alrokayan, S. A.; Alaamery, M. A.; Ansari, A. A., DNA-based nanobiosensors as an emerging platform for detection of disease. Sensors 2015, 15 (6), 14539-14568. 231. Martín-Fernández, B.; Miranda-Ordieres, A. J.; Lobo-Castañón, M. J.; Frutos- Cabanillas, G.; de-los-Santos-Álvarez, N.; López-Ruiz, B., Strongly structured DNA sequences as targets for genosensing: Sensing phase design and coupling to PCR amplification for a highly specific 33-mer gliadin DNA fragment. Biosensors and Bioelectronics 2014, 60, 244-251. 232. Peng, Y.; Jiang, J.; Yu, R., A sensitive electrochemical biosensor for microRNA detection based on streptavidin–gold nanoparticles and enzymatic amplification. Analytical Methods 2014, 6 (9), 2889-2893. 233. Dong, X.-Y.; Mi, X.-N.; Wang, B.; Xu, J.-J.; Chen, H.-Y., Signal amplification for DNA

detection based on the HRP-functionalized Fe3O4 nanoparticles. Talanta 2011, 84 (2), 531-537. 234. Loaiza, Ó. A.; Jubete, E.; Ochoteco, E.; Cabañero, G.; Grande, H.; Rodríguez, J., Gold coated ferric oxide nanoparticles based disposable magnetic genosensors for the detection of DNA hybridization processes. Biosensors and Bioelectronics 2011, 26 (5), 2194-2200.

235. Wang, J.; Palecek, E.; Nielsen, P. E.; Rivas, G.; Cai, X.; Shiraishi, H.; Dontha, N.; Luo, D.; Farias, P. A., Peptide nucleic acid probes for sequence-specific DNA biosensors. Journal of the American Chemical Society 1996, 118 (33), 7667-7670. 236. Hunt, E. A.; Broyles, D.; Head, T.; Deo, S. K., MicroRNA detection: current technology and research strategies. Annual Review of Analytical Chemistry 2015, 8, 217-237. 237. Aldalbahi, A.; Rahaman, M.; Almoiqli, M.; Hamedelniel, A.; Alrehaili, A., Single- Walled Carbon Nanotube (SWCNT) Loaded Porous Reticulated Vitreous Carbon (RVC) Electrodes Used in a Capacitive Deionization (CDI) Cell for Effective Desalination. Nanomaterials (Basel) 2018, 8 (7), 527. 238. Islam, M. N.; Masud, M. K.; Nguyen, N.-T.; Gopalan, V.; Alamri, H. R.; Alothman, Z. A.; Al Hossain, M. S.; Yamauchi, Y.; Lam, A. K.-Y.; Shiddiky, M. J., Gold-loaded nanoporous ferric oxide nanocubes for electrocatalytic detection of microRNA at attomolar level. Biosensors and Bioelectronics 2017. 239. Kokkinos, C.; Economou, A.; Prodromidis, M. I., Electrochemical immunosensors: Critical survey of different architectures and transduction strategies. TrAC Trends in Analytical Chemistry 2016, 79, 88-105. 240. Moreno-Guzmán, M.; González-Cortés, A.; Yáñez-Sedeño, P.; Pingarrón, J. M., A disposable electrochemical immunosensor for prolactin involving affinity reaction on streptavidin-functionalized magnetic particles. Analytica Chimica Acta 2011, 692 (1), 125-130. 241. Li, H.; Wei, Q.; He, J.; Li, T.; Zhao, Y.; Cai, Y.; Du, B.; Qian, Z.; Yang, M., Electrochemical immunosensors for cancer biomarker with signal amplification based on ferrocene functionalized iron oxide nanoparticles. Biosensors and Bioelectronics 2011, 26 (8), 3590-3595. 242. Li, F.; Li, Y.; Dong, Y.; Jiang, L.; Wang, P.; Liu, Q.; Liu, H.; Wei, Q., An ultrasensitive label-free electrochemical immunosensor based on signal amplification strategy of multifunctional magnetic graphene loaded with cadmium ions. Scientific Reports 2016, 6, 21281. 243. Zheng, T.; Fu, J.-J.; Hu, L.; Qiu, F.; Hu, M.; Zhu, J.-J.; Hua, Z.-C.; Wang, H., Nanoarchitectured electrochemical cytosensors for selective detection of leukemia cells and quantitative evaluation of death receptor expression on cell surfaces. Analytical Chemistry 2013, 85 (11), 5609-5616. 244. Zheng, T.; Zhang, Q.; Feng, S.; Zhu, J.-J.; Wang, Q.; Wang, H., Robust nonenzymatic hybrid nanoelectrocatalysts for signal amplification toward ultrasensitive 99

electrochemical cytosensing. Journal of the American Chemical Society 2014, 136 (6), 2288-2291. 245. Khoshfetrat, S. M.; Mehrgardi, M. A., Amplified detection of leukemia cancer cells using an aptamer-conjugated gold-coated magnetic nanoparticles on a nitrogen-doped graphene modified electrode. Bioelectrochemistry 2017, 114, 24-32. 246. Sun, D.; Lu, J.; Zhong, Y.; Yu, Y.; Wang, Y.; Zhang, B.; Chen, Z., Sensitive electrochemical aptamer cytosensor for highly specific detection of cancer cells based on the hybrid nanoelectrocatalysts and enzyme for signal amplification. Biosensors and Bioelectronics 2016, 75, 301-307. 247. Tian, L.; Qi, J.; Qian, K.; Oderinde, O.; Cai, Y.; Yao, C.; Song, W.; Wang, Y., An ultrasensitive electrochemical cytosensor based on the magnetic field assisted

binanozymes synergistic catalysis of Fe3O4 nanozyme and reduced graphene oxide/molybdenum disulfide nanozyme. Sensors and Actuators B: Chemical 2018. 248. Maier, S. A.; Brongersma, M. L.; Kik, P. G.; Meltzer, S.; Requicha, A. A.; Atwater, H. A., Plasmonics—a route to nanoscale optical devices. Advanced Materials 2001, 13 (19), 1501-1505. 249. Moro, L.; Turemis, M.; Marini, B.; Ippodrino, R.; Giardi, M. T., Better together: Strategies based on magnetic particles and quantum dots for improved biosensing. Biotechnology Advances 2017, 35 (1), 51-63. 250. VS, A. P.; Joseph, P.; SCG, K. D.; Lakshmanan, S.; Kinoshita, T.; Muthusamy, S., Colorimetric sensors for rapid detection of various analytes. Materials Science and Engineering: C 2017, 78, 1231-1245. 251. Sung, Y. J.; Suk, H.-J.; Sung, H. Y.; Li, T.; Poo, H.; Kim, M.-G., Novel antibody/gold nanoparticle/magnetic nanoparticle nanocomposites for immunomagnetic separation and rapid colorimetric detection of Staphylococcus aureus in milk. Biosensors and Bioelectronics 2013, 43, 432-439. 252. Suaifan, G. A.; Esseghaier, C.; Ng, A.; Zourob, M., Ultra-rapid colorimetric assay for protease detection using magnetic nanoparticle-based biosensors. Analyst 2013, 138 (13), 3735-3739. 253. Chinnappan, R.; Al Attas, S.; Kaman, W. E.; Bikker, F. J.; Zourob, M., Development of magnetic nanoparticle based calorimetric assay for the detection of bovine mastitis in cow milk. Analytical Biochemistry 2017, 523, 58-64.

100

254. Suaifan, G. A.; Alhogail, S.; Zourob, M., Paper-based magnetic nanoparticle-peptide probe for rapid and quantitative colorimetric detection of Escherichia coli O157: H7. Biosensors and Bioelectronics 2017, 92, 702-708. 255. Masson, J.-F., Surface plasmon resonance clinical biosensors for medical diagnostics. ACS Sensors 2017, 2 (1), 16-30. 256. Mauriz, E.; García-Fernández, M.; Lechuga, L. M., Towards the design of universal immunosurfaces for SPR-based assays: A review. TrAC, Trends Analytical Chemistry2016, 79, 191-198. 257. Wu, B.; Jiang, R.; Wang, Q.; Huang, J.; Yang, X.; Wang, K.; Li, W.; Chen, N.; Li, Q., Detection of C-reactive protein using nanoparticle-enhanced surface plasmon resonance using an aptamer-antibody sandwich assay. Chemical Communication 2016, 52 (17), 3568-3571. 258. Beccati, D.; Halkes, K. M.; Batema, G. D.; Guillena, G.; Carvalho de Souza, A.; van Koten, G.; Kamerling, J. P., SPR Studies of Carbohydrate–Protein Interactions: Signal Enhancement of Low‐Molecular‐Mass Analytes by Organoplatinum (II)‐Labeling. ChemBioChem 2005, 6 (7), 1196-1203. 259. Lin, K.; Lu, Y.; Chen, J.; Zheng, R.; Wang, P.; Ming, H., Surface plasmon resonance hydrogen sensor based on metallic grating with high sensitivity. Optics Express 2008, 16 (23), 18599-18604. 260. Tabassum, R.; Gupta, B. D. In Analysis of electric field distribution in SPR refractive index sensor using different conducting metal oxides, Laser Science, Optical Society of America, 2015, JW2A. 13. 261. He, L.; Smith, E. A.; Natan, M. J.; Keating, C. D., The distance-dependence of colloidal Au-amplified surface plasmon resonance. The Journal of Physical Chemistry B 2004, 108 (30), 10973-10980.

262. Wang, J.; Zhu, Z.; Munir, A.; Zhou, H. S., Fe3O4 nanoparticles-enhanced SPR sensing for ultrasensitive sandwich bio-assay. Talanta 2011, 84 (3), 783-788. 263. Nguyen, H.; Park, J.; Kang, S.; Kim, M., Surface plasmon resonance: a versatile technique for biosensor applications. Sensors 2015, 15 (5), 10481-10510.

264. Guo, X., Fe3O4@ Au nanoparticles enhanced surface plasmon resonance for ultrasensitive immunoassay. Sensors and Actuators B: Chemical 2014, 205, 276-280. 265. Cai, L.; Zhan, R.; Pu, K.-Y.; Qi, X.; Zhang, H.; Huang, W.; Liu, B., Butterfly-shaped conjugated oligoelectrolyte/graphene oxide integrated assay for light-up visual detection of heparin. Analytical Chemistry 2011, 83 (20), 7849-7855. 101

266. Wu, Q.; Sun, Y.; Zhang, D.; Li, S.; Wang, X.; Song, D., Magnetic field-assisted SPR biosensor based on carboxyl-functionalized graphene oxide sensing film and Fe3O4- hollow gold nanohybrids probe. Biosensors and Bioelectronics 2016, 86, 95-101. 267. Li, S.; Wu, Q.; Ma, P.; Zhang, Y.; Song, D.; Wang, X.; Sun, Y., A sensitive SPR biosensor based on hollow gold nanospheres and improved sandwich assay with PDA-Ag@

Fe3O4/rGO. Talanta 2018, 180, 156-161. 268. Unser, S.; Bruzas, I.; He, J.; Sagle, L., Localized surface plasmon resonance biosensing: current challenges and approaches. Sensors 2015, 15 (7), 15684-15716. 269. Storhoff, J. J.; Elghanian, R.; Mirkin, C. A.; Letsinger, R. L., Sequence-dependent stability of DNA-modified gold nanoparticles. Langmuir 2002, 18 (17), 6666-6670. 270. Premaratne, G.; Dharmaratne, A. C.; Al Mubarak, Z. H.; Mohammadparast, F.; Andiappan, M.; Krishnan, S., Multiplexed surface plasmon imaging of serum

biomolecules: Fe3O4@ Au Core/shell nanoparticles with plasmonic simulation insights. Sensors and Actuators B: Chemical 2019, 299, 126956. 271. Suthanthiraraj, P. P. A.; Sen, A. K., Localized surface plasmon resonance (LSPR) biosensor based on thermally annealed silver nanostructures with on-chip blood-plasma separation for the detection of dengue non-structural protein NS1 antigen. Biosensors and Bioelectronics 2019, 132, 38-46. 272. Jain, P. K.; Xiao, Y.; Walsworth, R.; Cohen, A. E., Surface plasmon resonance enhanced magneto-optics (SuPREMO): Faraday rotation enhancement in gold-coated iron oxide nanocrystals. Nano Letters 2009, 9 (4), 1644-1650. 273. Cai, Y.; Zhu, J.; He, J.; Yang, W.; Ma, C.; Xiong, F.; Li, F.; Chen, W.; Chen, P., Magnet Patterned Superparamagnetic Fe3O4/Au Core–Shell Nanoplasmonic Sensing Array for Label‐Free High Throughput Cytokine Immunoassay. Advanced Healthcare Materials 2019, 8 (4), 1801478. 274. Pavel, I.; McCarney, E.; Elkhaled, A.; Morrill, A.; Plaxco, K.; Moskovits, M., Label-free SERS detection of small proteins modified to act as bifunctional linkers. The Journal of Physical Chemistry C 2008, 112 (13), 4880-4883. 275. Ngo, H. T.; Wang, H.-N.; Fales, A. M.; Vo-Dinh, T., Plasmonic SERS biosensing nanochips for DNA detection. Analytical and Bioanalytical Chemistry 2016, 408 (7), 1773-1781. 276. Wang, H.-N.; Crawford, B. M.; Fales, A. M.; Bowie, M. L.; Seewaldt, V. L.; Vo-Dinh, T., Multiplexed detection of microRNA biomarkers using SERS-based inverse molecular

102

sentinel (iMS) nanoprobes. The Journal of Physical Chemistry C 2016, 120 (37), 21047- 21055. 277. Li, L.; Liao, M.; Chen, Y.; Shan, B.; Li, M., Surface-enhanced Raman spectroscopy (SERS) nanoprobes for ratiometric detection of cancer cells. Journal of Materials Chemistry B 2019, 7 (5), 815-822. 278. Laing, S.; Gracie, K.; Faulds, K., Multiplex in vitro detection using SERS. Chemical Society Reviews 2016, 45 (7), 1901-1918.

279. Pang, Y.; Wang, C.; Wang, J.; Sun, Z.; Xiao, R.; Wang, S., Fe3O4@Ag magnetic nanoparticles for microRNA capture and duplex-specific nuclease signal amplification based SERS detection in cancer cells. Biosensors and Bioelectronics 2016, 79, 574-580. 280. Zhang, H.; Yi, Y.; Zhou, C.; Ying, G.; Zhou, X.; Fu, C.; Zhu, Y.; Shen, Y., SERS detection of microRNA biomarkers for cancer diagnosis using gold-coated paramagnetic nanoparticles to capture SERS-active gold nanoparticles. RSC Advances 2017, 7 (83), 52782-52793. 281. Sun, Y.; Xu, L.; Zhang, F.; Song, Z.; Hu, Y.; Ji, Y.; Shen, J.; Li, B.; Lu, H.; Yang, H., A promising magnetic SERS immunosensor for sensitive detection of avian influenza virus. Biosensors and Bioelectronics . 2017, 89, 906-912.

282. Qiu, Y.; Deng, D.; Deng, Q.; Wu, P.; Zhang, H.; Cai, C., Synthesis of magnetic Fe3O4– Au hybrids for sensitive SERS detection of cancer cells at low abundance. Journal of Materials Chemistry B 2015, 3 (22), 4487-4495. 283. Ferrari, A. C.; Basko, D. M., Raman spectroscopy as a versatile tool for studying the properties of graphene. Nature Nanotechnology 2013, 8 (4), 235. 284. Chen, M.; Luo, W.; Zhang, Z.; Zhu, F.; Liao, S.; Yang, H.; Chen, X., Sensitive surface enhanced Raman spectroscopy (SERS) detection of methotrexate by core-shell-satellite magnetic microspheres. Talanta 2017, 171, 152-158. 285. Demeritte, T.; Viraka Nellore, B. P.; Kanchanapally, R.; Sinha, S. S.; Pramanik, A.; Chavva, S. R.; Ray, P. C., Hybrid graphene oxide based plasmonic-magnetic multifunctional nanoplatform for selective separation and label-free identification of Alzheimer’s disease biomarkers. ACS Applied Materials & Interfaces 2015, 7 (24), 13693-13700. 286. Zheng, W.; Jiang, X., Integration of nanomaterials for colorimetric immunoassays with improved performance: a functional perspective. Analyst 2016, 141 (4), 1196-1208.

103

287. Kaittanis, C.; Santra, S.; Perez, J. M., Role of nanoparticle valency in the nondestructive magnetic-relaxation-mediated detection and magnetic isolation of cells in complex media. Journal of the American Chemical Society 2009, 131 (35), 12780-12791. 288. Yang, Y.-C.; Wang, Y.-T.; Tseng, W.-L., Amplified peroxidase-like activity in iron oxide nanoparticles using adenosine monophosphate: Application to urinary protein sensing. ACS Applied Materials & Interfaces 2017, 9 (11), 10069-10077. 289. Masud, M. K.; Yadav, S.; Islam, M. N.; Nguyen, N.-T.; Salomon, C.; Kline, R.; Alamri, H. R.; Alothman, Z. A.; Yamauchi, Y.; Hossain, M. S. A., Gold-Loaded Nanoporous Ferric Oxide Nanocubes with Peroxidase-Mimicking Activity for Electrocatalytic and Colorimetric Detection of Autoantibody. Analytical Chemistry 2017, 89 (20), 11005- 11013. 290. Park, K. S.; Kim, M. I.; Cho, D. Y.; Park, H. G., Label‐Free Colorimetric Detection of Nucleic Acids Based on Target‐Induced Shielding Against the Peroxidase‐Mimicking Activity of Magnetic Nanoparticles. Small 2011, 7 (11), 1521-1525. 291. Li, Y.; Srinivasan, B.; Jing, Y.; Yao, X.; Hugger, M. A.; Wang, J.-P.; Xing, C., Nanomagnetic competition assay for low-abundance protein biomarker quantification in unprocessed human sera. Journal of the American Chemical Society 2010, 132 (12), 4388-4392. 292. Choi, J.; Gani, A. W.; Bechstein, D. J.; Lee, J.-R.; Utz, P. J.; Wang, S. X., Portable, one- step, and rapid GMR biosensor platform with smartphone interface. Biosensors and Bioelectronics 2016, 85, 1-7. 293. Krishna, V. D.; Wu, K.; Perez, A. M.; Wang, J.-P., Giant magnetoresistance-based biosensor for detection of influenza A virus. Frontiers in Microbiology 2016, 7, 400. 294. Zhou, Y.; Xie, Q., Hyaluronic acid-coated magnetic nanoparticles-based selective collection and detection of leukemia cells with quartz crystal microbalance. Sensors and Actuators B: Chemical 2016, 223, 9-14. 295. Sun, W.; Song, W.; Guo, X.; Wang, Z., Ultrasensitive detection of nucleic acids and proteins using quartz crystal microbalance and surface plasmon resonance sensors based on target-triggering multiple signal amplification strategy. Analytica Chimica Acta 2017, 978, 42-47. 296. Zhan, S.; Lou, X.; Zhou, P.; Xia, F., Sandwich Assays Based on SPR, SERS, GMR, QCM, Microcantilever, SAW, and RRS Techniques for Protein Detection. In Biosensors Based on Sandwich Assays, Springer, 2018, 69-91.

104

297. Körner, J.; Reiche, C. F.; Ghunaim, R.; Fuge, R.; Hampel, S.; Büchner, B.; Mühl, T.,

Magnetic properties of individual Co2FeGa Heusler nanoparticles studied at room temperature by a highly sensitive co-resonant cantilever sensor. Scientific Reports 2017, 7 (1), 8881. 298. Yue, Q.; Tao, L.; Hou, Y.; Zhang, C.; Wang, Y.; Hong, M.; Li, C.-Z., Assay of miRNA in cell samples using enhanced resonance light scattering technique based on self aggregation of magnetic nanoparticles. Nanomedicine 2018, 13 (18), 2301-2310. 299. Yuan, Y.-H.; Wu, Y.-D.; Chi, B.-Z.; Wen, S.-H.; Liang, R.-P.; Qiu, J.-D., Simultaneously electrochemical detection of microRNAs based on multifunctional magnetic nanoparticles probe coupling with hybridization chain reaction. Biosensors and Bioelectronics 2017, 97, 325-331. 300. Yu, N.; Wang, Z.; Wang, C.; Han, J.; Bu, H., Combining padlock exponential rolling circle amplification with CoFe2O4 magnetic nanoparticles for microRNA detection by nanoelectrocatalysis without a substrate. Analytica Chimica Acta 2017, 962, 24-31. 301. Islam, M. N.; Gorgannezhad, L.; Masud, M. K.; Tanaka, S.; Hossain, M. S. A.; Yamauchi, Y.; Nguyen, N.-T.; Shiddiky, M. J. A., Graphene-Oxide-Loaded Superparamagnetic Iron Oxide Nanoparticles for Ultrasensitive Electrocatalytic Detection of MicroRNA. ChemElectroChem 2018, 5 (17), 2488-2495. 302. Tavallaie, R.; McCarroll, J.; Le Grand, M.; Ariotti, N.; Schuhmann, W.; Bakker, E.; Tilley, R. D.; Hibbert, D. B.; Kavallaris, M.; Gooding, J. J., Nucleic acid hybridization on an electrically reconfigurable network of gold-coated magnetic nanoparticles enables microRNA detection in blood. Nature Nanotechnology 2018, 13 (11), 1066. 303. Li, Y.; Yu, C.; Yang, B.; Liu, Z.; Xia, P.; Wang, Q., Target-catalyzed hairpin assembly and metal-organic frameworks mediated nonenzymatic co-reaction for multiple signal amplification detection of miR-122 in human serum. Biosensors and Bioelectronics 2018, 102, 307-315. 304. Gorgannezhad, L.; Umer, M.; Kamal Masud, M.; Hossain, M. S. A.; Tanaka, S.; Yamauchi, Y.; Salomon, C.; Kline, R.; Nguyen, N.-T.; Shiddiky, M. J. A., Detection of FGFR2 : FAM76A Fusion Gene in Circulating Tumor RNA Based on Catalytic Signal Amplification of Graphene Oxide-loaded Magnetic Nanoparticles. Electroanalysis 2018, 30 (10), 2293-2301. 305. Zhang, T.; Chai, H.; Meng, F.; Guo, Z.; Jiang, Y.; Miao, P., DNA-Functionalized Porous Fe3O4 Nanoparticles for the Construction of Self-Powered miRNA Biosensor with

105

Target Recycling Amplification. ACS Applied Materials & Interfaces 2018, 10 (43), 36796-36804. 306. Tang, P.; Zhang, H.; Huo, J.; Lin, X., An electrochemical sensor based on iron(ii,iii)@graphene oxide@molecularly imprinted polymer nanoparticles for interleukin-8 detection in saliva. Analytical Methods 2015, 7 (18), 7784-7791. 307. Hwang, H.; Choi, E.; Han, S.; Lee, Y.; Choi, T.; Kim, M.; Shin, H.; Kim, J.; Choi, J., MESIA: Magnetic force-assisted electrochemical sandwich immunoassays for quantification of prostate-specific antigen in human serum. Analytica Chimica Acta 2019, 1061, 92-100. 308. Fang, S.; Dong, X.; Liu, S.; Penng, D.; He, L.; Wang, M.; Fu, G.; Feng, X.; Zhang, Z., A

label-free multi-functionalized electrochemical aptasensor based on a Fe3O4@3D- rGO@plasma-polymerized (4-vinyl pyridine) nanocomposite for the sensitive detection of proteins in whole blood. Electrochimica Acta 2016, 212, 1-9. 309. Yadav, S.; Masud, M. K.; Islam, M. N.; Gopalan, V.; Lam, A. K.; Tanaka, S.; Nguyen, N. T.; Hossain, M. S. A.; Li, C.; Yamauchi, M. Y.; Shiddiky, M. J. A., Gold-loaded nanoporous iron oxide nanocubes: a novel dispersible capture agent for tumor- associated autoantibody analysis in serum. Nanoscale 2017, 9 (25), 8805-8814. 310. Boriachek, K.; Masud, M. K.; Palma, C.; Phan, H.-P.; Yamauchi, Y.; Hossain, M. S. A.; Nguyen, N.-T.; Salomon, C.; Shiddiky, M. J. A., Avoiding Pre-Isolation Step in Exosome Analysis: Direct Isolation and Sensitive Detection of Exosomes Using Gold-Loaded Nanoporous Ferric Oxide Nanozymes. Analytical Chemistry 2019, 91 (6), 3827-3834. 311. Daneshpour, M.; Izadi, P.; Omidfar, K., Femtomolar level detection of RASSF1A tumor suppressor gene methylation by electrochemical nano-genosensor based on Fe3O4/TMC/Au nanocomposite and PT-modified electrode. Biosensors and Bioelectronics . 2016, 77, 1095-1103. 312. Omidfar, K.; Darzianiazizi, M.; Ahmadi, A.; Daneshpour, M.; Shirazi, H., A high

sensitive electrochemical nanoimmunosensor based on Fe3O4/TMC/Au nanocomposite and PT-modified electrode for the detection of cancer biomarker epidermal growth factor receptor. Sensors and Actuators B: Chemical 2015, 220, 1311-1319. 313. Lee, J.; Morita, M.; Takemura, K.; Park, E. Y., A multi-functional gold/iron-oxide nanoparticle-CNT hybrid nanomaterial as virus DNA sensing platform. Biosensors and Bioelectronics . 2018, 102, 425-431. 314. Wu, S.; Duan, N.; Qiu, Y.; Li, J.; Wang, Z., Colorimetric aptasensor for the detection of

Salmonella enterica serovar typhimurium using ZnFe2O4-reduced graphene oxide 106

nanostructures as an effective peroxidase mimetics. International Journal of Food Microbiology 2017, 261, 42-48. 315. Amini, B.; Kamali, M.; Salouti, M.; Yaghmaei, P., Fluorescence bio-barcode DNA assay based on gold and magnetic nanoparticles for detection of Exotoxin A gene sequence. Biosensors and Bioelectronics . 2017, 92, 679-686. 316. Hou, Y.-H.; Wang, J.-J.; Jiang, Y.-Z.; Lv, C.; Xia, L.; Hong, S.-L.; Lin, M.; Lin, Y.; Zhang, Z.-L.; Pang, D.-W., A colorimetric and electrochemical immunosensor for point- of-care detection of enterovirus 71. Biosensors and Bioelectronics . 2018, 99, 186-192. 317. Jimenez, A. M. J.; Rodrigo, M. A. M.; Milosavljevic, V.; Krizkova, S.; Kopel, P.; Heger, Z.; Adam, V., Gold nanoparticles-modified nanomaghemite and quantum dots-based hybridization assay for detection of HPV. Sensors and Actuators B: Chemical 2017, 240, 503-510. 318. Qian, J.; Xu, D.; Ho, S.-L.; Wang, K.; Wong, M. S.; Li, H.-W., Magnetically controlled immunosensor for highly sensitive detection of carcinoembryonic antigen based on an efficient “turn-on” cyanine fluorophore. Sensors and Actuators B: Chemical 2018, 258, 133-140. 319. Chen, Y.; Xianyu, Y.; Sun, J.; Niu, Y.; Wang, Y.; Jiang, X., One-step detection of pathogens and cancer biomarkers by the naked eye based on aggregation of immunomagnetic beads. Nanoscale 2016, 8 (2), 1100-1107. 320. Fu, G.; Sanjay, S. T.; Li, X., Cost-effective and sensitive colorimetric immunosensing using an iron oxide-to-Prussian blue nanoparticle conversion strategy. Analyst 2016, 141 (12), 3883-3889. 321. Jiang, L.-F.; Chen, B.-C.; Chen, B.; Li, X.-J.; Liao, H.-L.; Huang, H.-M.; Guo, Z.-J.; Zhang, W.-Y.; Wu, L., Detection of Aβ oligomers based on magnetic-field-assisted separation of aptamer-functionalized Fe3O4 magnetic nanoparticles and BaYF5: Yb, Er nanoparticles as upconversion fluorescence labels. Talanta 2017, 170, 350-357. 322. Narmani, A.; Kamali, M.; Amini, B.; Kooshki, H.; Amini, A.; Hasani, L., Highly sensitive and accurate detection of Vibrio cholera O1 OmpW gene by fluorescence DNA biosensor based on gold and magnetic nanoparticles. Process Biochemistry 2018, 65, 46-54.

323. Yang, L.; Li, N.; Wang, K.; Hai, X.; Liu, J.; Dang, F., A novel peptide/Fe3O4@SiO2-Au nanocomposite-based fluorescence biosensor for the highly selective and sensitive detection of prostate-specific antigen. Talanta 2018, 179, 531-537.

107

324. Chen, H.; Hou, Y.; Ye, Z.; Wang, H.; Koh, K.; Shen, Z.; Shu, Y., Label-free surface plasmon resonance cytosensor for breast cancer cell detection based on nano- conjugation of monodisperse magnetic nanoparticle and folic acid. Sensors and Actuators B: Chemical 2014, 201, 433-438. 325. Tang, L.; Casas, J.; Venkataramasubramani, M., Magnetic nanoparticle mediated enhancement of localized surface plasmon resonance for ultrasensitive bioanalytical assay in human blood plasma. Analytical Chemistry 2013, 85 (3), 1431-1439. 326. Mei, Z.; Dhanale, A.; Gangaharan, A.; Sardar, D. K.; Tang, L., Water dispersion of magnetic nanoparticles with selective Biofunctionality for enhanced plasmonic biosensing. Talanta 2016, 151, 23-29. 327. Lou, Z.; Han, H.; Zhou, M.; Wan, J.; Sun, Q.; Zhou, X.; Gu, N., Fabrication of magnetic conjugation clusters via intermolecular assembling for ultrasensitive surface plasmon resonance (SPR) detection in a wide range of concentrations. Analytical Chemistry 2017, 89 (24), 13472-13479. 328. Bedford, E. E.; Boujday, S.; Pradier, C.-M.; Gu, F. X., Spiky gold shells on magnetic particles for DNA biosensors. Talanta 2018, 182, 259-266. 329. Pang, Y.; Wang, C.; Wang, J.; Sun, Z.; Xiao, R.; Wang, S., Fe3O4@Ag magnetic nanoparticles for microRNA capture and duplex-specific nuclease signal amplification based SERS detection in cancer cells. Biosensors and Bioelectronics . 2016, 79, 574-580. 330. Wang, J.; Wu, X.; Wang, C.; Rong, Z.; Ding, H.; Li, H.; Li, S.; Shao, N.; Dong, P.; Xiao, R., Facile synthesis of Au-coated magnetic nanoparticles and their application in bacteria detection via a SERS method. ACS Applied Materials & Interfaces 2016, 8 (31), 19958-19967. 331. Xianyu, Y.; Wang, Q.; Chen, Y., Magnetic particles-enabled biosensors for point-of- care testing. TrAC, Trends Analytical Chemistry 2018, 106, 213-224. 332. Chung, H. J.; Pellegrini, K. L.; Chung, J.; Wanigasuriya, K.; Jayawardene, I.; Lee, K.; Lee, H.; Vaidya, V. S.; Weissleder, R., Nanoparticle detection of urinary markers for point-of-care diagnosis of kidney injury. PloS One 2015, 10 (7), e0133417. 333. Eltzov, E.; Guttel, S.; Low Yuen Kei, A.; Sinawang, P. D.; Ionescu, R. E.; Marks, R. S., Lateral flow immunoassays–from paper strip to smartphone technology. Electroanalysis 2015, 27 (9), 2116-2130. 334. Wang, Y.; Xu, H.; Wei, M.; Gu, H.; Xu, Q.; Zhu, W., Study of superparamagnetic nanoparticles as labels in the quantitative lateral flow immunoassay. Materials Science and Engineering: C 2009, 29 (3), 714-718. 108

335. Kim, M. S.; Kweon, S. H.; Cho, S.; An, S. S. A.; Kim, M. I.; Doh, J.; Lee, J., Pt- Decorated Magnetic Nanozymes for Facile and Sensitive Point-of-Care Bioassay. ACS Applied Materials & Interfaces 2017, 9 (40), 35133-35140. 336. Oliveira-Rodríguez, M.; Serrano-Pertierra, E.; García, A. C.; López-Martín, S.; Yañez- Mo, M.; Cernuda-Morollón, E.; Blanco-López, M. d. C., Point-of-care detection of extracellular vesicles: sensitivity optimization and multiple-target detection. Biosensors and Bioelectronics 2017, 87, 38-45. 337. Lago-Cachón, D.; Rivas, M.; Martínez-García, J. C.; Oliveira-Rodríguez, M.; Blanco- López, M. d. C.; García, J., High frequency lateral flow affinity assay using superparamagnetic nanoparticles. Journal of Magnetism and Magnetic Materials 2017, 423, 436-440. 338. Sebba, D.; Lastovich, A. G.; Kuroda, M.; Fallows, E.; Johnson, J.; Ahouidi, A.; Honko, A. N.; Fu, H.; Nielson, R.; Carruthers, E., A point-of-care diagnostic for differentiating Ebola from endemic febrile diseases. Science Translational Medicine 2018, 10 (471), eaat0944. 339. Petkovic, K.; Metcalfe, G.; Chen, H.; Gao, Y.; Best, M.; Lester, D.; Zhu, Y., Rapid detection of Hendra virus antibodies: an integrated device with nanoparticle assay and chaotic micromixing. Lab on a Chip 2017, 17 (1), 169-177. 340. Wei, X.; Zhou, W.; Sanjay, S. T.; Zhang, J.; Jin, Q.; Xu, F.; Dominguez, D. C.; Li, X., Multiplexed Instrument-Free Bar-Chart SpinChip Integrated with Nanoparticle- Mediated Magnetic Aptasensors for Visual Quantitative Detection of Multiple Pathogens. Analytical Chemistry 2018, 90 (16), 9888-9896. 341. Ren, W.; Mohammed, S. I.; Wereley, S.; Irudayaraj, J., Magnetic Focus Lateral Flow Sensor for Detection of Cervical Cancer Biomarkers. Analytical Chemistry 2019, 91 (4), 2876-2884. 342. Yan, W.; Wang, K.; Xu, H.; Huo, X.; Jin, Q.; Cui, D., Machine Learning Approach to Enhance the Performance of MNP-Labeled Lateral Flow Immunoassay. Nano-Micro Letters 2019, 11 (1), 7. 343. Xuhong, Y.; Sinong, Z.; Jianping, L.; Yu, C.; Juanli, Z.; Chao, Z.; Desheng, L.; Kai, H.; Yali, C.; Wenli, H., A PCR-lateral flow assay system based on gold magnetic nanoparticles for CYP2C19 genotyping and its clinical applications. Artificial cells, Nanomedicine, and Biotechnology 2019, 47 (1), 636-643.

109

344. Sharma, A.; Tok, A. I. Y.; Lee, C.; Ganapathy, R.; Alagappan, P.; Liedberg, B., Magnetic field assisted preconcentration of biomolecules for lateral flow assaying. Sensors and Actuators B: Chemical 2019, 285, 431-437. 345. Huang, Z.; Xiong, Z.; Chen, Y.; Hu, S.; Lai, W., A Sensitive and Matrix Tolerant Lateral Flow Immunoassay Based on Fluorescent Magnetic Nanobeads for the Detection of Clenbuterol in Swine Urine. Journal of Agricultural and Food Chemistry 2019. 346. Yang, D.; Ma, J.; Xue, C.; Wang, L.; Wang, X., One-pot synthesis of poly (acrylic acid)- stabilized Fe3O4 nanocrystal clusters for the simultaneously qualitative and quantitative detection of biomarkers in lateral flow immunoassay. Journal of Pharmaceutical and Biomedical Analysis 2018, 159, 119-126. 347. Hu, J.; Jiang, Y.-Z.; Tang, M.; Wu, L.-L.; Xie, H.-y.; Zhang, Z.-L.; Pang, D.-W., Colorimetric-Fluorescent-Magnetic Nanosphere-Based Multimodal Assay Platform for Salmonella Detection. Analytical Chemistry 2018, 91 (1), 1178-1184. 348. Wang, C.; Guan, D.; Chen, C.; He, S.; Liu, X.; Wang, C.; Wu, H., Rapid detection of unconjugated estriol in the serum via superparamagnetic lateral flow immunochromatographic assay. Analytical and Bioanalytical Chemistry 2018, 410 (1), 123-130. 349. Ren, W.; Cho, I.-H.; Zhou, Z.; Irudayaraj, J., Ultrasensitive detection of microbial cells using magnetic focus enhanced lateral flow sensors. Chemical Communication 2016, 52 (27), 4930-4933. 350. Liu, X.; Zhang, C.; Liu, K.; Wang, H.; Lu, C.; Li, H.; Hua, K.; Zhu, J.; Hui, W.; Cui, Y., Multiple SNPs detection based on lateral flow assay for phenylketonuria diagnostic. Analytical Chemistry 2018, 90 (5), 3430-3436. 351. Kwak, B.; Lee, J.; Lee, D.; Lee, K.; Kwon, O.; Kang, S.; Kim, Y., Selective isolation of magnetic nanoparticle-mediated heterogeneity subpopulation of circulating tumor cells using magnetic gradient based microfluidic system. Biosensors and Bioelectronics, 2017, 88, 153-158. 352. Yuan, K.; Mei, Q.; Guo, X.; Xu, Y.; Yang, D.; Sánchez, B.J.; Sheng, B.; Liu, C.; Hu, Z.; Yu, G.; Ma, H., Antimicrobial peptide based magnetic recognition elements and Au@ Ag-GO SERS tags with stable internal standards: a three in one biosensor for isolation, discrimination and killing of multiple bacteria in whole blood. Chemical Science, 2018, 9 (47), 8781-8795. 353. Ghasemi, R.; Mirahmadi-Zare, S.Z.; Nasr-Esfahani, M.H.; Allafchian, A.; Behmanesh, M., Optical biosensing of Streptococcus agalactiae based on core/shell magnetic 110

nanoparticle-quantum dot. Analytical and Bioanalytical Chemistry, 2019, 411 (25), 6733-6743.

354. Sharafeldin, M.; Bishop, G.W.; Bhakta, S.; El-Sawy, A.; Suib, S.L.; Rusling, J.F., Fe3O4 nanoparticles on graphene oxide sheets for isolation and ultrasensitive amperometric detection of cancer biomarker proteins. Biosensors and Bioelectronics, 2017, 91, 359- 366. 355. Castillo-Torres, K.Y.; Arnold, D.P.; McLamore, E.S., Rapid isolation of Escherichia coli from water samples using magnetic microdiscs. Sensors and Actuators B: Chemical, 2019, 291, 58-66. 356. Chattopadhyay, S.; Sabharwal, P.K.; Jain, S.; Kaur, A.; Singh, H., Functionalized polymeric magnetic nanoparticle assisted SERS immunosensor for the sensitive detection of S. typhimurium. Analytica Chimica Acta, 2019, 1067, 98-106. 357. Pang, Y.; Shi, J.; Yang, X.; Wang, C.; Sun, Z.; Xiao, R., Personalized detection of

circling exosomal PD-L1 based on Fe3O4@TiO2 isolation and SERS immunoassay. Biosensors and Bioelectronics, 2020, 148, 111800.

111

2.2. Nanoarchitecture frameworks for electrochemical miRNA detection** With revolutionary advances in next-generation sequencing, the human transcriptome has been comprehensively interrogated. These discoveries have highlighted the emerging functional and regulatory roles of a large fraction of RNAs suggesting the potential they might hold as stable and minimally invasive disease biomarkers. Although a plethora of molecular biology- and biosensor-based RNA-detection strategies have been developed, most remain non-applicable in clinical applications. Multifunctional nanomaterials coupled with sensitive and robust electrochemical readouts may prove useful in these applications. Herein, we summarise the major contributions of engineered nanomaterial-based electrochemical biosensing strategies for the analysis of microRNAs (miRNAs). With special emphasis on nanostructure-based detection, this review also chronicles the needs and challenges of miRNA detection and provides a future perspective on the presented strategies. Cell transcriptomes comprising various RNA species, such as messenger RNA (mRNA), microRNA (miRNA), ribosomal RNA (rRNA), transfer RNA (tRNA), small interfering RNA (siRNA), and long non-coding RNA (lncRNA) have emerged as prominent non-invasive or minimally invasive biomarkers due to their widespread role in many pathophysiological phenomena and cellular processes, including cell-cycle regulation, apoptosis, and post-transcriptional gene regulation.1-4 A growing body of evidence has suggested that dysregulated RNA molecules play a central role in the initiation and progression of several diseases, including cancers.3, 5-6 Various RNA species are not only up- or downregulated in cancers, but different genomic anomalies such as copy number variations, single nucleotide polymorphisms, mutations etc. are also reflected through them.4, 7 Therefore, monitoring the RNA component can prove to be a powerful diagnostic tool for cancer management. Ever since the first discovery of lin-4 in C. elegans,8 miRNAs have come to be recognized as important regulators of cellular function. It has been suggested that miRNAs modulate more than 30% of mammalian genes and participate in almost all known cellular processes,9 including development, DNA repair, cell cycle regulation, differentiation, apoptosis, as well as malignant cellular transformation and host-pathogen interaction.10 In addition to their well- known regulatory function, recent studies have revealed a novel role of miRNAs, i.e., intercellular and inter-organ communication both via extracellular vesicles (e.g., exosomes) mediated miRNA transfer as well as non-membrane-bound miRNAs.11 miRNAs effectuate their regulatory function through post-transcriptional repression of gene expression. However, the mechanisms that underlie miRNA mediated repression are different in animals and plants.12 112

With the exception of at least one miRNA (mir-196) whereby gene regulation is carried out via RNA-induced silencing complex (RISC) induced mRNA cleavage (explained below).13 miRNAs identified in the animal kingdom in most cases bind to their complementary sites in the 3′ UTR of their target genes and negatively regulate their expression. In contrast, binding sites for plant miRNAs are not restricted to the 3′ UTR only and can be found anywhere across the entire length of target mRNA. Therefore, plant miRNAs regulate gene expression via cleavage of target mRNA through the RNA-induced silencing complex (RISC). RISC is a ribonucleoprotein complex which incorporates a small ssRNA (e.g. miRNA) and cleaves target mRNA in the middle of the small ssRNA–mRNA duplex.9 RNA interference (RNAi) is an evolutionarily conserved gene regulation phenomenon which was identified as an endogenous defence mechanism against viral infection whereby foreign nucleic acids entering the cells are destroyed by RISC mediated cleavage.12 RNAi has been exploited as a possible therapeutic strategy against human diseases including cancer. A recent review of the literature noted that more than 14 RNAi based therapeutic programs are currently at various stages of clinical trials.14 For example, clinical trials are underway for Miravirsin (SPC3649) a locked nucleic acid (LNA) ribonucleotide based anti-miRNA drug candidate. Miravirsin targets mir-122, a liver-specific miRNA known to play an important role in the lifecycle of the hepatitis C virus (HCV).15

Although most types of RNAs have been reported to have potential applications in diagnostics, prognostics, and therapeutics, this current review will focus mostly on miRNA-based cancer diagnostics. Being a critical regulator of post-transcriptional gene expression miRNAs, a large group of small (~22 nucleotides) non-coding RNAs, have garnered a considerable recent interest as diagnostic biomarkers.1, 5, 16 Dysregulation of these highly conserved regulatory RNAs can potentially impact, progression and prognosis of cancers.17 Several studies have demonstrated that in comparison with normal cells from the same tissues, altered miRNA expression is a common feature of all human tumours.18 Over the years it has also been demonstrated in several studies that miRNAs circulating in body fluids can be used as disease and therapy response indicators.19 In 2008, Lawrie C.H. et al. first reported in a seminal study the diagnostic role of circulating miRNAs in B-cell lymphoma.20 The levels of miR-21 and miR-155 were reported to be significantly higher in the serum samples of lymphoma patients compared to that from normal subjects. As a diagnostic and prognostic biomarker circulating miRNAs offer several distinct advantages, such as the provision of early detection, better

113

stability, and the potential for a minimally invasive liquid biopsy-based monitoring of cancer and other diseases.

2.2.1. Challenges and considerations Despite the huge potential for miRNAs in the diagnosis and prognosis of cancer, their current detection approaches are generally confined to classic nucleic acid detection methods such as Northern blotting, microarray, reverse-transcription quantitative polymerase chain reaction (RT-qPCR), and next-generation sequencing, as well as relatively robust biosensor-based miRNA sensing. Although miRNA detection technologies have been advancing, several major inherent and technical challenges remain. In addition, several recent studies also challenged the merits of miRNA as an effective biomarker. The section that follows underlines these major challenges and considerations that need to be addressed to achieve the full functionality of miRNA diagnostics.

Stability and proper storage of miRNAs. The ribonuclease (RNase) associated degradation of RNAs at room temperature is considered one of the bottlenecks in RNA analysis. During incubation, both endogenous and exogenous RNases progressively degrade miRNAs, thereby affecting the accuracy of detection.1 The use of RNase inhibitors is considered a potential solution to this problem.21 However, this apprehension is not applicable for those miRNAs, are either confined inside of microvesicles (e.g., exosomes, apoptotic bodies, or microparticles) or in other cases are found to be associated with lipoproteins or RNA-binding proteins. These miRNAs are generally packed into their structures and protect themselves from RNase degradation. Moreover, they are very stable even under extreme conditions including long- term storage, freeze-thaw cycles, etc. and are thus considered as stable biomarkers for disease diagnosis.11, 22 A study of serum storage condition showed that miRNA in pure serum can be stored at -20oC for at least 2–4 years.23 Nevertheless, pre-processing time may influence stability significantly as in vitro-hemolysis can release miRNAs from blood peripheral blood cells.24 As no standard control is available for normalization of RNA level, it has been suggested that the blood samples should be processed within 2-4 hours of collection in ethylenediaminetetraacetic acid (EDTA) tubes.25 Moreover, RNase free solutions and accessories should be used for miRNA experiments in order to prevent and minimize RNAse mediated degradation.

Sample source selection and optimization of RNA extraction. To ensure the quality of miRNA expression profiling, standardization of sample sources and extraction procedures are

114

important, as a little discrepancy there could substantially affect the results. For example, Wang et al. reported that the miRNA concentration was higher in serum than in the plasma of the same individual.26 The presence of platelet-derived miRNAs obtained from the blood coagulation process presumably could result in this inconsistency. Several studies also showed that there is a significant difference in expression profiling of tissue bound and circulating miRNAs.27 EDTA is a commonly used anticoagulant but long term storage of blood samples in EDTA coated tubes may lead to inaccurate miRNA profiling.28 However, standardization of the holding period may help to overcome this challenge. In addition, careful attention should be given to retaining the small RNA fraction while extracting it from the complex biological sample to avoid a false-positive readout.29

The varying size of RNAs and associated challenges. Amplification-based detection of miRNAs is highly challenging. Primers cannot anneal with the small miRNA sequences because of the similar size of traditional primers and miRNA, and thus they cannot amplify them. To solve this problem, miRNAs are enzymatically polyadenylated using oligo(dT) primers to generate longer primer binding sequences for the reverse transcription step in RT- qPCR [30]30. Primer size matching can also be avoided by generating the reverse transcription priming site with a stem-loop primer.31

Poor discrimination of homologous miRNAs. Isolating a specific miRNA in the background of homologous miRNAs (high sequence similarity) originating from the same miRNA family is difficult to achieve reliably. For example, let-7a is under-expressed in lung cancer, while let- 7c is down-regulated in breast cancer.32 Remarkably, let-7a and let-7c have only a single base mismatch. An assay which is unable to differentiate this single base difference can thus easily compromise the diagnostic accuracy. To overcome this, the recognition probe should be highly specific to the target miRNA and be able to discriminate between sequences with a single base difference. One of the approaches to overcome this problem is to employ thermostable LNA probes, which upon incorporation with DNA probes increase the melting temperature (Tm) of the strands, therefore increases the binding affinity, resulting in the enhancement of both specificity and sensitivity.33 In another strategy, p19 binding protein was utilized to bind with dsRNA (p19 does not bind to mRNA, rRNA, ssRNA, ssDNA or dsDNA) to make sequence- independent bindings and reduce the responses from non-specific sequences.34 The nonspecific adsorption of RNAs onto nanoparticle (NP) surfaces is also responsible for biased 115

signals (please see later discussion), which can be overcome via designing target-specific nanoparticulates or engineering target specific signalling events. Recently, a 2D NP based combination nanosensor array was reported for sets of homologous miRNA analyses using a target-specific bind-and-release model, where the signal was achieved through the release of capture probe quenched fluorophores through the formation of very specific target-probe complexes, avoiding signals from any non-specific adsorption or bindings of biomolecules to the sensor surface or NPs.35

Nonspecific responses in the detection platform. In biosensor-based strategies, the clinical sample may have a complex biological matrix containing large amounts of unrelated biomolecules such as proteins and non-target nucleic acids, which could non-specifically be adsorbed on the sensor surface, resulting in a false readout. The use of a suitable blocking/antifouling agent such as mercaptohexanol, mercapto-ethanol, poly(ethylene glycol), or bovine serum albumin is highly recommended to prevent nonspecific binding. Additionally, RNA molecules are also prone to aggregation via ultra-violet induced cross-linking with other RNA or proteins.36 When nanomaterials are integrated into a sensor surface or are in solution for signal generating, interference might also come from non-specific adsorption (low specificity of nanomaterials for highly similar miRNAs) and/ or from an aggregation of NPs (when the amount of nanomaterials is much higher than analytes).37 Therefore, along with making nanomaterials biocompatible and dispersed in aqueous media, there is a need to utilize a very specific recognition probe, keeping away the sensors from a UV source and optimizing the amount needed to be incorporated into sensors.

Assay inconsistency due to platform-dependent variation. Substantial inter-platform differences (variation between different detection methods as well as between different laboratory setup) during miRNA analysis are gradually becoming a grave concern for clinical applications. Readers may refer to the critical reviews by Mestdagh et al., Witwer et al. and Sing et al. for the vast body of knowledge on this issue.38-40 Several works have reported contradictory specificity and inconsistent reproducibility of miRNA detection platforms. The enzymatic reaction or amplification steps during preparation or signal enhancement, the strength of hybridization, complementary probe design, laboratory practices, and detection techniques could be accountable for such inter-platform miRNA quantification variations.

116

Requirement for a highly sensitive assay to analyze clinically relevant miRNA concentrations. Detecting circulating miRNAs is challenging due to their low abundance in plasma and serum (e.g., femtomolar to picomolar levels/ few molecules per cell). Therefore, the development of a sensitive miRNA assay platform is crucial. To enable this, various enrichment and amplification steps are generally employed in conventional miRNA assays, whereas different novel transduction schemes and nanomaterial-based signal enhancement strategies can be incorporated into the sensing strategies.41

2.2.2. Nanoarchitectures for miRNA detection With recent advances, nanostructured materials are transitioning to applications in the fields of biosensing, medicine, and biotechnology. These materials are used in developing novel methods and devices for diagnosis and monitoring of specific diseases via the detection of levels of disease-specific biomolecules. Nanostructured materials generally possess flexible and modular structures, small size, high surface area, low toxicity, enzyme mimicking, and superparamagnetic behaviour, and biocompatibility.42-43 In the past decades, they have widely been used to develop many devices and methods with unprecedented power to comprehensively interrogate genetic, epigenetic, transcriptomic, proteomic, and cellular information from many diseases including cancer.44 In comparison with bulk materials of the same mass, nanostructured materials exhibit expressive advantages in molecular diagnostics, particularly in disease diagnosis applications: (i) the smaller size of nanostructured materials break down the barrier for structural miniaturization of diagnostic platforms, leading to the prospect of designing small-size, inexpensive, portable, low-cost tools in point-of-care diagnostics; (ii) nanostructured materials enable direct contact with the sensing environments (electrolyte, labeller, etc.), thereby accelerating signal transduction, and consequently boosting the robustness and sensitivity of the analysis and lowering the detection limit; (iii) they have the potential to provide reagent-less biosensing (via immobilizing receptor molecules), biomimetic, in-vivo detection; (iv) high surface area and superparamagnetic activity of nanostructured materials allows them to be used as carrier or capture-vehicles for loading a large amount of specific biological probes, and work as ‘dispersible capture agents’ for isolating and capturing circulating disease specific markers in body fluids; (v) optical (surface- enhanced Raman spectroscopy-SERS, surface plasmon resonance-SPR, and florescence) and electrochemical properties of NPs can be exploited to adopt many novel transduction schemes, and (vi) peroxidase-like activity of many materials has found a wide range of applications in ELISA-based sensing approaches. These unique properties of nanostructured materials have 117

also been widely explored for achieving sensitive biosensing of miRNAs.45 Figure 2.2.1 depicts the common nanostructures widely used in miRNA biosensing.

Gold nanoparticles. Gold NPs (AuNPs) have found multifaceted applications in biosensing systems. In addition to their ease of synthesis, the affinity of AuNPs surface with a variety of ligands has not only helped in the development of highly stable AuNPs but also facilitates facile bioconjugation with a variety of biomolecules.46 Furthermore, tunability of AuNP characteristics like conductivity, redox behaviour, and catalytic activity makes them excellent candidates for interfacing nucleotide recognition events with electronic signal transduction.47-48 Pioneering AuNP based DNA detection was reported by Mirkin and colleagues who utilized a distance-dependent change in the extinction spectrum of AuNPs to detect target DNA in a separation-free manner. Two different capture probes, non- complimentary to each other, were attached to two separate batches of AuNPs. Hybridization of target DNA molecules containing sequences complementary to both the surface-bound probes leads to particle aggregation and colour change.49 Since then AuNPs have been integrated into electrochemical nucleic acid-sensing systems, including miRNA detection platforms, in various roles such as (i) tags attached to oligonucleotides which can be electrochemically detected after acidic dissolution through stripping voltammetry, (ii) carriers/enhancers of other electroactive labels, (iii) as electrocatalytic signal amplifiers, and (iv) as dispersible nano-electrodes which seek out the analyte thereby substantially enhancing the rate of analyte mass transfer to the transducer.48, 50 Regardless of the potential of Au NPs to serve as a label, signalling molecules or dispersible electrode, the use of these materials in biosensing is yet limited because of their lack of long-term colloidal stability. They can easily be irreversibly aggregated and form large particles to reduce the surface energy upon the exposure of certain physical and chemical changes including contact with biofluids, freeze- drying, and buffer solutions. The stability of NPs is prerequisite for the use them in biosensing, which can be achieved by functionalizing with polymers (such as PEG), ligands (protein receptor) or surfactants.42,43 Another way for achieving stability is to form hybrid or bi-metallic NPs, where Au NPs the Au NPs didn't get enough space to interact and also possess intermolecular repulsion.45,46

Strong thiol-gold interaction provides the basis of many nucleic acid biosensors where thiol-derivatized nucleic acid fragments are used to modify AuNPs.51 Furthermore, direct adsorption of both single-stranded and double-stranded nucleic acids has also been observed 118

and several nucleic acid biosensors have been developed based on this principle.52-53 Direct adsorption arises from electrostatic interactions, hydrophobic forces, and sequence- dependent (DNA base-Au) affinity interactions. Over the past few years, several electrochemical miRNA sensors have been developed by our group as well as other researchers, where magnetically purified target miRNAs are directly adsorbed onto the gold electrode surfaces for subsequent electrochemical interrogation, which results in the sensitive detection.54-58 In addition to these functionalities, several amplification strategies coupled with AuNPs have been devised and have achieved ultra-sensitive miRNA detection.59-60

Figure 2.2.1: Commonly used nanoarchitecture in mirna biosensing. Schematic representation of nanostructured materials widely used in developing electrochemical miRNA biosensors. Abbreviations AgNPs/SWCNT, silver nanoparticles/ single-walled carbon nanotubes nanohybrid; Au-Fe2O3NC, Au-loaded iron oxide nanocube; AuNPs, gold nanoparticles;

AuNPs@MoS2, AuNPs decorated over molybdenum disulphide; Fe3O4@Ag Shell, core-shell nanostructure with magnetic Fe3O4 as core and Ag metallic shell; GO, graphene oxide; PdPt, palladium platinum; QD, quantum dot; Strep- MB, streptavidin-magnetic bead.

Silver nanoparticles. Silver NPs (AgNPs) are another class of metal nanoparticles which have found application in a range of electrochemical and optical sensor platforms owing to their easy signal amplification, higher extinction coefficients, higher scattering to extinction ratio, and high field enhancements. AgNPs and their network structure are easily oxidized, show well-defined amplified signals even at a lower potential, enabling their use as redox tags for electrochemical detection.61 For instance, AgNPs aggregates were employed as labels on

119

electrode surfaces for label-free (signal-on) electrochemical miRNA sensors.62 Moreover, AgNPs demonstrated higher extinction coefficient, stronger Raman and fluorescence enhancement compared with that of AuNPs of the same size, leveraging several developments in miRNA sensing technologies. These approaches are based on plasmonic properties of AgNPs, e.g., platforms incorporating SERS or localized surface plasmon resonance (LSPR) or metal-enhanced fluorescence (MEF) readouts. Oligonucleotides and ligand modified Ag nanocluster building blocks or tags, having luminescent properties and bio-specificity for nucleic acids, have been used in electrochemiluminescent miRNA sensors.63 Additionally, positively charged AgNPs can also be integrated with negatively charged gold loaded DNA nanostructures, resulting in an increased resonance shift for amplified SPR signals.62 Recently, based on Ag nanostructure such as the DNA-mediated Au-Ag nano-mushroom, as well as Au@R6G@AgAuNPs (rodhamine-6-G attached on AuNPs and then encapsulated in AgAu alloy shell NPs) as SERS signal reporters, several miRNA sensors have been reported for simultaneous and multiplexed detection of numerous RNA targets.64-66

Carbon and carbon-based nanomaterials. Carbon nanomaterials such as carbon nanotubes (CNTs), graphene, graphene oxide (GO), carbon nano-fibre, and carbon quantum dots (QDs) offer attractive opportunities for developing novel sensors and refining the analytical performance of already existing platforms. The intrinsic characteristics of CNTs such as their hollow core, high elastic modulus, as well as diverse electron transport capability make this material an ideal candidate as a building block, sensing layer, a surface modifier, or dispersible capture agent for loading a large number of biomolecules or assembling labels for signal amplification.67 In addition, CNTs are fluorescent in the near-infrared spectral region (tissue penetrating region) and their emission wavelength and intensity are sensitive up to single- molecule attachment, permitting perturbations at the nanotube surface to be transduced via modulation of their emission. Based on such (single-walled carbon nanotubes-SWCNT) modulation, an in vivo miRNA sensor have been fabricated for multiplex detection through multiple nanotube chiralities.68 In addition, graphenes (GO, reduced graphene oxide-rGO, graphite-GR) have unique electronic, adsorption, and fluorescence properties, and are emerging as powerful key foundations of miRNA sensors.69-71 The GO surface possesses many carboxylic acid and hydroxyl groups, making it more water-soluble and suitable for simple covalent functionalization and biomolecule adsorption.72 Therefore many recent biosensors have utilized GO and GO loaded metal composites to establish miRNA sensing platforms.73

120

Quantum dots. Quantum dots (QDs) are fluorescent semiconductor nanocrystals which have gained increasing application in the construction of electrochemical biosensors mainly because of their high electron transfer efficiency and high surface reaction activity. Their robust photo- stability and size-tunable light emission spectra also make them ideal labels for optical and chemiluminescent sensors. Several miRNA detection methods based on photophysical properties of QDs have been reported.74-75 Zhang et al. used graphene QDs as a substrate to immobilize horseradish peroxidase (HRP) enzyme onto a target miRNA sequence containing a dsDNA structure.76 Similarly, owing to their user-friendly bio-labelling capability, QDs have found application in paper-based biosensors. Deng et al. utilized QDs in a target recycled nonenzymatic-amplification-strategy-based strip (paper) sensor for obtaining efficient, reproducible, and portable detection of miRNA.77

Magnetic nanoparticles. Integration of magnetic materials with conventional NP-based miRNA detection strategies cannot only enhance their sensitivity and selectivity but also offers a low-cost miniaturization platform. In the absence of an external magnetic field, magnetic nanoparticles show negligible magnetic moment (superparamagnetism), which allows the formation of a stable colloidal suspension that can be manipulated and engineered by applying a magnetic field without any reversal aggregation. Having such superparamagnetic properties, easy bio-functionalization, and high surface-to-volume ratio, magnetic NPs facilitate magnet- based enrichment, capture, isolation, and purification of the target from a complex biomatrix. In addition, they can be directly used as a cost-effective electrode surface modification for biomolecule immobilization, as electrocatalysts for significant signal enhancement by increasing the electron transfer rate, and as a signalling label for sensitive bio-recognition and transducing events.78 Moreover, certain magnetic nanomaterials have been shown to possess intrinsic peroxidase-like activity, which has fuelled their application as labels in colourimetric miRNA sensing.9-80 In the past few years, we have employed gold and graphene oxide loaded superparamagnetic iron oxide nanocubes as a transducer surface and electrocatalysts for ultrasensitive miRNA detection.53-56, 81 Recently, Tavallaie, R. et al. utilized redox-labelled probe DNA modified magnetic NPs for sequence-specific hybridization with a target.50 However, while the rate of analyte capturing depends on the affinity towards the biomolecules and the total surface of magnetic particles, the higher amount of nanomaterials may increase the probability of particle aggregation and risk of non-specific adsorption. Therefore, a lesser amount of particles with a high surface-to-volume ratio (e.g., porous nanomaterials) and higher bio-affinity could be an excellent choice for biosensing. 121

Metal-organic frameworks. Metal-organic frameworks (MOFs) are a large family of crystalline materials with organic multidentate ligands as linkers and metal ions as nodes, which can form 3D frameworks with unique properties including high pore volume, large surface area, and extraordinary thermal stability. MOFs can bind with the negatively charged probe DNA sequences through electrostatic interactions and (or) π-stacking, presenting them as sensitive tracer labels for biosensing. Moreover, they are highly advantageous in biosensing as they provide high-loading of probe DNA and show resistance against probe DNA degradation82 MOFs are widely employed as quenching labels for disease-specific nucleic acid detection. In such strategies, the organic ligands tightly bind the probe DNA while the metal ions working as a coordination centre activate their intrinsic fluorescent quenching capability through a photo-induced electron transfer process83 Moreover, the presence of various functional groups, such as -COOH and -NH2 groups, makes MOFs an excellent platform for loading different signalling metal substrates and biological ligands.84 The structural modification of MOFs with other functional materials such as AuNPs, AgNPs, CuNPs (copper NPs), PdNPs (palladium NPs) or RuNPs (ruthenium NPs)produces a superior entity due to synergistic effects of the combined material compared with pure MOFs.85-86 Among them, Cu- MOFs have received special attention in electrochemical applications because of their well- defined configuration, unique electrical conductivity, and superior catalytic activity.87 For example, Wang and co-authors fabricated a paper-based electrochemical miRNA sensor using hairpin assembly target recycling for signal amplification and AuNP modified Cu-based metal- organic frameworks (Cu-MOFs) for catalysis.88

2.2.3. Nanoarchitecture based electrochemical miRNA detection Electrochemistry based readout has stimulated great interest in recent years mainly due to its inherent sensitivity, specificity, simplicity of operation, short response time, and broad dynamic range (over nine orders of magnitude greater than many nucleic acid biosensors). Additionally, electrochemistry based methods are portable and highly amenable to miniaturization, which can prove to be highly beneficial in achieving massively multiplexed electrochemical devices. Thus, electrochemistry based devices have the potential to solve most of the analytical challenges hampering the development of a point-of-care device for a miRNA biomarker. In these devices, electrode materials play the critical role in obtaining high- performance data via various electrochemical techniques such as voltammetry (CV: cyclic voltammetry, DPV: differential pulse voltammetry, LSV: linear sweep voltammetry, etc.), 122

coulometry, amperometry (i-t), and impedance spectroscopy. Incorporating engineered nanomaterials with common electrode surfaces can not only give synergistic effects related to biocompatibility, magnetism, catalysis, and conductivity but also provide accelerated signal transduction and amplified bio-recognition events, resulting in an ultrasensitive biosensing platform. Over the last few decades, significant research has been reported on the construction of various nanomaterials for the fabrication of functional electrode surfaces, as signalling tags or as electrocatalysts.89-91 Here, we review the pros and cons of recent (2015 to the present) NP-integrated electrochemical miRNA assays. Within this timeframe, all strategies from the literature will be discussed, based on their intrinsic properties towards biosensing. Table 2.4 summarises the most commonly used nanomaterials and their applications in miRNA detection in this timeframe and Figure 2.2.2 schematically outlines the major nanoarchitecture-based electrochemical strategies for miRNA detection.

Figure 2.2.2: Nanoarchitecture in miRNA biosensing. Top: miRNAs can be extracted from various biological samples such as cell lines, tumour tissues, organisms, exosomes, serum, plasma, and urine samples. Middle: after performing the extraction of total small RNA (i.e., a

123

pool of miRNAs) using a commercial extraction kit-based procedure, the target miRNA is isolated and purified from the miRNA pool using a capture probe bound magnetic bead (MB), adapted with the permission from [57]. Herein, streptavidin-coated MB bind with the biotinylated ed capture probe followed by the sequence-specific hybridization of capture probe with target miRNA. Heat release of isolated and purified hybrid gives the target miRNAs for subsequent electrochemical analysis. Bottom: nanostructure material act both as (left) electrode surface modifier and (right) signal amplification tags/labels. In the left figure, gold-loaded nanoporous ferric oxide nanocube (Au@NPFe2O3NC) is magnetically bound onto a screen- printed carbon electrode. A target miRNA is directly adsorbed onto the surface of the electrode- bound Au@NPFe2O3NC via affinity interaction between the Au and miRNA nucleobases. In the right figure, a target-specific capture probe pre-immobilized onto an electrode surface interacts with graphene nanosheet containing a large number of electroactive molecules, resulting in an enhanced electrochemical response. In the presence of target miRNAs, a successful hybridization event between the immobilized capture probes and target miRNAs reduces the attachment of graphene nanosheet based electroactive molecules onto the electrode surface, leading to a significant reduction in electrochemical response.

Nanostructures as a capture agent or vehicles for miRNA isolation and purification. Nanostructures with superparamagnetic properties are becoming a promising tool for the capture, and separation of biomolecules from their complex biological matrixes. These tools offer significant advantages over conventional separation techniques such as chromatography, because they possess enhanced functional area, can readily disperse in solution, and are quickly localized or retrieved using an external magnet. The materials can be synthesized at low cost and can be reused several times in bioassays.92 Over the last three years, we have developed several electrochemical sensors employing streptavidin-labelled magnetic beads (Dynabeads® MyOne™) for isolating target miRNAs.53-58 The 1-µm size beads possess a large surface area, -15 high binding capacity for avidin-biotin interaction (dissociation constant, Kd = 10 ), proficient magnetic pull, and slow sedimentation during incubation. In such bead-based isolation and purification steps (Figure 3.2.2), the magnetic beads are functionalized with complementary capture probes through biotin-avidin interactions, followed by the hybridization of target miRNAs. The bead-bound target miRNAs then undergo multiple washing and purification steps. The target is then heat released and subjected to electrochemical readouts.57 Trau et al. reported an amplification-free simultaneous detection (fluorescence and electrochemical) of RNA species, including miRNA, using parallel magnetic isolation of individual RNA targets, 124

and in situ poly(A-T)-templated CuNB synthesis.93 Very recently, an electric-field-induced reconfigurable gold-coated magnetic NP (Au@MNPs) as dispersible electrodes was reported by J. Gooding et al. for direct miRNA analysis (Figure 2.2.3(i)). In this remarkable report, the thiol-modified probe DNA (complementary to target miRNA) were mixed with an excess amount of Au@MNPs followed by mixing with samples for target miRNA hybridization.50 Au@MNPs-DNA/miRNA hybrid was then separated magnetically from unhybridized sequences and other biomatrix and collected onto a gold microelectrode surface. The square- wave voltammetric responses from the electronic reconfiguration of Au@MNPs before and after hybridization were taken to analyze the target present in samples. However, magnetic- nanostructure-based separation and purification depend on the shape and size of the magnetic cores and the functional surfaces. The isolation and purification process requires time- consuming multiple incubation steps. Nanostructures with high superparamagnetism (to avoid residual magnetism), enlarged functional surface (porous structure) area, and higher target affinity could be a possible route for robust miRNA isolation and purification.

125

Figure 2.2.3: Nanoarchitecture based electrochemical miRNA biosensing. (i) Representation of Au@MNPs dispersible electrode-based electrochemical miRNA detection. The Au@MNPs modified with a methylene-blue-labelled probe DNA (a), followed by mixing with the sample to capture target miRNA (b). After magnetic isolation and purification of miRNA hybrid (c), Au@MNPs are magnetically collected on to the electrode surface (d), for electrical reconfiguration of Au@MNPs through 10 cycles of square-wave voltammetry (e). (ii) Schematic illustration of Au nanostructure synthesis and electrochemical nucleic acid sensor design; (ii-a) represents the preparation of four different sizes and morphologies gold nanostructures electrode by electrodeposition on indium-tin-oxide (ITO); (ii-b) demonstration of smooth HFGNs gold nanostructure-based electrochemical biosensor for miR-21 detection 3+ 3+ employing Ru(NH3)6 as an electrochemical signal molecule. Ru(NH3)6 molecules were stoichiometrically bounded with the HFGMs/ITO electrode attached anionic phosphate of DNA/miRNA strands and thereby resulting SWV signals for target miR-21. Adapted, with permission, from [50, 99].

126

Table 2.4: Electrochemical miRNA detection based on nanoarchitectures. Nanoarchitecture Function Principle miRNA LOD Ref. AuNPs Selective pre-concentration of AuNPs Differential adsorption ability of nucleic acids let-7A 16 fM 94 on AuNPs AuNPs-MMB and Fc AuNPs-MMB for probe DNA Cyclic voltammetric detection of miRNA based miR-182 0.14 fM 95 capped AuNPs immobilization, target recognition, and on AuNP-MMBs and Fc-capped AuNPs signal magnetic separation; amplification Fc capped AuNPs for target recognition event amplifications.

Au-NPs modified Cu- Immobilization of probe DNA and signal Target miRNA triggered DPV signal miR-21 0.35 fM 88 MOFs amplifications amplification

96 N-doped MoSe2 electrode surface enables loading of Target miRNA assisted supersandwich structure let-7a 0.17 fM graphene/AuNPs and AuNPs for probe DNA and NG-AuNPs for hemin redox signal amplifications. flower-like MoSe2 employed as electrocatalyst towards the oxidation of hemin. PTh/rGO PTh/rGO serves as immobilizer for probe Nanocomposite based dual signal amplification miR-106a 0.06 fM 59

Au/TMC/Fe3O4 DNA and augmented electrode conductivity, for simultaneous detection of two miRNAs let-7a 0.02 fM CdSe@CdS/TMC/ and both nanocomposites acted as selective

Fe3O4 labels for two miRNAs

Cd2+ functionalized TiP nanospheres containing a large amount Target assisted sandwich structure for capturing miR-21 0.76 aM 97 titanium phosphate of Cd2+ acted as stripping label signal tags. nanosphere

127

(+)AuNPs Positively charged AuNPs stoichiometrically Dual mode (EIS and SWV) electrochemical miR-45 0.37 fM 59 bind with the negatively charged miRNA detection and attached ferrocene for SWV detection.

3+ 98 Flower-Like Gold Hierarchical Stoichiometric amount sof Ru(NH3)6 bound to miR-21 1 fM Nanostructures gold nanostructures for immobilization of a anionic phosphate of large amount of cDNA DNA/miRNA sequences provide electrochemical readout.

99 AuNPs@MoS2 As both electrode modifier and Nanocomposite based sandwich structure miR-21 0.78 fM nanoamplifier. empowers electrochemical signals using

3-/4- 3+ [Fe(CN)6] and [Ru(NH3)6] CNT Solid substrate for solid-state RCA strategy Target miRNA unfolded the stem-loop structure let-7 1.2 fM 100 and triggered CNT based RCA process family

AuNPs and hemin-rGO Achieving electrochemical transducing Decrease in the DPV response of hemin cDNA 0.14 aM 101 interface on GCE resultant dsDNA obtained via hybridization of probe DNA with complementary DNA

102 CoFe2O4 MNPs Nanoelectrocatalyst for toluidine blue (Tb) Target miRNA hybridizes with the padlock miR-21 0.3 fM catalysis. probe, resulting in the generation of padlock exponential rolling circle amplification (P- ERCA) products for circular exponential signal amplification.

128

103 Fe3O4 NPs To carry a large number of redox signalling Target-triggered HCR strategy for capturing miR-141 0.28 fM molecules thionine and Fc for obtaining large amounts of specific magnetic probes for and miR- 0.36 fM target specific DPV readout. simultaneous detection of miRNAs. 21.

104 MoS2-Thi-AuNPs Functionalization of GCE to probe DNA- Hybridization of the target with probe DNA miR-21 0.26 pM nanocomposites RNA complex. greatly hinders the electron transfer on the electrode, resulting in a decrease of thionine signals.

33 Au@NPFe2O3NC Immobilizing target miRNAs on AuNPs on Nanocube assisted electrocatalytic miR-21 1 pM nanocube and electrocatalysis of surface- chronocoulometric reduction of target miRNA

3+ 3+ bound Ru(NH3)6 bound reduction of Ru(NH3)6 GO-loaded-iron oxide Immobilizing target miRNAs on GO and Chronocoulometric readout of charge‐ miR-21 100 aM 56

3+ hybrid materials GO/IO based electrocatalysis of surface- compensating [Ru(NH3)6] followed by an 3+ bound Ru(NH3)6 enhancement in CC charge display through the 3+ 3− Ru(NH3)6] /[Fe(CN)6] system.

DNA-Au@MNPs Au@MNPs as dispersible electrodes square-wave voltammetric responses from the miR-21 10 aM 50 electric-field-induced reconfiguration of target bound DNA–Au@MNPs

PdNPs@Fe-MOFs Nanocarriers for signal probe Streptavidin-PdNPs@Fe-MOFs as tracer miR-122 0.003 fM 86 immobilization and redox probes and anchored on a target based sandwiched structure electrocatalysts for signal enhancement for electrocatalytic oxidation of TMB (in

presence of H2O2)

Abbreviations: CdSe@CdS, cadmium diselenide-cadmium sulphide; CNT, carbon nanotube; CoFe2O4, cobalt ferrite; Fc, ferrocene; MMB, magnetic microbeads; MNPs, magnetic NPs; MOFs, metal-organic frameworks; MoS2, molybdenum-disulfide; MoSe2, molybdenum diselenide; PdNPs, palladium NPs; PTh, polythiophene; rGO, reduced graphene oxide; Thi, thionine; TMC, N-trimethylchitosan polymer.

129

Nanostructures for miRNA immobilization. Immobilization of reporter or target miRNA on the transduction surfaces is one of the key requirements for designing electrochemical sensors. This immobilization step can be affected by denaturation, low hybridization efficiency, or poor transducing capabilities. Engineered nanostructures have been integrated on electrode surfaces as they provide a high surface area, a high density of capture sites, and high hybridization moiety for faster and efficient analyte binding, and enhanced amplification agent incorporation. Nevertheless, immobilization steps need to be carefully optimized to protect the nanostructured surface from undesired adsorbents and the ambient environment. The merits of the nanostructured electrode surface greatly depend on the size and morphology, however. Kelly research group et al. elaborately leveraged the effect of nanostructured features on the substrate angle of deflection, hybridization efficiency, and biomolecule–electrode connections.105-107 In 2009, they reported engineered highly nanostructured palladium microelectrodes (Pd NMEs) which were modified with PNA probes and then exposed with total RNA for hybridization. The microelectrode system was coupled with an electrocatalytic Ru3+/Fe3+ reporter system and achieved a detection limit of 10 aM for miR-21 electrochemically.108. In the following year, by manipulating the deposition time, applied potential, metal concentration, and electrolyte, three nanostructured palladium (II) electrodes with smooth hemispherical, moderately nanotextured (100-150 nm size), and finely nanotextured (20 -30 nm) electrodes were engineered.105 The highly nanostructured electrode with a surface coverage of ssDNA (∼9 × 1012 molecules/cm2) exhibited a high hybridization efficiency, while smooth hemispherical one showed lower hybridization efficiencies. Recently, Su et al. manufactured shape-controlled hierarchical flower-like gold nanostructures (HFGNs) on indium tin oxide (ITO) electrode for miR-21 detection from human lung cancer cells (A549) and achieved a LOD of 1 fM (Figure 3 (ii)).98 Although these nanostructures presented an excellent platform for sensing miRNA, the chemistry involved in fabrication steps limits their application in robust and multiplex point- of-care (POC) devices. The non-metallic nanostructures such as carbon derivative materials (CNT, SWCNTs, graphene, GO) have also become promising choices for the fabrication of recognition platforms, owing to their easy functionalization, fast electron transfer, physisorption capability, e.g. π-π noncovalent interactions, etc. For instance, CNTs as an excellent electron transfer electrode material were used as a signal transducer and solid primer for DNA hairpin probe immobilization for miRNA detection.100. GO contains carboxylic acid, and carbonyl and epoxy groups, and it has better solubility in aqueous and organic solvents. This material has been utilized for the modification of glassy carbon electrode, pencil electrodes, or screen-printed 130

carbon electrode (SPCE). The GO modified electrode surface offers a significantly enhanced surface for better interaction between capture probes and target miRNAs. The capture probes are easily immobilized at the GO surface and facilitate the hybridization with target miRNAs.109 Due to the high content of oxygenated functional groups, however, GO needs to be reduced to form rGO to retrieve the sp2 network and facilitate electron transport. This functionality can be easily achieved by reducing the GO electrochemically.110 The electrical conductivity can be further enhanced by introducing nitrogen atoms into the lattice of carbon during the reduction steps. Recently, a nitrogen-doped rGO (NRGO) was coupled with chitosan for increased adsorption capability to obtain a synergetic sensing platform for miRNA detection.111 In another recent report, the rGO electrode surface conductivity was improved by modifying SPCE with polythiophene (PTh) and rGO.112 In this work, the PTh and rGO coated SPCE provided an optimized platform for successful immobilization of capture probes and subsequent hybridization, signal amplification, and electrochemical readout steps. The combination of GO and gold nanorod (GNR) was also employed for enhancing the sensitivity of nanobiosensors, as both GO sheets and GNR provide high electron transport and a large surface area, while GNR facilitates easy and stable probe immobilization. Azimzadeh et al. introduced a thiolated probe-functionalized GNR decorated graphene oxide (GO) sheet on the surface of the glassy carbon electrode (GCE) for miR-155 detection.113 In this work, the increase in current responses to the intercalating agent oracet blue (OB) upon the hybridization of the target miRNA was measured by differential pulse voltammetry (DPV).

Nanostructures for electrochemical signal amplification. Functional nanostructures can not only generate a synergetic effect involving catalytic activity, conductivity, and biocompatibility to accelerate the signal transduction, but also amplify biorecognition events with specifically designed signal tags, leading to highly sensitive detection performance. Gao et al. first utilized isoniazid-capped OsO2 NP as a tag for obtaining enhanced signals for electrocatalytic oxidation of hydrazine and achieved a LOD of 80 fmol/droplet.114 Following this, numerous NPs have been employed in electrochemical sensors as signal amplifying tags for miRNA detection [59-61]59-60, 115. As discussed earlier, the pre-concentration of oligonucleotides onto AuNP surfaces or the controlled manipulation of AgNP aggregation in the colloidal solution can be integrated with the electrochemical transduction surface to obtain amplified signals for target miRNAs. The ssDNA can easily overcome the repulsion between the negative charge of the phosphate backbone and citrate ions of AuNPs and stick to the AuNPs, while the stronger repulsion and rigid structure of dsDNA make the adsorption onto 131

AuNPs unfavourable. Therefore, in the absence of target miRNAs, the probe ssDNA adsorbed onto the AuNPs and ssDNA-AuNPs were unable to bind with a 1, 6- hexanedithiol modified 3- Au electrode (HMAuE), resulting in significantly lower amperometric signals of [Fe(CN)6] .94 When the target miRNA hybridized with probe ssDNA to form a duplex strand, the AuNPs exist in a free-state and attached on HMAuE successfully, producing a significantly improved amperometric signal. Despite its simple assay design and portability, this assay is limited by poor detection sensitivity (16 fM). Encapsulation of Ag nanoclusters (NCs) onto oligonucleotides adsorbed on AuNPs could enhance the sensitivity, although the encapsulation involves complex and time-consuming chemistry.59 As an alternative, AgNP aggregates can be employed with specific recognition elements as electrochemical reporters. Recently, Liu et al. presented a 4-mercaptophenylboronic acid (MPBA) induced citrate-capped AgNP aggregation-based amplification strategy where boronate ester covalent interaction occurred between the MPBA and cis-diol at the 3′-terminal of miRNAs.116 In this strategy, target miRNAs were hybridized with a thiolated hairpin-like DNA probe assembled on an AuNP modified electrode. The MPBA was then anchored with the 3′-terminal of miRNAs to capture AgNPs. Meanwhile, free MPBA molecules in solution induced the in situ assemblies of AgNPs on the electrode surface, resulting in an enhancement of the electrochemical signals. The method could successfully detect a 20 aM level of miR-21 in human serum samples. Another strategy for obtaining ultra-low miRNA detection is to integrate nanostructures with enzymatic amplification steps such as electrocatalysis.91 the polymerase chain reaction,63 rolling circle amplification,117 isothermal amplification,118 the hybridization chain reaction, and duplex-specific nuclease (DSN) amplification.63 Controlled aggregation of AuNPs has been integrated with triple signal amplification involving miRNA triggered DSN - catalyzed cleavage cycles, bridge DNA−AuNPs, and AuNP based electrochemical species enrichment for ultrasensitive detection of miR-21.115 Upon the addition of target miRNA, the hairpin structure of the DNA probe on the Au electrode surface was opened, forming a DNA/RNA duplex, thereby facilitating the DSN digestion of duplex DNA, as shown in Figure 2.2.4. The remaining ssDNA sequence was then hybridized with the single-stranded ends of three AuNPs 3+ containing bridge DNA, which adsorbed a significantly large amount of [Ru(NH3)6] for obtaining enhanced chronocoulometric (CC) readouts. This ‘turn-on” (as target miRNA activates the signals) assay design achieved an ultra-high sensitivity (LOD of 6.8 aM) without the need for tedious thermal cycling or reverse transcription processes. The use of gold composites such as GO-AuNP hybrid as a signal carrier has been reported for electrochemical– chemical–chemical (ECC) detection of miRNA using MgO-nanoflowers/AuNPs as a sensing 132

platform.119. This strategy is advantageous for achieving high sensitivity as it utilizes triple mode signal enhancement steps (ECC) and has achieved a LOD of 50 aM.

Figure 2.2.4. Signal amplification. Triple signal amplification strategy for ultrasensitive determination of miRNAs based on duplex-specific nuclease (DSN) and bridge DNA−Gold NPs; adapted, with permission, from [61]. DNA nanocomposite was formed using specially designed bridge DNA and DNA probes modified AuNPs. In the absence of target miRNAs, the hairpin probes attached on a gold electrode were inactivated while the target miRNA opened the hairpin structure, allowing the formation of DNA/RNA duplex and digestion of duplex by a DSN (DSN- an enzyme, preferentially cleave the dsDNA and DNA in DNA- RNA hybrid duplexes but not to ssDNA or single or double-stranded RNA). The remaining DNA sequence on the electrode surface became activated and hybridized with DNA 3+ nanocomposites, facilitates the absorption of numerous [Ru(NH3)6] for generating enhanced chronocoulometric responses.

Due to the enzyme mimetic and nanoelectrocatalyst merits of magnetic nanomaterials, they have been widely used in electrochemical miRNA sensing for catalyzing redox markers.

Yuan et al. reported two distinguishable magnetic nanoprobes (DNA1/Fe3O4 NPs/Thi and 103 DNA2/Fe3O4 NPs/Fc) for simultaneous detection of miR-141 and miR-21. Upon the hybridization of the target miR-141 and miR-21 with the thiol-modified hairpin capture probes (HCP1 and HCP2) on the Au electrode surface, the hybridization chain reaction (HCR) generates a large number of sequences for binding magnetic nanoprobes. Boosted DPV responses were then obtained for plentiful magnetic nanoprobes attached to thionine (Thi) and

133

ferrocene (Fc) for target miRNAs by target-triggered HCR and the catalytic responses of magnetic nanoprobes. A similar nanocomposite (Au@CoFe2O4/Tb-Gra composites) tagged padlock exponential rolling circle amplification (P-ERCA) assay has also been reported for miR-21.102 CNTs can interact with DNA through the π–π stacking of DNA aromatic bases and van der Waals forces, but dsDNA tends to stack less than ssDNA. Based on this principle, Asadzadeh-Firouzabadi et al. designed an electrochemical nanogenosensor for miR-25 using an AgNP/SWCNT nanohybrid as a label.120 AuNP-modified Cu-based metal-organic frameworks (AuNPs@Cu-MOFs) as nanocarriers and catalyst have been reported for miR-155 detection using a target-triggered signal amplification cycling strategy.88

Recently, we have utilized specially engineered Au@NPFe2O3NC for SPCE modification to achieve amplification-free, non-enzymatic, and electrocatalytic miRNA-21 3+ detection using [Ru(NH3)6] as an intercalating agent, and achieved a 1.0 pM level of detection.54 Ru3+ is an excellent electrochemical reporter, although it does not yield high sensitivity because only one electron can be accepted by each Ru3+ acceptor. The limited concentration of Ru3+ is also inadequate to produce an enhanced signal. To increase the 3+ 3- sensitivity, the [Ru(NH3)6] system was later combined with [Fe(CN)6] , where the 3+ electrocatalytic reduction of miRNA intercalated [Ru(NH3)6] subsequently reduces the 3- 55 3− solution-phase [Fe(CN)6] , as shown in Figure 3.2.5. The negative charge of Fe(CN)6] prevents this complex from reaching the surface of the electrode because of the electrostatic repulsion of the negatively charged phosphate backbone. Thus, the Fe3+ complex remains in solution and acts as an oxidant when it interacts with outwardly diffusing Ru2+, regenerating Ru3+. By regenerating Ru3+, one bound Ru3+ complex can interact with the electrode on multiple occasions, causing a significant amplification of the target responding chronocoulometric signal (Figure 3.2.5). This Au-NPFe2O3NC catalyzed and 3+ 3 [Ru(NH3)6] /[Fe(CN)6] redox cycling amplified assay design for miR-107 achieved a LOD of 100 aM. Very similar assay for miR-21 detection was also reported using another graphene‐ oxide‐loaded iron oxide (GO/IO hybrid material).56, 81

134

Figure 2.2.5: Schematic representation of Au-Fe2O3NC catalyzed miR-107 assay using 3+ 3 [Ru(NH3)6] /[Fe(CN)6] . Target miR-107 were magnetically isolated and purified from cell lines followed by adsorption on an Au-Fe2O3NC modified SPCE. The electrocatalytic 3+ activity of Au-Fe2O3NC was used for the reduction of [Ru(NH3)6] bound with target miRNA. 3-/4- The catalytic signal was further amplified by using the [Fe(CN)6] system. Adapted with the permission from [55].

2.2.4. Concluding remarks and future perspectives With the increasing recognition of the potential of miRNAs as diagnostic and prognostic biomarkers, recent years have witnessed a rapid surge in novel molecular and nanotechnology- based miRNA detection technologies. The momentous progress on nanostructures as nanocarriers, catalysts, conductive surface modifiers, probe immobilizers, and signal generators and enhancers, as well as the simplicity, robustness, ultra-sensitivity, miniaturization, and cost-effectiveness of electrochemical readout, are continuing the dynamic progress on nanostructure-based EC miRNA sensors. This review has summarized the remarkable achievements of recently (2015 to 2018) reported engineered nanostructures for the development of electrochemical miRNA sensors. Moreover, with a few illustrations of RNA diagnostics and prognostics, the review has described the challenges and needs involved in current miRNA detection strategies and offered an outlook for the near future of the presented assay designs. The advances in nanostructure-based electrochemical miRNA quantification reported in the recent scientific literature are a sign of flourishing research, although a great deal of work still needs to be performed to make it suitable for a point-of-care platform. To date, most sensors are merely proof-of-concept demonstrations and highly dependent on laboratory-based set-ups. We suggest that a number of factors, such as a proper selection of the miRNA source,

135

the right extraction procedure, designing special primers, avoiding amplification bias, etc., should be carefully considered to avoid inconsistencies in RNA assays. It is also obvious that the majority of the reported nanostructures involve multi-step preparation processes that require complex probe functionalization and tagging steps. Moreover, two or more NPs are commonly used to form a composite to achieve nanostructures that have high signal enhancement capacity as well as a large functional surface area. These steps can be simplified by engineering porous composite nanomaterials that have an enlarged surface area. Single-step nanostructure synthesis with in-situ probe functionalization and easy electrochemical transduction and readout could also be considered for future nanostructure-based miRNA sensing. Incorporation of target-specific multifunctional nanostructures and miniaturized electrochemical readouts into portable paper-based microfluidics and lab-on-a-chip in an easy and user-friendly format needs to be explored for reducing the burdens (cost, complication, time frame) associated with the existing miRNA diagnostic technologies. An automated miRNA diagnostic platform that can perform without any human intervention is the future platform that may find applications beyond the classical laboratory. On that note, we believe, that further developments in miniaturized nanostructure-based electrochemical miRNA analysis platforms will shape the future of personalized disease diagnostics both in research and clinical settings.

2.2.5. References 1. Islam, M. N.; Masud, M. K.; Haque, M. H.; Hossain, M. S. A.; Yamauchi, Y.; Nguyen, N. T.; Shiddiky, M. J., RNA biomarkers: diagnostic and prognostic potentials and recent developments of electrochemical biosensors. Small Methods 2017, 1 (7), 1700131. 2. Bhat, M.; Robichaud, N.; Hulea, L.; Sonenberg, N.; Pelletier, J.; Topisirovic, I., Targeting the translation machinery in cancer. Nature Reviews Drug Discovery 2015, 14 (4), 261-278. 3. David, C. J.; Manley, J. L., Alternative pre-mRNA splicing regulation in cancer: pathways and programs unhinged. Genes & Development 2010, 24 (21), 2343-2364. 4. Hayes, J.; Peruzzi, P. P.; Lawler, S., MicroRNAs in cancer: biomarkers, functions and therapy. Trends in Molecular Medicine 2014, 20 (8), 460-469.

136

5. Bertoli, G.; Cava, C.; Castiglioni, I., MicroRNAs: new biomarkers for diagnosis, prognosis, therapy prediction and therapeutic tools for breast cancer. Theranostics 2015, 5 (10), 1122. 6. Cooper, T. A.; Wan, L.; Dreyfuss, G., RNA and disease. Cell 2009, 136 (4), 777-793. 7. Schmitt, A. M.; Chang, H. Y., Long noncoding RNAs in cancer pathways. Cancer cell 2016, 29 (4), 452-463. 8. Lee, R. C.; Feinbaum, R. L.; Ambros, V., The C. elegans heterochronic gene lin-4 encodes small RNAs with antisense complementarity to lin-14. Cell 1993, 75 (5), 843- 854. 9. Filipowicz, W.; Bhattacharyya, S. N.; Sonenberg, N., Mechanisms of post- transcriptional regulation by microRNAs: are the answers in sight? Nature Reviews Genetics 2008, 9 (2), 102. 10. Cai, Y.; Yu, X.; Hu, S.; Yu, J., A brief review on the mechanisms of miRNA regulation. Genomics, Proteomics & Bioinformatics 2009, 7 (4), 147-154. 11. Turchinovich, A.; Samatov, T. R.; Tonevitsky, A. G.; Burwinkel, B., Circulating miRNAs: cell–cell communication function? Frontiers in Genetics 2013, 4, 119. 12. He, L.; Hannon, G. J., MicroRNAs: small RNAs with a big role in gene regulation. Nature Reviews Genetics 2004, 5 (7), 522. 13. Yekta, S.; Shih, I.-h.; Bartel, D. P., MicroRNA-directed cleavage of HOXB8 mRNA. Science 2004, 304 (5670), 594-596. 14. Chakraborty, C.; Sharma, A. R.; Sharma, G.; Doss, C. G. P.; Lee, S.-S., Therapeutic miRNA and siRNA: moving from bench to clinic as next generation medicine. Molecular Therapy-Nucleic Acids 2017, 8, 132-143. 15. Gebert, L. F.; Rebhan, M. A.; Crivelli, S. E.; Denzler, R.; Stoffel, M.; Hall, J., Miravirsen (SPC3649) can inhibit the biogenesis of miR-122. Nucleic Acids Research 2013, 42 (1), 609-621. 16. Matin, F.; Jeet, V.; Clements, J. A.; Yousef, G. M.; Batra, J., MicroRNA theranostics in prostate cancer precision medicine. Clinical Chemistry 2016, 62 (10), 1318-1333. 17. Pritchard, C. C.; Cheng, H. H.; Tewari, M., MicroRNA profiling: approaches and considerations. Nature Reviews Genetics 2012, 13 (5), 358. 18. Calin, G. A.; Dumitru, C. D.; Shimizu, M.; Bichi, R.; Zupo, S.; Noch, E.; Aldler, H.; Rattan, S.; Keating, M.; Rai, K., Frequent deletions and down-regulation of micro-RNA genes miR15 and miR16 at 13q14 in chronic lymphocytic leukemia. Proceedings of the National Academy of Sciences 2002, 99 (24), 15524-15529. 137

19. Wang, H.; Peng, R.; Wang, J.; Qin, Z.; Xue, L., Circulating microRNAs as potential cancer biomarkers: the advantage and disadvantage. Clinical Epigenetics 2018, 10 (1), 1-10. 20. Lawrie, C. H.; Gal, S.; Dunlop, H. M.; Pushkaran, B.; Liggins, A. P.; Pulford, K.; Banham, A. H.; Pezzella, F.; Boultwood, J.; Wainscoat, J. S., Detection of elevated levels of tumour‐associated microRNAs in serum of patients with diffuse large B‐cell lymphoma. British Journal of Haematology 2008, 141 (5), 672-675. 21. Frei, A. P.; Bava, F.-A.; Zunder, E. R.; Hsieh, E. W.; Chen, S.-Y.; Nolan, G. P.; Gherardini, P. F., Highly multiplexed simultaneous detection of RNAs and proteins in single cells. Nature Methods 2016, 13 (3), 269. 22. Rabinowits, G.; Gerçel-Taylor, C.; Day, J. M.; Taylor, D. D.; Kloecker, G. H., Exosomal microRNA: a diagnostic marker for lung cancer. Clinical Lung Cancer 2009, 10 (1), 42-46. 23. Grasedieck, S.; Schöler, N.; Bommer, M.; Niess, J. H.; Tumani, H.; Rouhi, A.; Bloehdorn, J.; Liebisch, P.; Mertens, D.; Döhner, H., Impact of serum storage conditions on microRNA stability. Leukemia 2012, 26 (11), 2414. 24. Kirschner, M. B.; Edelman, J. J. B.; Kao, S. C.-H.; Vallely, M. P.; Van Zandwijk, N.; Reid, G., The impact of hemolysis on cell-free microRNA biomarkers. Frontiers in Genetics 2013, 4, 94. 25. Poel, D.; Buffart, T. E.; Oosterling-Jansen, J.; Verheul, H. M.; Voortman, J., Evaluation of several methodological challenges in circulating miRNA qPCR studies in patients with head and neck cancer. Experimental & Molecular Medicine 2018, 50 (3), e454. 26. Wang, K.; Yuan, Y.; Cho, J.-H.; McClarty, S.; Baxter, D.; Galas, D. J., Comparing the MicroRNA spectrum between serum and plasma. PloS one 2012, 7 (7), e41561. 27. Mitchell, P. S.; Parkin, R. K.; Kroh, E. M.; Fritz, B. R.; Wyman, S. K.; Pogosova- Agadjanyan, E. L.; Peterson, A.; Noteboom, J.; O'Briant, K. C.; Allen, A., Circulating microRNAs as stable blood-based markers for cancer detection. Proceedings of the National Academy of Sciences 2008, 105 (30), 10513-10518. 28. Leidinger, P.; Backes, C.; Rheinheimer, S.; Keller, A.; Meese, E., Towards clinical applications of blood-borne miRNA signatures: the influence of the anticoagulant EDTA on miRNA abundance. PLoS One 2015, 10 (11), e0143321. 29. McDonald, J. S.; Milosevic, D.; Reddi, H. V.; Grebe, S. K.; Algeciras-Schimnich, A., Analysis of circulating microRNA: preanalytical and analytical challenges. Clinical Chemistry 2011, 57 (6), 833-840. 138

30. Shi, R.; Chiang, V. L., Facile means for quantifying microRNA expression by real-time PCR. Biotechniques 2005, 39 (4), 519-525. 31. Chen, C.; Ridzon, D. A.; Broomer, A. J.; Zhou, Z.; Lee, D. H.; Nguyen, J. T.; Barbisin, M.; Xu, N. L.; Mahuvakar, V. R.; Andersen, M. R., Real-time quantification of microRNAs by stem–loop RT–PCR. Nucleic Acids Research 2005, 33 (20), e179-e179. 32. Takamizawa, J.; Konishi, H.; Yanagisawa, K.; Tomida, S.; Osada, H.; Endoh, H.; Harano, T.; Yatabe, Y.; Nagino, M.; Nimura, Y., Reduced expression of the let-7 microRNAs in human lung cancers in association with shortened postoperative survival. Cancer Research 2004, 64 (11), 3753-3756. 33. Kaur, H.; Arora, A.; Wengel, J.; Maiti, S., Thermodynamic, counterion, and hydration effects for the incorporation of locked nucleic acid nucleotides into DNA duplexes. Biochemistry 2006, 45 (23), 7347-7355. 34. Labib, M.; Khan, N.; Ghobadloo, S. M.; Cheng, J.; Pezacki, J. P.; Berezovski, M. V., Three-mode electrochemical sensing of ultralow microRNA levels. Journal of the American Chemical Society 2013, 135 (8), 3027-3038. 35. Hizir, M. S.; Nandu, N.; Yigit, M. V., Homologous miRNA analyses using a combinatorial nanosensor array with two-dimensional nanoparticles. Analytical Chemistry 2018, 90 (10), 6300-6306. 36. Quarless, S. A.; Cantor, C. R., Analysis of RNA structure by ultraviolet crosslinking and denaturation gel electrophoresis. Analytical Biochemistry 1985, 147 (2), 296-300. 37. Fiammengo, R., Can nanotechnology improve cancer diagnosis through miRNA detection? Biomarkers in Medicine 2017, 11 (1), 69-86. 38. Mestdagh, P.; Hartmann, N.; Baeriswyl, L.; Andreasen, D.; Bernard, N.; Chen, C.; Cheo, D.; D'andrade, P.; DeMayo, M.; Dennis, L., Evaluation of quantitative miRNA expression platforms in the microRNA quality control (miRQC) study. Nature Methods 2014, 11 (8), 809. 39. Witwer, K. W., Circulating microRNA biomarker studies: pitfalls and potential solutions. Clinical Chemistry 2015, 61 (1), 56-63. 40. Singh, R.; Ramasubramanian, B.; Kanji, S.; Chakraborty, A. R.; Haque, S. J.; Chakravarti, A., Circulating microRNAs in cancer: Hope or hype? Cancer Letters 2016, 381 (1), 113-121. 41. Chen, Y.-X.; Huang, K.-J.; Niu, K.-X., Recent advances in signal amplification strategy based on oligonucleotide and nanomaterials for microRNA detection-a review. Biosensors and Bioelectronics 2018, 99, 612-624. 139

42. Beveridge, J. S.; Stephens, J. R.; Williams, M. E., The use of magnetic nanoparticles in analytical Chemistry. Annual Review of Analytical Chemistry 2011, 4, 251-273. 43. Reddy, L. H.; Arias, J. L.; Nicolas, J.; Couvreur, P., Magnetic nanoparticles: design and characterization, toxicity and biocompatibility, pharmaceutical and biomedical applications. Chemical reviews 2012, 112 (11), 5818-5878. 44. Perfézou, M.; Turner, A.; Merkoçi, A., Cancer detection using nanoparticle-based sensors. Chemical Society Reviews 2012, 41 (7), 2606-2622. 45. Xia, N.; Zhang, L., Nanomaterials-based sensing strategies for electrochemical detection of microRNAs. Materials 2014, 7 (7), 5366-5384. 46. Saha, K.; Agasti, S. S.; Kim, C.; Li, X.; Rotello, V. M., Gold nanoparticles in chemical and biological sensing. Chemical reviews 2012, 112 (5), 2739-2779. 47. Turner, M.; Golovko, V. B.; Vaughan, O. P.; Abdulkin, P.; Berenguer-Murcia, A.; Tikhov, M. S.; Johnson, B. F.; Lambert, R. M., Selective oxidation with dioxygen by gold nanoparticle catalysts derived from 55-atom clusters. Nature 2008, 454 (7207), 981. 48. Daniel, M.-C.; Astruc, D., Gold nanoparticles: assembly, supramolecular Chemistry, quantum-size-related properties, and applications toward biology, catalysis, and nanotechnology. Chemical Reviews 2004, 104 (1), 293-346. 49. Mirkin, C. A.; Letsinger, R. L.; Mucic, R. C.; Storhoff, J. J., A DNA-based method for rationally assembling nanoparticles into macroscopic materials. Nature 1996, 382 (6592), 607. 50. Tavallaie, R.; McCarroll, J.; Le Grand, M.; Ariotti, N.; Schuhmann, W.; Bakker, E.; Tilley, R. D.; Hibbert, D. B.; Kavallaris, M.; Gooding, J. J., Nucleic acid hybridization on an electrically reconfigurable network of gold-coated magnetic nanoparticles enables microRNA detection in blood. Nature Nanotechnology 2018, 13 (11), 1066. 51. Cárdenas, M.; Barauskas, J.; Schillén, K.; Brennan, J. L.; Brust, M.; Nylander, T., Thiol-specific and nonspecific interactions between DNA and gold nanoparticles. Langmuir 2006, 22 (7), 3294-3299. 52. Koo, K. M.; Sina, A. A.; Carrascosa, L. G.; Shiddiky, M. J.; Trau, M., DNA–bare gold affinity interactions: mechanism and applications in biosensing. Analytical Methods 2015, 7 (17), 7042-7054. 53. Islam, M. N.; Gopalan, V.; Haque, M. H.; Masud, M. K.; Al Hossain, M. S.; Yamauchi, Y.; Nguyen, N.-T.; Lam, A. K.-Y.; Shiddiky, M. J., A PCR-free electrochemical method

140

for messenger RNA detection in cancer tissue samples. Biosensors and Bioelectronics 2017, 98, 227-233. 54. Masud, M. K.; Islam, M. N.; Haque, M. H.; Tanaka, S.; Gopalan, V.; Alici, G.; Nguyen, N.-T.; Lam, A. K.; Hossain, M. S. A.; Yamauchi, Y., Gold-loaded nanoporous superparamagnetic nanocubes for catalytic signal amplification in detecting miRNA. Chemical Communications 2017, 53 (58), 8231-8234. 55. Islam, M. N.; Masud, M. K.; Nguyen, N.-T.; Gopalan, V.; Alamri, H. R.; Alothman, Z. A.; Al Hossain, M. S.; Yamauchi, Y.; Lamd, A. K.; Shiddiky, M. J., Gold-loaded nanoporous ferric oxide nanocubes for electrocatalytic detection of microRNA at attomolar level. Biosensors and Bioelectronics 2018, 101, 275-281. 56. Islam, M. N.; Gorgannezhad, L.; Masud, M. K.; Tanaka, S.; Hossain, M. S. A.; Yamauchi, Y.; Nguyen, N. T.; Shiddiky, M. J., Graphene‐Oxide‐Loaded Superparamagnetic Iron Oxide Nanoparticles for Ultrasensitive Electrocatalytic Detection of MicroRNA. ChemElectroChem 2018, 5 (17), 2488-2495. 57. Koo, K. M.; Carrascosa, L. G.; Shiddiky, M. J.; Trau, M., Amplification-free detection of gene fusions in prostate cancer urinary samples using mrna–gold affinity interactions. Analytical Chemistry 2016, 88 (13), 6781-6788. 58. Boriachek, K.; Umer, M.; Islam, M. N.; Gopalan, V.; Lam, A. K.; Nguyen, N.-T.; Shiddiky, M. J., An amplification-free electrochemical detection of exosomal miRNA- 21 in serum samples. Analyst 2018, 143 (7), 1662-1669. 59. Jolly, P.; Batistuti, M. R.; Miodek, A.; Zhurauski, P.; Mulato, M.; Lindsay, M. A.; Estrela, P., Highly sensitive dual mode electrochemical platform for microRNA detection. Scientific Reports 2016, 6, 36719. 60. Fredj, Z.; Azzouzi, S.; Turner, A. P.; Ali, M. B.; Mak, W. C., Neutravidin biosensor for direct capture of dual-functional biotin-molecular beacon-AuNP probe for sensitive voltammetric detection of microRNA. Sensors and Actuators B: Chemical 2017, 248, 77-84. 61. Li, H.; Xu, D., Silver nanoparticles as labels for applications in bioassays. TrAC Trends in Analytical Chemistry 2014, 61, 67-73. 62. Liu, R.; Wang, Q.; Li, Q.; Yang, X.; Wang, K.; Nie, W., Surface plasmon resonance biosensor for sensitive detection of microRNA and cancer cell using multiple signal amplification strategy. Biosensors and Bioelectronics 2017, 87, 433-438. 63. Chen, A.; Ma, S.; Zhuo, Y.; Chai, Y.; Yuan, R., In situ electrochemical generation of electrochemiluminescent silver naonoclusters on target-cycling synchronized rolling 141

circle amplification platform for microRNA detection. Analytical Chemistry 2016, 88 (6), 3203-3210. 64. Su, J.; Wang, D.; Nörbel, L.; Shen, J.; Zhao, Z.; Dou, Y.; Peng, T.; Shi, J.; Mathur, S.; Fan, C., Multicolor gold–silver nano-mushrooms as ready-to-use SERS probes for ultrasensitive and multiplex DNA/miRNA detection. Analytical Chemistry 2017, 89 (4), 2531-2538. 65. Ma, D.; Huang, C.; Zheng, J.; Tang, J.; Li, J.; Yang, J.; Yang, R., Quantitative detection of exosomal microRNA extracted from human blood based on surface-enhanced Raman scattering. Biosensors and Bioelectronics 2018, 101, 167-173. 66. Miao, X.; Cheng, Z.; Ma, H.; Li, Z.; Xue, N.; Wang, P., Label-free platform for microRNA detection based on the fluorescence quenching of positively charged gold nanoparticles to silver nanoclusters. Analytical Chemistry 2017, 90 (2), 1098-1103. 67. Pasinszki, T.; Krebsz, M.; Tung, T. T.; Losic, D., Carbon nanomaterial based biosensors for non-invasive detection of cancer and disease biomarkers for clinical diagnosis. Sensors 2017, 17 (8), 1919. 68. Harvey, J. D.; Jena, P. V.; Baker, H. A.; Zerze, G. H.; Williams, R. M.; Galassi, T. V.; Roxbury, D.; Mittal, J.; Heller, D. A., A carbon nanotube reporter of microRNA hybridization events in vivo. Nature Biomedical Engineering 2017, 1 (4), 0041. 69. Bo, X.; Zhou, M.; Guo, L., Electrochemical sensors and biosensors based on less aggregated graphene. Biosensors and Bioelectronics 2017, 89, 167-186. 70. Fan, X.; Qi, Y.; Shi, Z.; Lv, Y.; Guo, Y., Molecular mechanism of helicase on graphene-based hybridization reaction platform for microRNA detection. RSC Advances 2017, 7 (58), 36444-36449. 71. Fan, X.; Qi, Y.; Shi, Z.; Lv, Y.; Guo, Y., A graphene-based biosensor for detecting microRNA with augmented sensitivity through helicase-assisted signal amplification of hybridization chain reaction. Sensors and Actuators B: Chemical 2018, 255, 1582- 1586. 72. Gao, Y.-P.; Wu, X.; Huang, K.-J.; Xing, L.-L.; Zhang, Y.-Y.; Liu, L., Two-dimensional transition metal diseleniums for energy storage application: a review of recent developments. CrystEngComm 2017, 19 (3), 404-418. 73. Zhu, X.; Shen, Y.; Cao, J.; Yin, L.; Ban, F.; Shu, Y.; Li, G., Detection of microRNA SNPs with ultrahigh specificity by using reduced graphene oxide-assisted rolling circle amplification. Chemical Communications 2015, 51 (49), 10002-10005.

142

74. Miao, P.; Tang, Y.; Wang, B.; Meng, F., Near-infrared Ag2S quantum dots-based DNA logic gate platform for miRNA diagnostics. Analytical Chemistry 2016, 88 (15), 7567- 7573. 75. Qiu, X.; Hildebrandt, N., Rapid and multiplexed microRNA diagnostic assay using quantum dot-based forster resonance energy transfer. ACS Nano 2015, 9 (8), 8449- 8457. 76. Hu, T.; Zhang, L.; Wen, W.; Zhang, X.; Wang, S., Enzyme catalytic amplification of miRNA-155 detection with graphene quantum dot-based electrochemical biosensor. Biosensors and Bioelectronics 2016, 77, 451-456. 77. Deng, H.; Liu, Q.; Wang, X.; Huang, R.; Liu, H.; Lin, Q.; Zhou, X.; Xing, D., Quantum dots-labeled strip biosensor for rapid and sensitive detection of microRNA based on target-recycled nonenzymatic amplification strategy. Biosensors and Bioelectronics 2017, 87, 931-940. 78. Yadav, S.; Masud, M. K.; Islam, M. N.; Gopalan, V.; Lam, A. K.-y.; Tanaka, S.; Nguyen, N.-T.; Al Hossain, M. S.; Li, C.; Yamauchi, M. Y., Gold-loaded nanoporous iron oxide nanocubes: a novel dispersible capture agent for tumor-associated autoantibody analysis in serum. Nanoscale 2017, 9 (25), 8805-8814. 79. Wei, H.; Wang, E., Nanomaterials with enzyme-like characteristics (nanozymes): next- generation artificial enzymes. Chemical Society Reviews 2013, 42 (14), 6060-6093. 80. Masud, M. K.; Yadav, S.; Islam, M. N.; Nguyen, N.-T.; Salomon, C.; Kline, R.; Alamri, H. R.; Alothman, Z. A.; Yamauchi, Y.; Hossain, M. S. A., Gold-loaded nanoporous ferric oxide nanocubes with peroxidase-mimicking activity for electrocatalytic and colorimetric detection of autoantibody. Analytical Chemistry 2017, 89 (20), 11005- 11013. 81. Gorgannezhad, L.; Umer, M.; Kamal Masud, M.; Hossain, M. S. A.; Tanaka, S.; Yamauchi, Y.; Salomon, C.; Kline, R.; Nguyen, N. T.; Shiddiky, M. J., Detection of FGFR2: FAM76A Fusion Gene in Circulating Tumor RNA Based on Catalytic Signal Amplification of Graphene Oxide‐loaded Magnetic Nanoparticles. Electroanalysis 2018, 30 (10), 2293-2301. 82. Qiu, G.-H.; Lu, W.-Z.; Hu, P.-P.; Jiang, Z.-H.; Bai, L.-P.; Wang, T.-R.; Li, M.-M.; Chen, J.-X., A metal-organic framework based PCR-free biosensor for the detection of gastric cancer associated microRNAs. Journal of Inorganic Biochemistry 2017, 177, 138-142.

143

83. Wu, Y.; Han, J.; Xue, P.; Xu, R.; Kang, Y., Nano metal–organic framework (NMOF)- based strategies for multiplexed microRNA detection in solution and living cancer cells. Nanoscale 2015, 7 (5), 1753-1759. 84. Chen, M.; Gan, N.; Zhou, Y.; Li, T.; Xu, Q.; Cao, Y.; Chen, Y., A novel aptamer-metal ions-nanoscale MOF based electrochemical biocodes for multiple antibiotics detection and signal amplification. Sensors and Actuators B: Chemical 2017, 242, 1201-1209. 85. Shao, H.; Lu, J.; Zhang, Q.; Hu, Y.; Wang, S.; Guo, Z., Ruthenium-based metal organic framework (Ru-MOF)-derived novel Faraday-cage electrochemiluminescence biosensor for ultrasensitive detection of miRNA-141. Sensors and Actuators B: Chemical 2018, 268, 39-46. 86. Li, Y.; Yu, C.; Yang, B.; Liu, Z.; Xia, P.; Wang, Q., Target-catalyzed hairpin assembly and metal-organic frameworks mediated nonenzymatic co-reaction for multiple signal amplification detection of miR-122 in human serum. Biosensors and Bioelectronics 2018, 102, 307-315. 87. Wang, J.-S.; Jin, F.-Z.; Ma, H.-C.; Li, X.-B.; Liu, M.-Y.; Kan, J.-L.; Chen, G.-J.; Dong, Y.-B., Au@ Cu (II)-MOF: Highly efficient bifunctional heterogeneous catalyst for successive oxidation–condensation reactions. Inorganic Chemistry 2016, 55 (13), 6685-6691. 88. Wang, H.; Jian, Y.; Kong, Q.; Liu, H.; Lan, F.; Liang, L.; Ge, S.; Yu, J., Ultrasensitive electrochemical paper-based biosensor for microRNA via strand displacement reaction and metal-organic frameworks. Sensors and Actuators B: Chemical 2018, 257, 561- 569. 89. Jamali, A. A.; Pourhassan-Moghaddam, M.; Dolatabadi, J. E. N.; Omidi, Y., Nanomaterials on the road to microRNA detection with optical and electrochemical nanobiosensors. TrAC Trends in Analytical Chemistry 2014, 55, 24-42. 90. Labib, M.; Berezovski, M. V., Electrochemical sensing of microRNAs: avenues and paradigms. Biosensors and Bioelectronics 2015, 68, 83-94. 91. Kilic, T.; Erdem, A.; Ozsoz, M.; Carrara, S., microRNA biosensors: opportunities and challenges among conventional and commercially available techniques. Biosensors and Bioelectronics 2018, 99, 525-546. 92. Yildiz, I., Applications of magnetic nanoparticles in biomedical separation and purification. Nanotechnology Reviews 2016, 5 (3), 331-340. 93. Koo, K. M.; Carrascosa, L. G.; Trau, M., DNA-directed assembly of copper nanoblocks with inbuilt fluorescent and electrochemical properties: application in simultaneous 144

amplification-free analysis of multiple RNA species. Nano Research 2018, 11 (2), 940- 952. 94. Li, Y.; Tian, R.; Zheng, X.; Huang, R., Amplified electrochemical detection of nucleic acid hybridization via selective preconcentration of unmodified gold nanoparticles. Analytica Chimica Acta 2016, 934, 59-65. 95. Lu, Z.; Tang, H.; Wu, D.; Xia, Y.; Wu, M.; Yi, X.; Li, H.; Wang, J., Amplified voltammetric detection of miRNA from serum samples of glioma patients via combination of conducting magnetic microbeads and ferrocene-capped gold nanoparticle/streptavidin conjugates. Biosensors and Bioelectronics 2016, 86, 502- 507. 96. Wang, Y.-H.; Huang, K.-J.; Wu, X.; Ma, Y.-Y.; Song, D.-L.; Du, C.-Y.; Chang, S.-H., Ultrasensitive supersandwich-type biosensor for enzyme-free amplified microRNA detection based on N-doped graphene/Au nanoparticles and hemin/G-quadruplexes. Journal of Materials Chemistry B 2018, 6 (14), 2134-2142. 97. Cheng, F.-F.; He, T.-T.; Miao, H.-T.; Shi, J.-J.; Jiang, L.-P.; Zhu, J.-J., Electron transfer mediated electrochemical biosensor for microRNAs detection based on metal ion functionalized titanium phosphate nanospheres at attomole level. ACS Applied Materials & Interfaces 2015, 7 (4), 2979-2985. 98. Su, S.; Wu, Y.; Zhu, D.; Chao, J.; Liu, X.; Wan, Y.; Su, Y.; Zuo, X.; Fan, C.; Wang, L., On‐Electrode Synthesis of Shape‐Controlled Hierarchical Flower‐Like Gold Nanostructures for Efficient Interfacial DNA Assembly and Sensitive Electrochemical Sensing of MicroRNA. Small 2016, 12 (28), 3794-3801. 99. Su, S.; Cao, W.; Liu, W.; Lu, Z.; Zhu, D.; Chao, J.; Weng, L.; Wang, L.; Fan, C.; Wang, L., Dual-mode electrochemical analysis of microRNA-21 using gold nanoparticle-

decorated MoS2 nanosheet. Biosensors and Bioelectronics 2017, 94, 552-559. 100. Tian, Q.; Wang, Y.; Deng, R.; Lin, L.; Liu, Y.; Li, J., Carbon nanotube enhanced label-free detection of microRNAs based on hairpin probe triggered solid-phase rolling-circle amplification. Nanoscale 2015, 7 (3), 987-993. 101. Ye, Y.; Gao, J.; Zhuang, H.; Zheng, H.; Sun, H.; Ye, Y.; Xu, X.; Cao, X., Electrochemical gene sensor based on a glassy carbon electrode modified with hemin- functionalized reduced graphene oxide and gold nanoparticle-immobilized probe DNA. Microchimica Acta 2017, 184 (1), 245-252.

145

102. Yu, N.; Wang, Z.; Wang, C.; Han, J.; Bu, H., Combining padlock exponential rolling circle amplification with CoFe2O4 magnetic nanoparticles for microRNA detection by nanoelectrocatalysis without a substrate. Analytica Chimica Acta 2017, 962, 24-31. 103. Yuan, Y.-H.; Wu, Y.-D.; Chi, B.-Z.; Wen, S.-H.; Liang, R.-P.; Qiu, J.-D., Simultaneously electrochemical detection of microRNAs based on multifunctional magnetic nanoparticles probe coupling with hybridization chain reaction. Biosensors and Bioelectronics 2017, 97, 325-331. 104. Zhu, D.; Liu, W.; Zhao, D.; Hao, Q.; Li, J.; Huang, J.; Shi, J.; Chao, J.; Su, S.; Wang, L., Label-free electrochemical sensing platform for microRNA-21 detection using thionine and gold nanoparticles co-functionalized MoS2 nanosheet. ACS Applied Materials & Interfaces 2017, 9 (41), 35597-35603. 105. Sage, A. T.; Besant, J. D.; Lam, B.; Sargent, E. H.; Kelley, S. O., Ultrasensitive electrochemical biomolecular detection using nanostructured microelectrodes. Accounts of Chemical Research 2014, 47 (8), 2417-2425. 106. Bin, X.; Sargent, E. H.; Kelley, S. O., Nanostructuring of sensors determines the efficiency of biomolecular capture. Analytical Chemistry 2010, 82 (14), 5928-5931. 107. Soleymani, L.; Fang, Z.; Sun, X.; Yang, H.; Taft, B. J.; Sargent, E. H.; Kelley, S. O., Nanostructuring of patterned microelectrodes to enhance the sensitivity of electrochemical nucleic acids detection. Angewandte Chemie International Edition 2009, 48 (45), 8457-8460. 108. Yang, H.; Hui, A.; Pampalakis, G.; Soleymani, L.; Liu, F. F.; Sargent, E. H.; Kelley, S. O., Direct, electronic microRNA detection for the rapid determination of differential expression profiles. Angewandte Chemie International Edition 2009, 48 (45), 8461- 8464. 109. Congur, G.; Eksin, E.; Erdem, A., Impedimetric detection of microRNA at graphene oxide modified sensors. Electrochimica Acta 2015, 172, 20-27. 110. Guo, H.-L.; Wang, X.-F.; Qian, Q.-Y.; Wang, F.-B.; Xia, X.-H., A green approach to the synthesis of graphene nanosheets. ACS Nano 2009, 3 (9), 2653-2659. 111. Eksin, E.; Bikkarolla, S. K.; Erdem, A.; Papakonstantinou, P., Chitosan/nitrogen doped reduced graphene oxide modified biosensor for impedimetric detection of microRNA. Electroanalysis 2018, 30 (3), 551-560. 112. Daneshpour, M.; Karimi, B.; Omidfar, K., Simultaneous detection of gastric cancer- involved miR-106a and let-7a through a dual-signal-marked electrochemical nanobiosensor. Biosensors and Bioelectronics 2018, 109, 197-205. 146

113. Azimzadeh, M.; Rahaie, M.; Nasirizadeh, N.; Ashtari, K.; Naderi-Manesh, H., An electrochemical nanobiosensor for plasma miRNA-155, based on graphene oxide and gold nanorod, for early detection of breast cancer. Biosensors and Bioelectronics 2016, 77, 99-106. 114. Gao, Z.; Yang, Z., Detection of microRNAs using electrocatalytic nanoparticle tags. Analytical Chemistry 2006, 78 (5), 1470-1477. 115. Bo, B.; Zhang, T.; Jiang, Y.; Cui, H.; Miao, P., Triple signal amplification strategy for ultrasensitive determination of miRNA based on duplex specific nuclease and bridge DNA–gold nanoparticles. Analytical Chemistry 2018, 90 (3), 2395-2400. 116. Liu, L.; Chang, Y.; Xia, N.; Peng, P.; Zhang, L.; Jiang, M.; Zhang, J.; Liu, L., Simple, sensitive and label–free electrochemical detection of microRNAs based on the in situ formation of silver nanoparticles aggregates for signal amplification. Biosensors and Bioelectronics 2017, 94, 235-242. 117. Yang, J.; Tang, M.; Diao, W.; Cheng, W.; Zhang, Y.; Yan, Y., Electrochemical strategy for ultrasensitive detection of microRNA based on MNAzyme-mediated rolling circle amplification on a gold electrode. Microchimica Acta 2016, 183 (11), 3061- 3067. 118. Zhou, Y.; Yin, H.; Li, J.; Li, B.; Li, X.; Ai, S.; Zhang, X., Electrochemical biosensor for microRNA detection based on poly (U) polymerase mediated isothermal signal amplification. Biosensors and Bioelectronics 2016, 79, 79-85. 119. Shuai, H.-L.; Huang, K.-J.; Zhang, W.-J.; Cao, X.; Jia, M.-P., Sandwich-type microRNA biosensor based on magnesium oxide nanoflower and graphene oxide–gold nanoparticles hybrids coupling with enzyme signal amplification. Sensors and Actuators B: Chemical 2017, 243, 403-411. 120. Asadzadeh-Firouzabadi, A.; Zare, H. R., Preparation and application of AgNPs/SWCNTs nanohybrid as an electroactive label for sensitive detection of miRNA related to lung cancer. Sensors and Actuators B: Chemical 2018, 260, 824-831.

147

Chapter 3

Nanoarchitectured peroxidase-mimetic nanozymes: mesoporous nanocrystalline α- or γ-iron oxide?*

*Sections of this chapter are based on Masud, M.K., et al., Journal of Materials Chemistry B, 2019, 7, 5412-5422.

148

3. Nanoarchitectured peroxidase-mimetic nanozymes: mesoporous nanocrystalline α- or γ-iron oxide?

3.1. Introduction Inorganic nanoparticles (NPs) with an enzyme-mimicking ability (nanozymes) have gained significant attention in the fields of analytical sensing, biosensing, and nanomedicine.1-2 Nano- zymes exhibit several advantages over natural enzymes due to their high stability against protease digestion or denaturation, structural, morphological, and functional integrity under diﬀerent environmental stress (low pH, high temperature or presence of salt), inexpensive preparation and wide application as an electrocatalyst.3-4 Similar to horseradish peroxidase (HRP), nanozymes can catalyze the oxidation of chromogenic substances in the presence of hydrogen peroxide (H2O2) to generate colourimetric (colour change) and electrochemical (current) signals. In recent years, various nanozymes with peroxidase-like activity (e.g., 5 peroxidase, oxidase, and catalase) have been employed for: (a) determination of H2O2, (b) glucose detection in biological (blood and urine) samples,6 (c) evaluation of antioxidants,7 and (d) nanozyme-based colourimetric and electrochemical immunosensors3 and nucleic acid sensors.8 Apart from high nanozyme activity, easy separation of NPs or NPs-attached target biomolecules (target biomolecules can be fastened by functionalizing NPs with an antibody or nucleic acid probe relevant to the target) from complex biological systems using an external magnet and re-using them is a highly desirable feature for a biosensor.9 In the past decade, a plethora of NPs and composites have shown enzyme-mimicking properties, and they have been utilized in biosensors for the detection of specific biomolecules.10 Among them, iron oxide (IO) nanostructures, owing to their superparamagnetism, high catalytic activity, low toxicity and ease of synthesis, have been widely utilized as nanozymes. However, many previously reported iron oxides demonstrated their activity at a relatively high temperature (32-45 oC), which limited their applications in biosensors since biomolecules tend to degrade or be denatured at elevated temperatures.11 Recently, several nanozymes have been reported to exhibit mimetic activity at room temperature; however, they required cumbersome modifications with biomolecules (e.g., ATP, AMP) or polymer-based engineering (e.g. molecular imprinting).12- 16 Therefore, the preparation of nanozymes with high catalytic activity at room temperature (without any functionalization or modification) is highly desirable. Mesoporous iron oxide with high porosity and surface area, narrow pore size distribution, and controllable wall composition may potentially achieve enhanced peroxidase mimetic activity at room temperature. The high electrocatalytic and enzymatic performance of mesoporous iron oxide in terms of turnover rate

149

and selectivity are contributed by its high active surface area, exposed Fe atoms and enhanced electronic effect.3-4 Moreover, the highly interconnected pores of mesoporous iron oxide could lead to higher fluxes of diffusing redox species over the entire area. Iron(III) oxide (Fe2O3) possesses

Iron(III) oxide (Fe2O3) possesses four crystal phases: alpha- (α-), beta- (β-), gamma- 17 (γ-), and epsilon-Fe2O3 (ε-Fe2O3). Among all these phases, α-Fe2O3 (hematite) and γ-Fe2O3 (maghemite) are the most common and widely investigated phases of iron(III) oxide.15 The stability and unique magnetism of α- Fe2O3 and γ-Fe2O3 render them popular as nanozymes in biosensing platforms. Nevertheless, their intrinsic properties and functionalities largely depend 18-19 on their morphology, size, and crystal structure. The α-Fe2O3 phase possesses a rhombohedral-centred hexagonal structure with a close-packed oxygen lattice,20-21 while γ- 22-23 Fe2O3 has a cubic crystal structure of an inverse spinel-type lattice structure. Owing to their diﬀerent crystal structures, they typically exhibit diﬀerent activity toward the peroxidase substrate. To date, most of the reported iron oxide nanozymes have either been α-Fe2O3 or γ-

Fe2O3 or their composite. Although admirable research and results in the field of IO-based nanozymes have been achieved, few reports have investigated the relationship between their nanozyme activity and crystal phases. To the best of our knowledge, a systematic comparison of the catalytic activity between a- and γ- Fe2O3 nanozymes towards chromogenic substances has been rarely reported. The detailed mechanism underlying the structural eﬀect of the crystal phases (mesoporous γ-Fe2O3 and α-Fe2O3) toward the nanozyme activity will be useful for designing artificial nanozymes for future biosensing applications.

In this work, mesoporous IOs with two crystal phases (γ-Fe2O3 and a-Fe2O3) have been synthesized by a hard-templating approach using mesoporous carbon (CMK-3). The intrinsic nanozyme (peroxidase-mimetic) activity of these two phases toward the oxidation of TMB (in the presence of H2O2) was tested at room temperature and used to determine the crystal phase which could give superior activity. To achieve room temperature nanozyme activity, a porous structure is integrated into the surface of both crystal phases. Finally, the as-prepared mesoporous IO samples were employed for both naked eye and colourimetric detection of glucose. Our mesoporous γ- Fe2O3 sample shows ultrahigh sensitivity toward the detection of glucose with an impressive limit of detection (LOD) of 0.9 mM, indicating its promising potential for nanozyme-based sensing of glucose and other biomolecules.

150

3.2. Experimental 3.2.1. Materials Iron(III) nitrate nonahydrate, iron(III) oxide nanopowder, glycerol (C3H8O3, 99.5%), isopropanol, sodium acetate, hydrogen peroxide, 2,2'-azino-bis (ABTS), terephthalic acid and absolute ethanol were purchased from Sigma-Aldrich, Australia. 3,3’,5,5’- Tetramethylbenzidine, was purchased from Sigma Life Science Australia. Dimethyl Sulfoxide AR (DMSO) was purchased from Chem-Supply, Australia. All the chemicals were used as received without further purification.

3.2.2. Characterization All the samples were systematically characterized by various analytical, spectroscopic and imaging techniques. Powder X-ray diffraction (XRD) patterns were obtained by using a Bruker D8 Focus Advance Diffractometer equipped with Lynx Eye detector with a Ni-filtered Cu Kα radiation (λ = 1.5406 Å) operating at 40 kV and 40 mA in 2θ range of 1.0 to 7.0 with a step size of 0.01o and a step time of 2 s. The diffraction patterns were recorded with a divergent slit of 0.298° over the 2 θ range of 1.5 to 7.5° and 5-50° with step size = 0.01° and a step time of 2 s. The lattice parameters were refined by the Rietveld method using the TOPAS program. Transition electron microscopy (TEM) images and selected area electron diffraction (SAED) patterns were obtained by using a 2100 JEOL microscope operated at 120 keV and 200 keV. Prior to the analysis, the samples were ultrasonicated in absolute ethanol for 20 min and then dropped on 200-mesh lacey foam coated copper grids and dried at room temperature before loading into the microscope for analysis. Nitrogen adsorption-desorption isotherms were measured at 77 K using Micromeritics ASAP 2020 surface area analyzer. The samples were initially outgassed at 300 °C for 4 h before the measurements. Helium was used to carry out free space measurements. The specific surface area was calculated by using the BET (Brunauer-Emmett-Teller) equation, micro- and mesopore-size distribution was obtained from the desorption branch of the isotherm by HK (Horvath-Kawazoe) and BJH (Barrett-Joyner- Halenda) methods, respectively. Diffuse-reflectance ultraviolet and visible spectra of the materials were recorded by using Thermo-Scientific Evolution 600 spectrophotometer (190 nm - 800 nm range) with barium sulfate as a standard under ambient conditions. Prior to the analysis, the BaSO4 background spectrum was taken followed by the analysis of the samples. Electron paramagnetic resonance (ESR) of the samples was recorded on a Miniflex-II electron spin resonance spectrometer (X-band). The EPR spectra of all the samples were recorded at room temperature with a field modulation of 100 KHz. Magnetization measurements were 151

carried out at 300 K by employing a Lakeshore 7400 vibrating sample magnetometer (VSM). Temperature dependence of magnetization was measured under zero-field cooling (ZFC) and field cooling (FC) with an applied magnetic field of 1000 Oe and temperature range of 10 K to 300 K.

3.2.3. Synthesis of CMK-3 Mesoporous silica (SBA-15) was synthesized according to the procedure reported in our earlier work with some modifications.24-26 SBA-15 was synthesized to prepare the CMK-3 template.

The synthesis gel with composition TEOS/P123/HCl/H2O = 1.0/0.017/ 5.9/193.0 was hydrothermally treated at 413 K for 24 h to form large pores. The obtained products were filtered, dried at 373 K overnight and calcined at 773 K for 6 h. CMK-3 was synthesized according to the procedure reported in our previous reports.25, 27 In a typical process, 1.0 g of dried SBA-15 powder was placed into an acidic sucrose solution (1.25 g sucrose, 0.14 g H2SO4 o o conc. solution, 5.0 g H2O). The obtained mixture was then air-dried at 100 C and 160 C for a period of 6 h each. This process was repeated once again to form a coloured sample, which o was then carbonized at 900 C under an N2 atmosphere for 7 h. The obtained silica/carbon composite was then etched with a 10–15% HF solution to remove the silica template. The etching solution was washed, filtered, and dried at 120 oC to yield the CMK-3.

3.2.4. Synthesis of α-Fe2O3 and γ-Fe2O3 26 To prepare the mesoporous γ-Fe2O3, 0.3 g of dried CMK-3 powder was dissolved in 20 mL of 0.5 M iron nitrate solution (in ethanol) and then stirred for 6 h. The mixture (solution) was then filtered and dried. The dried powder was further treated with 0.05 M NaOH solution (10 mL) and mixed under magnetic stirring for 3 h, followed by filtration and drying at 373 K. The dried powder was subsequently calcined at 523 K in a quasi-sealed container. To remove the carbon scaﬀold, the obtained sample was heated at 573 K for 2 h under a nitrogen atmosphere and at 623 K for 1 h under an air atmosphere to achieve the mesoporous γ-Fe2O3. The as- obtained mesoporous γ-Fe2O3 was further heated at 623 K in the air to obtain the mesoporous

α-Fe2O3.

3.2.5. Nanozyme activity

The nanozyme activities (peroxidase-like) of both γ-Fe2O3 and α-Fe2O3 were investigated by using 5 µg of NPs to catalyze a TMB substrate solution at room temperature. The substrate solution was prepared by dissolving 800 µM TMB (TMB in DMSO) and 500 mM H2O2 in 80 152

mL of 0.2 M sodium acetate (NaAc) buffer solution (pH 3.5). The reaction was conducted in the dark for 10 min, and the nanozyme catalyzed the oxidation of the colourless TMB solution to a blue-coloured solution. The intensity (absorbance at 652 nm) of the coloured TMB (blue) solution was measured by using a plate reader in time scan mode. To obtain the steady-state kinetics of the nanozyme toward TMB oxidation, different concentrations of TMB (0.01 to 1.0 mM) were employed under typical reaction conditions (5 mg iron oxide, 500 mM H2O2 in 80 mL of 0.2 M NaAc buffer, pH 3.5, room temperature). Similar reactions were also conducted for H2O2 by varying its concentration from 0.01 to 1.0 M. The apparent kinetic parameters for both TMB and H2O2 (Km and Vmax) were estimated by considering the Michaelis–Menten equation28 and using the Lineweaver–Burk equation.29

3.2.6. Investigation of nanozyme activity through scavenging hydroxyl radical (·OH)

The formation of ·OH radicals by the iron oxide nanozyme through the cleavage of H2O2 was monitored by scavenging ·OH radicals using isopropyl alcohol (IPA), methyl alcohol (MA), and terephthalic acid (TA). In a typical procedure, different amounts of IPA and TA were added into the nanozyme-oxidized blue coloured TMB solution (5 µg of iron oxide, 700 µM TMB and 500 mM H2O2 in 0.2 M NaAc, pH 3.5 buffer) and the decrease in the colour intensity was monitored by both naked-eye observation and absorbance spectra. The TA reacts with ·OH radicals, and the formation of 2-hydroxy-terephthalic acid (i.e., scavenging of ·OH radicals) was monitored by the fluorescence spectra obtained for hydroxy-terephthalic acid. In our experiments, different amounts of the iron oxide nanozyme (both γ-Fe2O3 and α-Fe2O3) ranging from 1 to 40 mg were added into a mixture solution consisting of a 0.2 M NaAC (300 mL) buffer solution, 5 mL of H2O2 solution and 20 mL of 0.2 M TA followed by 12 h incubation in the dark. The fluorescence spectra were then measured by using a plate reader at an excitation wavelength of 315 nm.

3.2.7. Naked-eye observation and colourimetric estimation of glucose

To assess the glucose-sensing capabilities of both α-Fe2O3 and γ-Fe2O3 phases, 30 μL of glucose solution with a different concentration ranging from 1.0 µM to 1000 µM was mixed with 3.0 μL of 20 mg/mL GOx solution in PBS (10 mM, pH 7.4) and incubated for 30 minutes at 45 oC. After that, 17 µL of TMB solution (1mg/mL in DMSO) and 5 µL of nanozyme (IOs) (1mg/mL) and 78 µL 0.2 M NaAc buffer (pH 3.5) was added to the mixture and incubated for 10 minutes in dark at room temperature. The glucose responding colour changes from

153

colourless TMB to oxidized blue colour TMB was observed by the naked eye and estimated by using calorimetry.

3.3. Result and Discussion

3.3.1. Structural Characterization of the Nanozymes

The synthesis of the ordered mesoporous IO samples (γ-Fe2O3 and α-Fe2O3) was achieved by using the nanocasting strategy with CMK-3 as the hard-template (the detailed experimental procedure is given in the Experimental section). The high structural stability and inter-pore connectivity of the starting hard template are critical for ensuring the successful replication of the ordered mesoporosity of the CMK-3 template in the final mesoporous metal oxide product. Therefore, we have synthesized an ordered mesoporous carbon, CMK-3, with thicker walls (d = 10.8 nm), as shown in Table 1.

Table 3.1. Structural information of SBA-15, CMK-3, and the resulting mesoporous γ-Fe2O3 and α-Fe2O3.

a b 2 −1 c d 3 −1 e Material d (nm) L (nm) SBET (m g ) DBJH (nm) Vp (cm g )

XRD TEM

SBA-15 10.8 ------480 8.7 1.29

CMK-3 9.8 ------1141 2.1 1.35

-Fe2O3 12.3 10.2 9.2 125 8.5 0.20

α-Fe2O3 11.7 18.8 --- 144 8.7 0.25 a Mesostructural periodicity; b Average crystallite size; c BET surface area; d Average pore size by the BJH method; e Total pore volume

154

Figure 3.1: (a) Low- and wide-angle XRD patterns of mesoporous γ-Fe2O3 and α-Fe2O3. (b)

N2 adsorption-desorption isotherms and pore size distribution curves of mesoporous γ-Fe2O3 and α-Fe2O3 (c) TEM image with ED patterns and (d) high-resolution TEM image of mesoporous γ-Fe2O3.

Figure 3.1a presents the wide-angle XRD diﬀraction patterns of the obtained mesoporous α-Fe2O3 and γ-Fe2O3, which show the reflection characteristics of the trigonal

(R3c) α-Fe2O3 (ICDD No. 33-0664) and cubic γ-Fe2O3 (P4132) (ICDD No. 39-1346) crystal systems, respectively. The line broadening of the reflections can be attributed to the nanocrystalline nature of the pore wall. The SEM and TEM images of both γ-Fe2O3 and α-

Fe2O3 are depicted in Figure 3.2.

155

Figure 3.2: (a, b) SEM image of (a) mesoporous γ-Fe2O3 and (b) mesoporous α-Fe2O3. (c and d) Enlarged TEM images of mesoporous γ-Fe2O3.

The XRD patterns were refined by the Rietveld method. Figure 3.3 depicts the Rietveld refinements of the XRD patterns of mesoporous α- Fe2O3 and γ-Fe2O3, respectively. The low- angle XRD patterns of both samples show a broad and less resolved reflection at around 0.81 (Figure 3.1a; inset), indicating the disordered nature of the mesopores (Table 1). This disorder may be attributed to the repeated synthetic steps, which would collapse the original mesostructure of CMK-3, and random growth and orientation of the IO crystals during the nanocasting process. Furthermore, we analyzed the carbon content. The α-Fe2O3 and γ-Fe2O3 samples possess 0.67% and 1.71%, respectively.

156

Figure 3.3: Rietveld refined XRD patterns of (a) mesoporous γ-Fe2O3 and (b) mesoporous α-

Fe2O3. The calculated lattice parameters are a, b, c = 8.374(7) Å (for mesoporous γ-Fe2O3) and a, b = 5.0327(9) Å; c = 13.761(3) Å (for mesoporous α-Fe2O3), respectively.

Figure 3.1b depicts the N2 adsorption-desorption isotherms of mesoporous α-Fe2O3 and

γ-Fe2O3, showing type IV isotherms with small hysteresis loops, which are characteristic of mesoporous materials. This is further supported by the broad pore size distribution (Figure 3.1b; inset), which clearly indicates the formation of disordered mesostructure. The textural properties of all the samples, including the starting templates (SBA-15 and CMK-3), are summarised in Table 3.1. The TEM images and selected area electron diffraction (ED) patterns of the γ-Fe2O3 are shown in Figure 3.1c and d, revealing that the resulting γ-Fe2O3 exhibits wormhole-like mesopores (the enlarged images are given in Figure 3.2c and d). Furthermore, the pore walls of the mesoporous γ-Fe2O3 are crystalline, which is also evident from the lattice fringes and ring-like ED patterns. The crystallite size (L) calculated from TEM is in good agreement with the value obtained by XRD data (Table 3.1). Magnetic NPs are highly useful for magnet-based pre-concentration, isolation, separation, and purification of target analytes from complex biological systems. Because of their small size and volume, magnetic NPs are mostly superparamagnetic, i.e. without the presence of an external magnetic field, the NPs possess no net magnetic moment; whereas, in the presence of an external magnetic field, a magnetic dipole is instigated, and a net arrangement of magnetic moments occur. The magnetic moment of NPs returns to its native random orientation when the external magnetic field is removed, and in turn, they become nonmagnetic. This characteristic is a prerequisite for magnetic manipulation of the sample.

157

Figure 3.4a and b display the magnetization isotherms of mesoporous γ-Fe2O3 and bulk-

γ-Fe2O3 (prepared without the CMK-3 template) measured at various temperatures. It is well established that their domain structures greatly influence the magnetic properties of IOs along with their size and shape.30 The reduction of the crystal size to nano dimensions can result in competition between thermal energy and magnetic anisotropy, which aﬀects the magnetic behaviour of the materials. In particular, this feature is observed in the case of γ-Fe2O3, which results in the transition from a ferromagnetic state to a superparamagnetic state by decreasing 31 the crystal size. The magnetic saturation of mesoporous γ-Fe2O3 sample (Figure 3.4a) is lower than that of the bulk-γ-Fe2O3 sample (Figure 3.4b). Besides, the S-shaped hysteresis loops with low values of coercivity (HC) and remanent magnetization (MR) along with a high value of saturation magnetization (MS) clearly indicate that the mesoporous γ-Fe2O3 exhibits a superparamagnetic behaviour at room temperature in contrast to the ferromagnetic bulk γ-

Fe2O3. This is because the magnetic moment of each nanocrystallite structure is too small to be aﬀected by thermal excitation. The HC observed in ferromagnetic bulk γ-Fe2O3 is due to the weak interaction between neighbouring nanocrystallites.

Figure 3.4: M-H curves of (a1) of mesoporous γ -Fe2O3, (a2) bulk-γ-Fe2O3, (b1) mesoporous

α-Fe2O3, and (b2) bulk-α-Fe2O3.

158

This is further evident from the ZFC curve (Figure 3.5), which shows a broad cusp at ~85 K, owing to the blocking transition temperature, which is a typical characteristic of 32 superparamagnetic behaviour and is not evident in the M–T curves of bulk γ-Fe2O3. The temperature dependence of the M–H curves is indicated by the increase in the broadness of the hysteresis loop at low temperatures of 20 K and 70 K. In contrast to the bulk sample, the M–H curves of mesoporous α-Fe2O3 do not show antiferromagnetic ordering even at low temperatures of 20 K and 70 K, which clearly indicates the suppression of Morin-transitions owing to its smaller crystallite size (Figure 3.4c and d).33

Figure 3.5: M-T curves of (a) mesoporous γ-Fe2O3 and (b) bulk γ-Fe2O3.

3.3.2. Nanozyme activity

Similar to HRP, IO NPs can catalyze the oxidation reaction of TMB in the presence of H2O2 at pH ranging from 3.0-6.5 or in an acidic buffer.34 This catalytic process converts the colourless TMB into a blue-coloured TMB (a charged transfer diamine complex), which gives a maximum absorbance at 652 nm. Upon the addition of an acid (or stop solution), the blue- coloured solution changes to a yellow-coloured solution with a maximum absorbance at 450 nm. This nanozyme-based reaction has widely been used to develop a large number of biosensors for the detection of H2O2, glucose, cellular biomarkers, proteins, bacteria, and clinically-relevant biomolecules and biomarkers (exosomes, DNA, RNA, autoantibodies, etc.).3-4, 8 Herein, we have intricately studied the nanozyme activity of two different crystal phases of mesoporous IO (mesoporous γ-Fe2O3 and α-Fe2O3). To examine the nanozyme activity of both mesoporous IO samples, we have performed a set of experiments in the presence and absence (referred to as a control) of the mesoporous iron oxide samples (Figure 3.6a). Both samples were used to catalyze the TMB substrate solution by incubating for 10 min in the dark at room temperature. In comparison with the two sets of controls (without H2O2 and

159

IOs), both mesoporous IO samples generate a clear blue-coloured solution, while the substrate solutions for the two controls remain colourless. As seen in Figure 3.6b, both crystal phases show much higher absorbance (at 652 nm) than the two control samples. Surprisingly, the mesoporous γ-Fe2O3 sample generates a considerably higher absorbance compared to the control sample by almost 48-fold (0.059 vs. 2.941) (Figure 3.6b).

Figure 3.6: (a) Schematic illustration of the nanozyme activity of mesoporous IO for the oxidation of TMB in the presence of H2O2. (b) The peroxidase-like activities of the two mesoporous IO samples in comparison with the two control samples. Experiments were carried out by using 5 µg of mesoporous iron oxide in a reaction volume of 0.2 ml, in 0.2 M NaAc o buffer, pH 3.5 at 25 C. The H2O2 concentration was 500 mM, and the TMB concentration was 700 µM unless otherwise stated. The maximum point was set at 100%. (c) Time-dependent peroxidase-like activity of mesoporous α-Fe2O3 and γ-Fe2O3. The probable mechanism for the IO (ferric oxide)-induced peroxidase mimetic activity is the presence of a large number of ferric ions (Fe3+) on the surface of the mesoporous IO, which initiates the oxidation of TMB by generating hydroxyl free radicals (·OH) from H2O2 following the Fenton reaction, as shown in equation (1)-(4).3, 35 The Fenton reaction is considered to follow two cascade steps, where initially H2O2 attaches to the exposed surface of the nanozyme and the ·OH free radical is generated in the second step. The rate constant for

160

equation (1) (the attachment of H2O2 on the ferric ion surface) is much lower than that of step (3) (generation of the ·OH radical) (0.002 L vs. 76 L mol-1s-1), presenting equation (1) as the rate-limiting step.16 Therefore, the nanozyme activity predominantly depends on the exposed area, i.e., the adsorption sites present on the nanozyme surface. 3+ 2+ + Fe + H2O2 → FeOOH + H (i)

2+ → 2+ FeOOH Fe + HO2· (ii)

2+ 3+ - Fe + H2O2 → Fe + OH + ·OH (iii)

·OH + TMB (colourless) → TMBox (blue) (iv)

Notably, the high peroxidase mimetic activity of the as-prepared IO samples at room temperature may be attributed to their mesoporous structure. The mesopores can improve the catalytic activity by increasing the adsorption of substrates (both H2O2 and TMB). Also, the position of ferric ions and the presence of holes in the crystal of the IOs play essential roles in enhancing their catalytic activity for the Fenton reaction. As seen in Figure 3.6c, the rate of increase in absorbance is much higher for γ-Fe2O3 than that for a-Fe2O3. Astoundingly, the mesoporous γ-Fe2O3 sample exhibits around a seven-times higher response (2.941 vs. 0.407) than that of mesoporous α-Fe2O3. This may be ascribed to the fact that the crystal structure of

γ-Fe2O3 possesses cation vacancies (in γ-Fe2O3, all the irons exist as trivalent and the presence of cation vacancies gives rise to charge neutrality) at the octahedral position.36 In brief, the crystal structure γ-Fe2O3 is generally obtained by creating one-third vacancies out of the 8a (tetrahedral sites; A-sites) and 16d (octahedral sites; B-sites) Fe sites in the Fd3m space group.37 The crystallographic formula of γ-Fe2O3 can be represented as [Fe]8a [Fe5/3◘1/3]16d

O4 where ◘ is a symbol that represents the vacancies. Furthermore, Oosterhout and Rooijmans studies revealed that the spinel tetragonal superstructure of γ-Fe2O3 possessed an ordered distribution of the vacancies with tetragonal symmetry and a three- times doubling along the c-axis, where the Fe atoms are completely ordered.37-38 These characteristic cation vacancies and the ordered exposure of Fe atoms present on the γ-Fe2O3 surface can facilitate increased adsorption of H2O2, triggering the rate-limiting step (equation 1). However, these cation vacancies are not present on the surface of α-Fe2O3, making it less attractive towards the substrate H2O2. This excellent phenomenon results in the relatively weak H2O2 attachment, as well as sevenfold lower activity of α-Fe2O3, compared to that of γ-Fe2O3. Moreover, we have also compared the nanozyme activity (towards TMB) of

161

the mesoporous γ-Fe2O3 with the non-porous γ-Fe2O3 samples, and it has been found that the mesoporous IO gives almost five- times (% of activity; 100 vs. 21.35) higher activity than that of the non-porous IO sample (Figure 3.7), which is due to the pore-induced higher surface area and exposed Fe atoms.

Figure 3.7: The nanozyme activity of non-porous and mesoporous γ-Fe2O3 towards the oxidation of TMB (5 µg NPs, 700 μM TMB, 500 mM H2O2 in 0.2 M NaAc buffer; pH 3.5). Error bars represent the standard error derived from three repeated measurements.

To check further, we have employed another chromogenic substrate 2,2´-azino-bis(3- ethylbenzothiazoline-6-sulphonic acid) (ABTS) to assess the nanozyme activity of both crystal phases. Both nanozymes catalyze the oxidation of ABTS and generate a green- coloured solution with a characteristic absorbance at 405 nm.39 Similar to the case of TMB, the mesoporous γ-Fe2O3 sample exhibits around 4 times higher absorbance (0.4658 vs. 1.5587 at

405 nm) than the mesoporous α-Fe2O3 sample (Figure 3.8)

Figure 3.8: (a) The absorbance spectra obtained for the oxidation of ABTS by peroxidase mimetic activity of iron oxide nanozymes where the mesoporous γ-Fe2O3 exhibits a much higher nanozyme activity toward ABTS oxidation than the mesoporous α-Fe2O3. (b) The

162

corresponding bar diagram for absorbance at 405 nm; inset shows the color intensity generated by both samples with respect to the control sample (no iron oxide).

Figure 3.9: The responses of absorbance change (at 652 nm) after the addition of hydroxide radical scavengers; (a) isopropanol (IPA) and methyl alcohol (MA) for both mesoporous γ-

Fe2O3 and (b) mesoporous α-Fe2O3 (TMBOX: Scavenger = 1: 0.25) (b); insets show the corresponding images of the colour intensity reduction after reacting with scavengers. Figure c and d depicts the absorbance responses after the addition of different amount both IPA and

MA scavengers towards the mimetic activity of mesoporous γ-Fe2O3 respectively. Error bars represent the standard error derived from three repeated measurements. To investigate the mechanism involved in the nanozyme activity toward the oxidation of TMB, hydroxyl radical scavenging experiments were performed. In our experiments, IPA and MA were added to the nanozyme-oxidized TMB solution, and the decrease of the colour intensity and absorbance were measured. IPA and MA have been considered as scavengers of hydroxyl radicals, and hence they reduce the number of hydroxyl radicals present in the system.40 As seen in Figure 3.9, IPA and MA reduce the colour intensity (fading of the blue colour compared to the control) and absorbance (at 652 nm) of both mesoporous γ-Fe2O3 and

α-Fe2O3, demonstrating the TMB oxidation progresses through the formation of ·OH radicals. With an increasing amount of IPA and MA, the peroxidase mimetic activity of mesoporous γ-

163

Fe2O3 decreased, where an equivalent amount (% v/v) of IPA and MA decreases the activity to around 78% and 68% respectively, revealing that the scavengers absorb ·OH radicals and hence lessen the absorbance (at 652 nm). To gain deeper insights into the participation of the hydroxyl radical, another set of experiments were conducted, in which TA was employed to react with the hydroxyl radical generated by the nanozyme activity of the iron oxides. TA reacts with ·OH radicals and creates a highly fluorescent compound, 2-hydroxy-terephthalic acid, which exhibits emission at 430 nm at an excitation wavelength of 315 nm.41

Figure 3.10: Investigating hydroxyl ion (·OH) radical formation through the fluorescence spectra of 2-hydroxyterephthalic acid produced from the oxidation of terephthalic acid (TA) by ·OH in the presence of both (a) mesoporous γ-Fe2O3 and (b) mesoporous α-Fe2O3.

As seen in Figure 3.10, without the nanozyme, the control sample (0 mg) does not generate any fluorescence signal, whereas the sample containing the nanozyme generates fluorescence signals. With increasing nanozyme amount (from 1 to 40 µg), the amount of generated ·OH radicals increases, which results in the higher fluorescence signals of 2-hydroxy terephthalic acid, representing the proportional relationship between the amount of nanozyme and the quantity of ·OH radicals generated. These results evidently demonstrated and confirmed that both iron oxide nanozymes cleaved the H2O2 to generate ·OH radicals, which subsequently promotes the oxidation of TMB (or ABTS). It is noteworthy that the mesoporous

γ-Fe2O3 gives a much more intense fluorescence spectrum (Figure 3.9a) than the mesoporous

α-Fe2O3 (Figure 3.9b). Similar to the natural enzyme HRP, the nanozyme activity of IO is dependent on the substrate pH, and the amount of IO used. To study the eﬀect of pH, we have studied the nanozyme activity of both mesoporous IO samples by using solutions with diﬀerent pH ranging from 2.5 to 5.5. With increasing pH value, the nanozyme activity of both mesoporous IO

164

samples decreases (Figure 3.11). This may be due to the excessive generation of FeOOH2+ and 3 HO2·, which convert to molecular oxygen, thereby reducing the nanozyme activity. Although pH 2.5 gives the highest activity among all the pH values, under acidic conditions (pH less than 3.0), leaching of iron from the mesoporous IO samples may occur. Therefore, pH 3.5 was chosen as the optimal pH for this study. The amount of nanozyme (i.e. mesoporous IO) can also aﬀect the activity. As shown in Figure 3.12, the nanozyme activity increases with an increasing amount of IO added into the reaction. As substrate solutions containing 5, 10, 15 and 20 µg of IO give nearly similar responses (absorbance), 5 µg was selected as the optimum amount for subsequent experiments.

Figure 3.11: The pH-dependent peroxidase-like activities of (a) mesoporous γ-Fe2O3 and (b) o α-Fe2O3 samples (5 µg). Mean values of absorbance at designated pH of NaAc (at 25 C) buffer solution and the concentrations of TMB and H2O2 were 700 μM and 500 mM, respectively. Error bars represent the standard error derived from three repeated measurements.

Figure 3.12: The peroxidase-like activities of (a) mesoporous γ-Fe2O3 and (b) mesoporous α-

Fe2O3 samples with different loading amounts of the samples. Mean values of absorbance obtained with different amount of mesoporous iron oxide (0 to 20 µg) in NaAc (at 25 oC) buffer solution with the concentrations of TMB and H2O2 being kept at 700 μM and 500 mM, respectively. Error bars represent the standard error derived from three repeated measurements.

To further explore the peroxidase mimetic activity of both mesoporous γ-Fe2O3 and α-

Fe2O3, the apparent steady-state kinetics toward TMB and H2O2 oxidation were ascertained by

165

3 varying the concentrations of TMB and H2O2 by using the initial rate method. Similar to HRP, a regular Michaelis-Menten curve was obtained for both mesoporous γ-Fe2O3 and α-Fe2O3 within the examined concentration ranges for both TMB (Figure 3.13a and c) and H2O2 (Figure

3.13b and d). The catalytic parameters Km (known as the Michaelis-Menten constant) and Vmax were evaluated from the typical Lineweaver–Burk double- reciprocal plot (1/[V0] vs. 1/[S]) 28-29 (Figure 3.12, inset). The Km value signifies the affinity of the enzyme (nanozyme) toward the substrate. A lower Km value implies a higher affinity of the nanozyme for its substrate and vice versa. The estimated Km for mesoporous γ-Fe2O3 toward TMB is significantly lower compared to HRP (0.0997 vs. 0.434 mM), implying that the mesoporous γ-Fe2O3 sample exhibits a much higher nanozyme activity for TMB than natural HRP. This outcome indicates the potential of the as-prepared mesoporous γ-Fe2O3 as a substitute for HRP. However, in the case of mesoporous a-Fe2O3, the Km value is almost similar or slightly higher (0.5304 vs. 0.434 mM) than that of HRP, indicating that the mesoporous α-Fe2O3 sample is not a suitable alternative to HRP. Nevertheless, the Km values of both the mesoporous α-Fe2O3 and γ-Fe2O3 samples toward H2O2 are substantially higher than HRP (144.30 and 127.92, respectively, vs.

3.70 mM), implying that a higher amount of H2O2 is needed to achieve significant nanozyme activity (at room temperature) for the two mesoporous IO samples. One of the possible reasons for the enhanced nanozyme activity of both mesoporous IO samples (at room temperature) is their mesopore-induced high surface area (enabling more Fe3+ ions to interact with the substrate) and interaction volume. These factors augment the mass transfer and cascade catalysis by keeping the substrates on the surface moiety and increasing the rate of electron transfer, thereby proliferating the kinetics of the oxidation reaction. Another probable reason is the transfer of a higher amount of lone-pair electron cloud (charge) from the amino group (TMB) to the vacant d-orbital of Fe3+, which enhances the mobility and electron density of the 3, 42 IOs to enhance the catalytic activity. Nonetheless, the significantly lower Km value of γ-

Fe2O3 than that of α-Fe2O3 indicates its superiority for biosensing. As stated earlier, α-Fe2O3 possesses a rhombohedral-centred hexagonal structure of the corundum type with a close- packed oxygen lattice with the R3ch(167) space group, where two-thirds of the octahedral sites 3+ 43 are occupied by Fe ions. On the other hand, γ-Fe2O3 possesses a cubic crystal structure of an inverse spinel-type and crystallised in the Fd3m space group, where the oxygen anions have a cubic close-packed array and the Fe3+ ions spread out throughout the tetrahedral sites (FeA) 36 and the octahedral sites (FeB) (inset; Figure 3). Moreover, the crystal structure of γ-Fe2O3 possesses cation vacancies in the octahedral sites. Besides, most of these cation vacancies are located on the surface of γ-Fe2O3. These sites act as adsorption sites for the substrates. 166

Therefore, the mesoporous γ-Fe2O3 can adsorb and hold a much higher amount of H2O2 as well as TMB, leading to its significantly higher peroxidase-mimetic activity toward TMB oxidation.

As evident from Table 3.2, the mesoporous γ-Fe2O3 sample shows a much better nanozyme performance compared to previously reported iron oxide-based nanomaterials and composites. For instance, recently reported mesoporous Fe2O3 nanoflakes exhibited a Km value of 0.24 mM at room temperature, which is almost three times higher than this work.4 In another research study, Prussian blue-modified γ-Fe2O3 exhibited a Km value of 0.307 mM for TMB oxidation at room temperature, which is even higher than the Km value of the as-prepared 13 mesoporous γ-Fe2O3. Fe3O4 NPs were reported to possess a very similar Km value to this work (0.098 vs. 0.0997 mM); however, this activity was achieved at 40 oC.34 This high temperature is not suitable for sensings of biological molecules, such as proteins, antibodies or cells, as biomolecules tend to degrade at elevated temperatures.44 In the past few years, several composite materials and biological molecule-based nanozymes with low Km values toward

TMB oxidation have been reported. For instance, ATP- mediated Fe3O4 showed a three times higher Km value than this report. However, this was achieved by modifying the Fe3O4 NPs with adenosine triphosphate (ATP) and carrying out the nanozyme- catalytic reaction at 40 oC.12

Recently, adenosine 5-monophosphate (AMP)-modified Fe3O4 was also reported, where the modification of the Fe3O4 NPs with phosphate-containing adenosine analogs greatly enhanced the peroxidase mimetic activity toward the oxidation of amplex Ultra-red (AU) (a Km value of 14 0.036 mM) and the adenosine analogs followed the trend: AMP-Fe3O4 > ATP-Fe3O4. To achieve such high activity, the Fe3O4 NPs needed to be modified with AMP under proper o conditions, including a high temperature of 37 C. Remarkably, our mesoporous γ-Fe2O3 can achieve a similarly high level of activity without any chemical or biological modification, indicating the benefit of the mesoporous structure and importance of phase tuning.

167

Figure 3.13: Steady-state kinetics and catalytic mechanism of mesoporous Fe2O3. The velocity

(Vo) of the reaction was measured by using 5 µg of Fe2O3 IOs in 200 mL of 0.2 M NaAc buffer, pH 3.5 at 25 oC. (a and b) Peroxidase-memetic responses over varying concentrations of TMB with the concentration of H2O2 being kept constant and over varying levels of H2O2 with the concentration of TMB being kept constant, respectively, for mesoporous γ-Fe2O3. (c and d) Peroxidase-memetic responses over varying concentrations of TMB with the concentration of

H2O2 being kept constant and over varying concentrations of H2O2 with the concentration of

TMB being held constant, respectively, for mesoporous α-Fe2O3. Inset images show double- reciprocal plots for the activity of respective IOs. Error bars represent the standard error derived from three repeated measurements.

3.3.3. Detection and estimation of glucose

To check the applicability and superiority of the mesoporous γ-Fe2O3 over a-Fe2O3, we have utilized both nanozymes for colourimetric (naked eye) glucose detection. Glucose is one of the most prominent biomolecules, and its aberrancy signifies the onset and progression of several diseases, especially diabetes mellitus and diabetes-related diseases, such as kidney failure, heart disease, and blindness.45-46 Therefore, a great deal of research has been carried out to develop robust and simple point-of-care (POC) diagnostics to monitor and manage glucose- associated diseases in an easy and simple manner. In recent years, nanozymes have shown

168

promising potential in glucose sensors.47 In a typical nanozyme-based glucose detection, glucose solution was initially oxidized by the GOx enzyme to generate glucose equivalent to the generated H2O2, which is subsequently detected by employing a TMB substrate solution 48 and nanozyme, as depicted in Figure 3.14a. In the presence of the generated H2O2, the nanozyme oxidizes the colourless TMB solution to a blue colour TMBox charge-transfer complex solution, which can be quantified by measuring its absorbance at 652 nm. The intensity or absorbance of the generated blue colour is equivalent to the generated H2O2 as well as the amount of glucose present in the sample. To check the capability of both mesoporous IO samples toward glucose detection, we have employed both samples as nanozymes with the same concentration of glucose solution (250 µM). In comparison to the control sample (no mesoporous IO), the mesoporous α-Fe2O3 sample generates a pale blue colour, whereas the mesoporous γ-Fe2O3 sample generates a highly intense blue-coloured solution (inset, Figure

3.14b). The mesoporous γ-Fe2O3 sample gives a much higher absorbance of 0.816 (at 652 nm) compared to 0.089 for mesoporous α-Fe2O3, indicating its superiority and excellent potentiality for glucose detection. To estimate the sensitivity for glucose detection, a series of diluted glucose solutions with concentrations ranging from 1 mM to 1 µM were tested. As shown in Figure 3.14d (inset), there is an increasing trend in the intensity of the blue colour and the absorbance with an increasing concentration of glucose (Figure 3.14c). This can be explained by the fact that a higher amount of glucose can generate a higher amount of H2O2 upon oxidation by GOx, which consequently enhances the nanozyme-catalyzed oxidation of TMB. As evident in Figure 3.14e, there is a linear increase in absorbance (colourimetric readout) with an increased amount of glucose. The linear regression equation can be approximated as y = 0.2133x - 0.1429, with a r2 (correlation coeﬃcient) of 0.9603. The limit of detection (LOD) for the naked eye was evaluated by observing the colour intensity and found to be 1 µM, which is clearly distinguishable from the no-target/ template (NoT) control (PBS was used instead of glucose). The colourimetric LOD was also evaluated by considering a signal-to-noise ratio of 3.0 and determined to be 0.9 µM with good inter-assay reproducibility (relative standard deviation <5%, for n = 3), which is much more sensitive than recently reported nanozyme-based glucose sensors. For instance, this assay is almost ten-times more sensitive (a LOD of 0.9 µM vs. 8.16 µM) than recently reported GOx@ZIF-8(NiPd) nanoflowers.49 A very similar LOD (0.9 µM vs. 1.12 µM) for glucose was reported for Fe3O4@C yolk-shell nanoparticles. However, such a low LOD was achieved by conducting the TMB oxidation reaction at 60 oC.50 Such a high- temperature reaction is not suitable for achieving a simple and portable glucose sensor. The 169

mesoporous γ-Fe2O3 sample has shown good potential to replace natural enzymes for TMB oxidation and reduce the burdens (cost, handling, and storage) associated with natural enzymes. Moreover, it exhibits good stability even after seven days of preparation. The application of this mesoporous IO sample is not limited to glucose detection; it may also be applied as an ideal alternative to conventional ELISA-based sensors for the detection of many other clinically relevant protein-based biomarkers by choosing proper recognition molecules, targets, and antibody functionalization with the signalling nanozyme. Notably, the as-prepared mesoporous IO samples (without any modification and functionalization) are capable of oxidizing TMB even at room temperature, which renders them highly suitable for paper and strip-based nanozyme- modified biosensors.

Figure 3.14: (a) Schematic representation of nanozyme-based glucose detection. (b)The mean responses of absorbance obtained for the assay with mesoporous α-Fe2O3 versus γ-Fe2O3 and a control sample (NoT, PBS was used instead of glucose). The inset shows the corresponding color change. (c) The absorbance spectra obtained for the designated concentration of standard glucose samples ranging from 1 µM to 1000 µM. The corresponding bar diagram for standard

170

glucose concentration are depicted in (d, inset shows the corresponding colour change). (e)

Calibration curve for glucose detection using γ-Fe2O3 as nanozyme with the range of glucose concentration from 1 µM to 1000 µM

Table 3.2: Comparison of the peroxidase-mimicking activities (kinetic parameters and conditions) of the as-prepared mesoporous α-Fe2O3 and γ-Fe2O3 with recently reported iron oxide-based nanostructures and composites for TMB/H2O2 substrate.

-1 o Sample Substrate Km (mM) Vmax (Ms ) pH Temp. ( C) Ref. Mesoporous TMB 0.0997 5.20×10-7

-8 γ-Fe2O3 H2O2 144.30 1.84×10 Mesoporous TMB 0.5304 5.43×10-8 3.5 25 This

-8 work α-Fe2O3 H2O2 127.92 3.77×10 Horseradish peroxidase TMB 0.434 10.0×10-8 4 40 34

-8 (HRP) H2O2 3.7 8.71×10 N-doped porous carbon TMB 0.135 6.13×10-8 4.5 40 51

-8 nanospheres (N-PCNSs) H2O2 161 11.7×10 Iron oxide nanoflakes TMB 0.24 3.07×10-8 3.5 25 4

o -8 (C@250 C) H2O2 150.47 3.12×10

-8 8 Mesoporous Fe2O3 TMB 0.298 8.71×10 3.5 25

-8 H2O2 146.7 10×10

52 Au/CeO2 CSNPs TMB 0.29 3.9 4 40

H2O2 44.69 2.23

-8 12 ATP-mediated Fe3O4 TMB 0.374 2.6×10 4 40

-8 H2O2 54.6 1.8×10 Prussian Blue-modified γ- TMB 0.307 1.06×10-6 4.6 25 13

-6 Fe2O3 H2O2 323.6 1.17×10 Graphene oxide (GO)- TMB 0.118 5.38×10-8 3.6 45 6

-7 Fe2O3 H2O2 305 1.01×10

-7 53 FeNPs@Co3O4 hollow TMB 0.488 2.06×10 3.5 35

-7 nanocages H2O2 0.019 0.17×10

-6 14 AMP-Fe3O4 NPs AU 0.036 1.55×10 7.0 35

171

3.4. Conclusion This work has demonstrated that mesoporous IO samples with two diﬀerent crystal phases (a- and g-) exhibit diﬀerent peroxidase mimetic activity. Both mesoporous α-Fe2O3 and γ-Fe2O3 exhibit room-temperature peroxidase mimetic activity. Owing to the favourable crystal phase, the mesoporous γ-Fe2O3 sample exhibited around seven-time higher responses in peroxidase activity than the mesoporous α-Fe2O3 sample. The applicability of these mesoporous IOs as nanozymes was verified by the colourimetric detection of glucose, where the mesoporous γ-

Fe2O3 sample shows almost an eight times higher response than mesoporous α-Fe2O3. The mesoporous γ-Fe2O3 sample exhibits very high sensitivity for colourimetric glucose detection with an impressive LOD of 0.9 µM. The findings of this work will provide an in-depth understanding of the eﬀect of the crystal phase of IOs on their peroxidase mimetic activity. We envisage that this work will provide useful guidance for designing highly eﬀective nanozymes based on IO nanoarchitectures for the sensing of glucose and other biomolecules in the future.

3.5. References 1. Gao, L.; Fan, K.; Yan, X., Iron oxide nanozyme: a multifunctional enzyme mimetic for biomedical applications. Theranostics 2017, 7 (13), 3207. 2. Benedetti, T. M.; Andronescu, C.; Cheong, S.; Wilde, P.; Wordsworth, J.; Kientz, M.; Tilley, R. D.; Schuhmann, W.; Gooding, J. J., Electrocatalytic nanoparticles that mimic the three-dimensional geometric architecture of enzymes: nanozymes. Journal of the American Chemical Society 2018, 140 (41), 13449-13455. 3. Masud, M. K.; Yadav, S.; Islam, M. N.; Nguyen, N.-T.; Salomon, C.; Kline, R.; Alamri, H. R.; Alothman, Z. A.; Yamauchi, Y.; Hossain, M. S. A., Gold-loaded nanoporous ferric oxide nanocubes with peroxidase-mimicking activity for electrocatalytic and colorimetric detection of autoantibody. Analytical Chemistry 2017, 89 (20), 11005- 11013. 4. Tanaka, S.; Masud, M. K.; Kaneti, Y. V.; Shiddiky, M. J.; Fatehmulla, A.; Aldhafiri, A. M.; Farooq, W. A.; Bando, Y.; Hossain, M. S. A.; Yamauchi, Y., Enhanced Peroxidase Mimetic Activity of Porous Iron Oxide Nanoflakes. ChemNanoMat 2019, 5 (4), 506-513. 5. Dutta, A. K.; Maji, S. K.; Srivastava, D. N.; Mondal, A.; Biswas, P.; Paul, P.; Adhikary,

B., Peroxidase-like activity and amperometric sensing of hydrogen peroxide by Fe2O3

and Prussian Blue-modified Fe2O3 nanoparticles. Journal of Molecular Catalysis A: Chemical 2012, 360, 71-77. 172

6. Song, Y.; Qu, K.; Zhao, C.; Ren, J.; Qu, X., Graphene oxide: intrinsic peroxidase catalytic activity and its application to glucose detection. Advanced Materials 2010, 22 (19), 2206-2210. 7. Huang, Y.; Liu, Z.; Liu, C.; Ju, E.; Zhang, Y.; Ren, J.; Qu, X., Self‐assembly of multi‐ nanozymes to mimic an intracellular antioxidant defense system. Angewandte Chemie International Edition 2016, 55 (23), 6646-6650. 8. Bhattacharjee, R.; Tanaka, S.; Moriam, S.; Masud, M. K.; Lin, J.; Alshehri, S. M.; Ahamad, T.; Salunkhe, R. R.; Nguyen, N.-T.; Yamauchi, Y., Porous nanozymes: the peroxidase-mimetic activity of mesoporous iron oxide for the colorimetric and electrochemical detection of global DNA methylation. Journal of Materials Chemistry B 2018, 6 (29), 4783-4791. 9. Boriachek, K.; Masud, M. K.; Palma, C.; Phan, H.-P.; Yamauchi, Y.; Hossain, M. S. A.; Nguyen, N.-T.; Salomon, C.; Shiddiky, M. J., Avoiding Pre-Isolation Step in Exosome Analysis: Direct Isolation and Sensitive Detection of Exosomes Using Gold- Loaded Nanoporous Ferric Oxide Nanozymes. Analytical Chemistry 2019, 91 (6), 3827-3834. 10. Wang, Q.; Wei, H.; Zhang, Z.; Wang, E.; Dong, S., Nanozyme: An emerging alternative to natural enzyme for biosensing and immunoassay. TrAC Trends in Analytical Chemistry 2018, 105, 218-224. 11. Wei, H.; Wang, E., Nanomaterials with enzyme-like characteristics (nanozymes): next- generation artificial enzymes. Chemical Society Reviews 2013, 42 (14), 6060-6093. 12. Vallabani, N. S.; Karakoti, A. S.; Singh, S., ATP-mediated intrinsic peroxidase-like

activity of Fe3O4-based nanozyme: one step detection of blood glucose at physiological pH. Colloids and Surfaces B: Biointerfaces 2017, 153, 52-60. 13. Zhang, X.-Q.; Gong, S.-W.; Zhang, Y.; Yang, T.; Wang, C.-Y.; Gu, N., Prussian blue modified iron oxide magnetic nanoparticles and their high peroxidase-like activity. Journal of Materials Chemistry 2010, 20 (24), 5110-5116. 14. Yang, Y.-C.; Wang, Y.-T.; Tseng, W.-L., Amplified peroxidase-like activity in iron oxide nanoparticles using adenosine monophosphate: Application to urinary protein sensing. ACS applied Materials & Interfaces 2017, 9 (11), 10069-10077. 15. Zhang, Z.; Zhang, X.; Liu, B.; Liu, J., Molecular imprinting on inorganic nanozymes for hundred-fold enzyme specificity. Journal of the American Chemical Society 2017, 139 (15), 5412-5419.

173

16. Roy, A.; Sahoo, R.; Ray, C.; Dutta, S.; Pal, T., Soft template induced phase selective

synthesis of Fe2O3 nanomagnets: one step towards peroxidase-mimic activity allowing colorimetric sensing of thioglycolic acid. RSC Advances 2016, 6 (38), 32308-32318. 17. Sakurai, S.; Namai, A.; Hashimoto, K.; Ohkoshi, S.-i., First observation of phase

transformation of all four Fe2O3 phases (γ→ ε→ β→ α-phase). Journal of the American Chemical Society 2009, 131 (51), 18299-18303. 18. Chaudhari, K. N.; Chaudhari, N. K.; Yu, J.-S., Peroxidase mimic activity of hematite

iron oxides (α-Fe2O3) with different nanostructures. Catalysis Science & Technology 2012, 2 (1), 119-124.

19. Liu, J.; Yang, H.; Xue, X., Preparation of different shaped α-Fe2O3 nanoparticles with large particles of iron oxide red. CrystEngComm 2019, 21 (7), 1097-1101. 20. Zboril, R.; Mashlan, M.; Petridis, D., Iron (III) oxides from thermal processes synthesis, structural and magnetic properties, Mössbauer spectroscopy characterization, and applications. Chemistry of Materials 2002, 14 (3), 969-982.

21. Chen, J.; Xu, L.; Li, W.; Gou, X., α‐Fe2O3 nanotubes in gas sensor and lithium‐ion battery applications. Advanced Materials 2005, 17 (5), 582-586. 22. Petkov, V.; Cozzoli, P. D.; Buonsanti, R.; Cingolani, R.; Ren, Y., Size, shape, and internal atomic ordering of nanocrystals by atomic pair distribution functions: a

comparative study of γ-Fe2O3 nanosized spheres and tetrapods. Journal of the American Chemical Society 2009, 131 (40), 14264-14266. 23. Jiao, F.; Jumas, J.-C.; Womes, M.; Chadwick, A. V.; Harrison, A.; Bruce, P. G.,

Synthesis of ordered mesoporous Fe3O4 and γ-Fe2O3 with crystalline walls using post- template reduction/oxidation. Journal of the American Chemical Society 2006, 128 (39), 12905-12909. 24. Krishna, N. V.; Selvam, P., Acid‐Mediated Synthesis of Ordered Mesoporous Aluminosilicates: The Challenge and the Promise. Chemistry–A European Journal 2017, 23 (7), 1604-1612. 25. Kuppan, B.; Selvam, P., Platinum-supported mesoporous carbon (Pt/CMK-3) as anodic catalyst for direct methanol fuel cell applications: The effect of preparation and deposition methods. Progress in Natural Science: Materials International 2012, 22 (6), 616-623. 26. Benyettou, F.; Ocadiz Flores, J. A.; Ravaux, F.; Rezgui, R.; Jouiad, M.; Nehme, S. I.; Parsapur, R. K.; Olsen, J. C.; Selvam, P.; Trabolsi, A., Mesoporous γ‐iron oxide

174

nanoparticles for magnetically triggered release of doxorubicin and hyperthermia treatment. Chemistry–A European Journal 2016, 22 (47), 17020-17028. 27. Jun, S.; Joo, S. H.; Ryoo, R.; Kruk, M.; Jaroniec, M.; Liu, Z.; Ohsuna, T.; Terasaki, O., Synthesis of new, nanoporous carbon with hexagonally ordered mesostructure. Journal of the American Chemical Society 2000, 122 (43), 10712-10713. 28. Lehninger, A. L.; Nelson, D. L.; Cox, M. M.; Cox, M. M., Lehninger principles of biochemistry. Macmillan: 2005. 29. Lineweaver, H.; Burk, D., The determination of enzyme dissociation constants. Journal of the American chemical society 1934, 56 (3), 658-666. 30. Ren, Y.; Ma, Z.; Bruce, P. G., Ordered mesoporous metal oxides: synthesis and applications. Chemical Society Reviews 2012, 41 (14), 4909-4927. 31. Kucheryavy, P.; He, J.; John, V. T.; Maharjan, P.; Spinu, L.; Goloverda, G. Z.; Kolesnichenko, V. L., Superparamagnetic iron oxide nanoparticles with variable size and an iron oxidation state as prospective imaging agents. Langmuir 2013, 29 (2), 710- 716. 32. Wang, Y.; Ren, J.; Liu, X.; Wang, Y.; Guo, Y.; Guo, Y.; Lu, G., Facile synthesis of

ordered magnetic mesoporous γ-Fe2O3/SiO2 nanocomposites with diverse mesostructures. Journal of Colloid and Interface Science 2008, 326 (1), 158-165. 33. Moriya, T., Anisotropic superexchange interaction and weak ferromagnetism. Physical Review 1960, 120 (1), 91. 34. Gao, L.; Zhuang, J.; Nie, L.; Zhang, J.; Zhang, Y.; Gu, N.; Wang, T.; Feng, J.; Yang, D.; Perrett, S., Intrinsic peroxidase-like activity of ferromagnetic nanoparticles. Nature Nanotechnology 2007, 2 (9), 577. 35. Wang, H.; Wan, K.; Shi, X., Recent advances in nanozyme research. Advanced Materials 2018, 1805368. 36. Machala, L.; Tucek, J.; Zboril, R., Polymorphous transformations of nanometric iron (III) oxide: a review. Chemistry of Materials 2011, 23 (14), 3255-3272. 37. Babay, S.; Mhiri, T.; Toumi, M., Synthesis, structural and spectroscopic characterizations of maghemite γ-Fe2O3 prepared by one-step coprecipitation route. Journal of Molecular Structure 2015, 1085, 286-293. 38. Van Oosterhout, G.; Rooijmans, C., A new superstructure in gamma-ferric oxide. Nature 1958, 181 (4601), 44.

175

39. André, R.; Natálio, F.; Humanes, M.; Leppin, J.; Heinze, K.; Wever, R.; Schröder, H.

C.; Müller, W. E.; Tremel, W., V2O5 nanowires with an intrinsic peroxidase‐like activity. Advanced Functional Materials 2011, 21 (3), 501-509. 40. Singh, V. K.; Yadav, P. K.; Chandra, S.; Bano, D.; Talat, M.; Hasan, S. H., Peroxidase mimetic activity of fluorescent NS-carbon quantum dots and their application in

colorimetric detection of H2O2 and glutathione in human blood serum. Journal of Materials Chemistry B 2018, 6 (32), 5256-5268. 41. Barreto, J. C.; Smith, G. S.; Strobel, N. H.; McQuillin, P. A.; Miller, T. A., Terephthalic acid: a dosimeter for the detection of hydroxyl radicals in vitro. Life sciences 1994, 56 (4), PL89-PL96. 42. Xing, Z.; Tian, J.; Asiri, A. M.; Qusti, A. H.; Al-Youbi, A. O.; Sun, X., Two-

dimensional hybrid mesoporous Fe2O3–graphene nanostructures: a highly active and reusable peroxidase mimetic toward rapid, highly sensitive optical detection of glucose. Biosensors and Bioelectronics 2014, 52, 452-457. 43. Li, S.-S.; Li, W.-J.; Jiang, T.-J.; Liu, Z.-G.; Chen, X.; Cong, H.-P.; Liu, J.-H.; Huang, Y.-Y.; Li, L.-N.; Huang, X.-J., Iron oxide with different crystal phases (α-and γ-Fe2O3) in electroanalysis and ultrasensitive and selective detection of lead (II): an advancing approach using XPS and EXAFS. Analytical chemistry 2015, 88 (1), 906-914. 44. Meersman, F.; Heremans, K., Temperature-induced dissociation of protein aggregates: accessing the denatured state. Biochemistry 2003, 42 (48), 14234-14241. 45. Alicic, R. Z.; Rooney, M. T.; Tuttle, K. R., Diabetic kidney disease: challenges, progress, and possibilities. Clinical Journal of the American Society of Nephrology 2017, 12 (12), 2032-2045. 46. Blake, D. R.; Meigs, J. B.; Muller, D. C.; Najjar, S. S.; Andres, R.; Nathan, D. M., Impaired glucose tolerance, but not impaired fasting glucose, is associated with increased levels of coronary heart disease risk factors: results from the Baltimore Longitudinal Study on Aging. Diabetes 2004, 53 (8), 2095-2100. 47. Masud, M. K.; Kim, J.; Billah, M. M.; Wood, K.; Shiddiky, M. J.; Nguyen, N.-T.; Parsapur, R. K.; Kaneti, Y. V.; Alshehri, A. A.; Alghamidi, Y. G., Nanoarchitectured peroxidase-mimetic nanozymes: mesoporous nanocrystalline α-or γ-iron oxide? Journal of Materials Chemistry B 2019, 7 (35), 5412-5422.

48. Wei, H.; Wang, E., Fe3O4 magnetic nanoparticles as peroxidase mimetics and their

applications in H2O2 and glucose detection. Analytical Chemistry 2008, 80 (6), 2250- 2254. 176

49. Wang, Q.; Zhang, X.; Huang, L.; Zhang, Z.; Dong, S., GOx@ ZIF‐8 (NiPd) nanoflower: an artificial enzyme system for tandem catalysis. Angewandte Chemie International Edition 2017, 56 (50), 16082-16085. 50. Lu, N.; Zhang, M.; Ding, L.; Zheng, J.; Zeng, C.; Wen, Y.; Liu, G.; Aldalbahi, A.; Shi,

J.; Song, S., Yolk–shell nanostructured Fe3O4@C magnetic nanoparticles with

enhanced peroxidase-like activity for label-free colorimetric detection of H2O2 and glucose. Nanoscale 2017, 9 (13), 4508-4515. 51. Fan, K.; Xi, J.; Fan, L.; Wang, P.; Zhu, C.; Tang, Y.; Xu, X.; Liang, M.; Jiang, B.; Yan, X., In vivo guiding nitrogen-doped carbon nanozyme for tumor catalytic therapy. Nature Communications 2018, 9 (1), 1440. 52. Bhagat, S.; Vallabani, N. S.; Shutthanandan, V.; Bowden, M.; Karakoti, A. S.; Singh, S., Gold core/ceria shell-based redox active nanozyme mimicking the biological multienzyme complex phenomenon. Journal of Colloid and Interface Science 2018, 513, 831-842.

53. Zhao, J.; Dong, W.; Zhang, X.; Chai, H.; Huang, Y., FeNPs@ Co3O4 hollow nanocages hybrids as effective peroxidase mimics for glucose biosensing. Sensors and Actuators B: Chemical 2018, 263, 575-584.

177

Chapter 4

Gold-Loaded Nanoporous Ferric Oxide Nanocubes with Peroxidase- Mimicking Activity for Electrocatalytic and Colorimetric Detection of Autoantibody*

*Sections of this chapter are based on Masud, M.K., et al., Analytical Chemistry, 2017, 89 (20), 11005-11013.

178

4. Gold-Loaded Nanoporous Ferric Oxide Nanocubes with Peroxidase-Mimicking Activity for Electrocatalytic and Colorimetric Detection of Autoantibody

4.1. Introduction Iron oxide (IO) nanostructures have widely been used in a variety of biomedical applications, such as tissue engineering, magnetic resonance imaging, hyperthermia, drug and gene targeting, isolation and separation of proteins or cells from samples, and in-vivo cell tracking.1- 2 The paramagnetic properties of IO nanoparticles also allow for contactless sample preparation and handling using the emerging technology of micromagnetofluidics.3 In recent years, it has been demonstrated that IO based nanoparticles (NPs) possess an intrinsic horseradish peroxidase (HRP)-mimicking activity towards the oxidation of common chromogenic substances such as 3,3ˊ,5,5ˊ-tertamethylbenzidine (TMB), di-azo-aminobenzene, and o- phenylenediamine , which have widely been used in catalytic decomposition of hydrogen peroxides or developing non-enzymatic glucose sensors.4-6 For instance, the peroxidase-like activity of Fe2O3, Prussian blue/Fe2O3, graphene/Fe2O3, GO_MNP(Fe3O4)/Pt, ZnFe2O4NPs have extensively been used to develop several high-performance biosensors for detecting 7-11 glucose, H2O2, cancer cells, L-cysteine etc. Similar to IO based NPs, other nanomaterials such as AuNPs, Eu2O2SNPs, CuONPs, copper–creatinine complex, AgX (X= Cl, Br, I), polyoxometalates, PtNPs, carbon nanotubes and nanodots have also been reported to exhibit peroxidase-like activity.6, 12-19 The use of these peroxidase-mimetic nanomaterials for developing biosensors is highly attractive due to several reasons. First, unlike natural peroxidase (i.e., HRP) they are stable towards protease digestion or denaturation, and their structure, morphology and function are not affected by environmental stresses.20 Second, they are relatively easy and inexpensive to prepare and highly suitable for benchtop storage and handling. Despite these advantages, most of these materials demonstrate their highest peroxidase-like activity at a higher temperature (40 – 45 °C), which limits their applications in disease-specific biomolecules detection at room temperature. Therefore, the design and synthesis of NPs with enhanced peroxidase-mimetics activity at room temperature are highly desirable. Recently, we have synthesized a new class of gold-loaded superparamagnetic ferric 21 oxide nanocube (Au-NPFe2O3NC). This Au-NPFe2O3NC exhibits multiple functionalities. First, it shows enhanced catalytic activity towards the common electroactive molecules (i.e., 22 hexaammineruthenium(III) chloride, (Ru(NH3)6Cl3)). Second, highly porous framework of gold loaded Fe2O3 nanocubes allows for the direct adsorption of a large amount of target DNA,

179

RNA or protein via gold-DNA/RNA/Proteins affinity interactions.21, 23-26 Third, as these nanocubes are paramagnetic, they can be used as dispersible capture vehicles to isolate the circulating biomarkers in body fluids (i.e., they can be dispersed into the sample and bind to the analytes of interest).21 As the capture event of the target analytes of interest temporally and spatially separated from the electrochemical measurement, this alleviates biofouling issues of the electrode. The Au-NPFe2O3NC nanocubes also provide a means to reduce the biological noises associated with adsorption of non-specific species present in the body fluids via 1 magnetic enrichment, separation, and purification steps. Fourth, the Au-NPFe2O3NC nanocubes can also be used as nanoenzymes with enhanced peroxidase-like activity. Although the functionalities of Au-NPFe2O3NC as electrocatalyst and dispersible capture vehicle have 21-22 already been explored, the peroxidase-like activity of Au-NPFe2O3NC has yet to be demonstrated. In this paper, we studied this peroxidase-like activity for the development of a proof-of-concept molecular sensor for detecting circulating autoantibodies in serum or plasma samples. Circulating autoantibodies that are elicited in responses to the tumor-associated antigen (TAA) are emerging biomarkers for the early detection of cancer, as they are produced by the patient’s immune system long (several months or years) before the onset of the clinical symptoms of diseases.27-29 The direct quantification of TAAs in clinical samples possess severe challenges due to their low abundance and associated difficulties in identifying minor structural modification or mutation. In contrast, serum autoantibodies are relatively more stable (i.e., longer half-lives due to limited proteolysis and clearance) and present as large quantities in clinical samples and therefore they have been used as circulating reporters for the early or preclinical detection of various cancers including ovarian, lung, and breast cancer.30-34 It is also important to note that, with the progression of cancer (stages), the amount of autoantibody (produced) compared to that of TAAs is highly significant and readily detectable with the conventional detection techniques.35 Most of the widely used methods for serum autoantibody detection are mainly based on ELISA or protein arrays that employ HRP conjugated protein- specific secondary antibodies to read the target autoantibody.31-37 These methods are, however, expensive, relatively less sensitive and can only provide qualitative or semi-quantitative results. Until recently, the most advanced assay for the sensitive detection of serum autoantibody is that developed by Asensio et al., where HaloTag fusion p53 protein modified commercial magnetic beads (MBs) were used as magnetic microcarriers for capturing the autoantibodies in sera.38 HRP conjugated anti-human IgG was then used to quantify captured autoantibody via colourimetry and amperometry. More recently, we have also developed a method using Au- 180

21 NPFe2O3NC as a dispersible capture agent for autoantibody isolation and detection. Although the analytical performance of these methods are superior, they still rely on HRP-based enzymatic reaction.

Herein, we first studied the peroxidase-like activity of Au-NPFe2O3NC nanocubes towards the catalytic oxidation of TMB in the presence of H2O2. This feature of Au-

NPFe2O3NC was then used to develop a molecular sensor for the colourimetric (naked-eye) and electrochemical detection of p53 autoantibody in serum or plasma samples. The method was first tested on the sample obtained from commercial p53 ELISA kit and finally challenged with a small cohort of plasma samples derived from epithelial ovarian cancer high-grade serous subtype (EOCHGS).

4.2. Experimental Section 4.2.1. Materials and instrumentations Unless otherwise stated, the reagents and chemicals used for this study were of analytical grade. Recombinant human p53 protein were purchased from Abcam (Australia), anti-human IgG antibody was obtained from Thermo Fisher Scientific (Australia). Bovine serum albumin (BSA) was used as a blocking agent and purchased from Life Technologies (Australia). p53 autoantibody ELISA kit was purchased from Dianova GmbH ( Germany). PBS tablet (0.01M phosphate buffer, 0.0027M potassium chloride and 0.137M sodium chloride, pH 7.4 at 25oC) and TMB were purchased from Sigma life Science (Australia). Analytical grade H2O2, dimethyl sulfoxide (DMSO), hydrochloric acid (HCl) were purchased from chem-supply (Australia). Extravidin modified screen-printed carbon electrode (SPCE-XTR) (DRP- 110XTR) and screen-printed gold electrode (SPE-Au) (220BT) with a three-electrode system printed on a ceramic substrate were acquired from Dropsens (Spain). In the three-electrode system of SPCE-XTR, working, counter and reference electrodes were extravidin/carbon, carbon and silver-modified respectively, while SPE-Au was comprised of gold working and auxiliary electrodes with Ag-modified reference electrode. All chemical and reagent were used as received without additional purification. UltrapureTM DNase/RNase-free distilled water (Invitrogen, Australia) was used throughout the experiments. Scanning electron microscope (SEM) images were taken with a Hitachi S-4800 scanning microscope with the accelerating voltage of 10 kV. Wide-angle X-ray diffraction (XRD) patterns were obtained using Rigaku RINT 2500X with monochromated Cu-Ka radiation (λ = 1.54 °A, 40 kV, 40 mA). Scanning electron microscope (SEM) images were obtained with a Hitachi SU8000 operated at an accelerating of 5 kV. Cross-sectional 181

transmission electron microscope (TEM) and HAADF-STEM (high-angle annular dark-field scanning transmission electron microscope) images were taken with a JEM-2100F operated at an accelerating voltage of 200 kV. The elemental chemical analysis was performed by X-ray photoelectron spectroscopy (XPS, PHI Quantera SXM, ULVAC-PHI Inc., Japan). Samples were degassed in a vacuum before carrying out the electrochemical measurements. All electrochemical measurements were carried out with a CHI650 electrochemical workstation

(CH instrument, USA). Au-NPFe2O3NC nanocubes were sonicated with a controlled ultrasonic water bath (Soniclean, Australia) before conducting the experiments. A DynaMag 2 magnetic separation rack from Thermo Fisher Scientific (Australia) and microtube mixer from Eppendorf (Germany) were also employed for magnetic washing and functionalization of nanocubes with IgG.

4.2.2. Study group and ovarian cancer samples Staged samples (cross-sectional) were collected at the Ochsner Baptist Medical Center in the clinical trials and obtained via The UQ Centre for Clinical research. Plasma samples were obtained following the declaration of Helsinki and approved by the Ethics Committee of The University of Queensland and the Ochsner Medical Center (New Orleans, USA). Plasma was separated from whole blood by centrifugation (2000 g x 10 min at Room temperature) and stored at -80 oC until analyses. Ovarian cancer samples were collected prospectively and assigned according to the histotypes classification (e.g. stage I, and stage III) and stored to -80 °C in the Biobank units. Only patients with epithelial ovarian cancer high-grade serous subtype (n = 2) and benign controls (n = 2) were included in this study.

4.2.3. Determination of the surface area of the electrode Cyclic voltammetry (CV) measurements were performed between -200 and 600 mV unless stated otherwise. The effective areas of SPC-Au was determined by the measurement of the peak current obtained as a function of scan rate under cyclic voltammetric conditions for the 3-/4- one-electron reduction of [Fe(CN)6] [2.5 mM in PBS (0.5 M KCl)] and by using the Randles-Sevcik equation (eq 1),39-40

5 3/ 2 1/ 2 1/ 2 ip  (2.6910 ) n AD C ...... … … (1)

3+ 2+ where, ip is the peak current (A), n is the number of electrons transferred (Fe → Fe , n = 1), 2 3- A is the effective area of the electrode (cm ), D is the diffusion coefficient of [Fe(CN)6] (taken to be 8.0 × 10-6 cm2s-1, C is the concentration (mol cm-3), ν is the scan rate (Vs-1).41

182

4.2.4. Preparation of IgG/Au-NPFe2O3NC nanocatalyst 42 IgG/Au-NPFe2O3NC was prepared following a published protocol. Briefly, 50 µL of nanocubes (1mg/mL) and 55 μL of IgG (100ng/mL) antibody were mixed thoroughly for 2 min by pipetting. 1.5 μL of 1% PEG was then added to the mixture and stirred in a microtube mixer for 30 min at room temperature. To remove the excess (unbounded or loosely bounded) IgG, the mixture was placed on a magnetic separation rack and IgG/Au-NPFe2O3NC nanocatalysts was magnetically separated. The clear solution was carefully pipetted off. 50 μL PBS solution was then added to the residue, mixed and nanocatalysts was magnetically separated. This washing was repeated for three times. The IgG/Au-NPFe2O3NC nanocatalysts was finally resuspended in 50 μL PBS buffer containing 1.5 μL of 1% PEG. The conjugated nanocatalysts was stored at 4 oC for further experiment.

4.2.5. The peroxidase-mimetic activity of Au-NPFe2O3NC

Unless otherwise stated, the peroxidase-like activity of Au-NPFe2O3NC was carried out at room temperature using 5 µg of Au-NPFe2O3NC in 80 µL of reaction buffer (0.2 M sodium acetate (NaAc), pH 3.5) in the presence of 800 µM freshly prepared TMB (TMB dissolved in

DMSO) and 700 mM H2O2. The formation of blue coloured solution was monitored and measured in time scan mode at 652 using a spectrophotometer (SpectraMax). The reaction was quenched by adding 2.0 µL of stop solution. The end-point of the resultant yellow colour product was measured using colourimetry (at 452 nm) and chronoamperometry. The steady- state kinetic assays were carried out using standard reaction condition (described above) by varying the concentration of H2O2 (0.01 to 1.1 M) at a fixed concentration of TMB (800 µM) and vice versa for the varying the concentration of TMB (0.01 to 1.0 mM) at 700 mM H2O2. The apparent kinetic parameters was calculated by considering a typical enzyme catalytic reaction;

where E, S, ES and P represent the enzyme, substrate, enzyme-substrate adduct and product respectively. The Michaelis-Menten equation for the catalytic system is expressed as follows,43

183

Vmax [S] Vo  … … … (2) K m  [S]

In this equation, Vo is the rate of substrate conversion to the product, Vmax is the maximum rate of conversion, which is attained when the active (catalytic) sites on the enzyme are saturated with substrate, [S] is the substrate concentration, and Km is the Michaelis-Menten constant (denotes the affinity of the enzyme for the substrate), which is equivalent to the substrate concentration at the conversion rate is half of Vmax. The rearrangement of the Michaelis-Menten equation gives the Lineweaver–Burk equation,44 which was used to determine enzyme kinetics terms Km and Vmax. 1 K 1 1  m  … … … (3) V o Vmax [S] Vmax

4.2.6. Colorimetric and amperometric detection of serum p53 autoantibody The SPCE-XTR was kept at room temperature for 1 h and washed with PBS. 5.0 μL (100 ng/mL) p53 antigen was incubated onto the electrode surface for 40 min. After the incubation, the electrode surface was washed with PBS three times followed by further incubation with 1% BSA solution (blocking agent) for 15 min. 5.0 μL of diluted serum sample or tested serum sample (1:100 dilution) were then added to the electrode surface and incubated for 1 h to capture the p53-specific autoantibodies present in the sample. After washing away the unbounded or loosely attached serum proteins with PBS, the complex is incubated with 5.0 μL of freshly prepared IgG/Au-NPFe2O3NC for 40 min followed by several PBS wash to remove all unbounded IgG/Au-NPFe2O3NC. Finally, 50 μL of freshly prepared TMB substrate solution

(800 µM TMB, 700 mM H2O2 in 0.2 M NaAc buffer, pH-3.5) was added and incubated for 30 min in dark. The color change was visually observed. For quantitative measurements of the amount of oxidized TMB, 2.0 μL of stop solution (2.0 M HCl) was added and absorbance was recorded at 452 nm with the SpectraMax spectrophotometer. For the electrochemical detection, 30 μL of the oxidized TMB solution was pipetted onto another clean SPE-Au, and amperometric response (i-t) were measured at 150 mV over 120 s. At least three replicates were measured for each standard/sample. All measurements were performed at room temperature.

184

4.3. Result and Discussion

4.3.1. The peroxidase-mimetic activity of Au-NPFe2O3NC It is now well established that natural peroxidase (HRP) can catalyse the oxidation of TMB in the presence of H2O2. This catalytic reaction generates two coloured products. First, HRP/H2O2 catalysed the oxidation of TMB that produced a blue-coloured charge-transfer complex of parent TMB (diamine) and TMB oxidised product (diimine), which could be used for qualitative (naked-eye) evaluation. Second, this blue-coloured complex turned yellow after the addition of an acid.4, 45 The yellow product is electroactive and stable at acidic pH, thus can be quantified by UV-Vis (semi-quantitative, colourimetry) and electrochemical detection 21- methods. We have recently synthesized a novel class of superparamagnetic Au-NPFe2O3NC. 22 The details characterization data of this material have already been described in the next chapter. Briefly, Au-NPFe2O3NC was synthesized by depositing AuNPs on to the nanoporous 21,22 iron oxide nanocubes (NPFe2O3NCs) derived from Prussian blue (PB) nanocubes. Bright field TEM and HAADF-STEM image for this materials clearly showed that they possessed a highly porous structure (Figure 4.1). SEM images in Figure 4.2 resulted that the Au content is well distributed over the entire particles. The peaks in the wide-angle XRD pattern are 46 assignable to Au, α-Fe2O3, and γ-Fe2O3 (Figure 4.3). This material exhibits peroxidase-like activity for the oxidation TMB in the presence of H2O2 at room temperature. Figure 4.4(a), represents the mechanism of the catalysis of TMB (in presence of H2O2) oxidation reaction by

Au-NPFe2O3NC.

Figure 4.1: Bright field TEM image and HAADF-STEM image for gold-loaded nanoporous ferric oxide nanocubes (Au-NPFe2O3NC).

185

Figure 4.2: (a) SEM image and (b-d) elemental mapping for Au-NPFe2O3NC [(b) Fe, (c) Au, and (d) O, respectively].

Figure 4.3: XRD pattern for Au-NPFe2O3NC

186

To assess peroxidase-mimetics of nanocubes, a set of control experiments were conducted. TMB substrate solution in the presence (positive control) and absence (negative control) of nanocubes was incubated in dark. After 10 min of incubation, a positive control sample generated a clear blue coloured solution, while negative control was colourless. Positive control gave an absorbance of 0.96 at 652 nm which is 15-times higher than that of negative control (abs. 0.96 versus 0.053) samples (Figure 1b). Addition of 2.0 M HCl into the solution, turned the blue-coloured complex to an electroactive yellow-coloured diimine product. The electrochemical quantification of the oxidised product was conducted by placing the reaction mixture onto a screen-printed gold electrode (SPE-Au) via of amperometry. Figure 1(c) shows the amperometric signals where positive control generated 86.78 µA cm-2 current density compared to that of the negative control (4.59 µA cm-2). These experiments (i.e., naked- eye observation, colourimetric and amperometric readouts) clearly confirm the peroxidase-like activity of our novel Au-NPFe2O3NC nanocubes. Notably, unlike most of the existing nanomaterials,7-18 which possess the highest peroxidase-like activity at 40 – 45 °C, our Au-

NPFe2O3NC nanocubes exhibit relatively enhanced activity at room temperature.

Figure 4.4: (a) Schematic illustration of peroxidase-mimicking activity of Au-NPFe2O3NC for the oxidation of TMB in the presence of H2O2. Mean values of (b) absorbance (UV-vis) and (c) amperometric current signals for the negative and positive control samples (inset in (b) and (c) are the corresponding photos for the naked eye evaluation and i-t curves respectively).

187

The level of peroxidase-like activity of Au-NPFe2O3NC nanocubes at room temperature is sufficient to generate approximately 20-times higher response in comparison with the negative control sample (Figure 1C). One possible explanation of this enhanced response could be due to (i) the large surface of the Au-NPFe2O3NC nanocubes (i.e., highly porous) that facilitates the binding of an increased amount of positively charged TMB and hence the TMB/H2O2 reaction, and (ii) the presence of 2% AuNPs in the nanocubes that also catalyse the TMB/H2O2 reaction.

In this nanocubes catalysed TMB/H2O2 reaction, ferric ions (from nanocubes) initiate . the reaction by generating hydroxyl free radical ( OH) from H2O2 following the Fenton reaction.47-48 TMB is then oxidised by the generated .OH, as shown below

3 2  Fe  H 2O2  FeOOH  H (1)

2 2 FeOOH  Fe  HO2  (2)

2 3  Fe  H 2O2  Fe  OH OH (3)

OH TMB TMBoxBlue (4)

Similar to HRP, the activity of Au-NPFe2O3NC is also dependent on the solution pH and amount of nanocubes. In order to check the effect of pH, we studied the peroxidase-like activity of nanocubes using different solution pH ranging from 2.5 to 5.5. We observed that with increasing pH of the solution the peroxidase-like activity of Au-NPFe2O3NC decrease (Figure 4.5a). This is because the addition of nanocubes may accelerate the processes (equation

2 1) and (equation 2) (see above) and produce an excessive amount of FeOOH and HO2 . At

- 49 higher pH, can be ionized to O2 , as shown by equation (equation 5). The can also instantly react with hydroxyl radicals and produce oxygen (equation 6).

  HO2 .  H  O2 (5)

 OH  HO2  /O2  H 2O  O2 (6) Among all studied pH values, pH 2.5 resulted in the highest responses both in colourimetry and amperometric readouts. As the iron could be leached from the nanocubes in the solution pH of <3,7 we selected pH 3.5 as the optimal pH for conducting subsequent experiments in our study. To optimize the amount of nanocubes in our assay, 5 μL of designated concentrations (i.e., 2.5 (0.5 μg/μL), 5 (0.5 μg/μL), 10, (2 μg/μL) and 20 μg (4 μg/μL)) of nanocubes were used. As can be seen in Figure 4.5b1; 5, 10 and 20 μg of nanocubes produced almost similar responses both in colourimetry and amperometric readouts, and therefore we chose 5 μg as the optimal nanocube amount for subsequent experiments.

188

Figure 4.5: Optimization of the pH and nanocube concentration for the peroxidase-like activity of Au-NPFe2O3NC nanocubes. Mean values of absorbance (a1, b1) and amperometric current density (a2 and b2) at designated pH (a1 and a2) of NaAc buffer solution and amount of nanocubes (b1 and b2). The concentration of TMB and H2O2 were 800 µM and 700 mM respectively. For a1 and a2, the concentration of nanocube was 5 µg. For b1 and b2, the pH of NaAc buffer solution was 3.5.

4.3.2. Steady-State Kinetic for Au-NPFe2O3NC To investigate the peroxidase-like activity of nanocubes, apparent steady-state kinetic parameters for TMB oxidation were determined by varying the concentration of H2O2 and TMB (Figure 4.6), a similar phenomenon commonly used for HRP enzymes.2 The experiments were carried out using 5 μg nanocubes in a reaction volume of 60 µL (0.2 M NaAc buffer, pH 3.5) at room temperature. The kinetic parameters were estimated by the initial rate method7, 50 The absorbance data were converted to corresponding concentration by the Beer-Lambert Law using the value of ɛ = 39,000 M-1cm-1 (at 652 nm) for the oxidized product of TMB.51 Typical Michaelis-Menten-like curve was obtained within the suitable concentration range for both

H2O2 (Figure 4.7a) and TMB (Figure 4.7b).

189

Figure 4.6: Dependence of the UV-vis absorbance (at 652 nm) of Au@NPFe2O3NC catalysed

TMB/H2O2 reaction on the concentration of (a) H2O2, (b) TMB in the range from 0.01 to 1.1 M and 0.01 to 1.0 mM respectively.

To obtain the catalytic parameters (Michaelis-Menten constant (Km) and maximum velocity (Vmax)), the data were fitted to the Michaelis-Menten- kinetic model using a nonlinear least square fitting.43 All kinetic parameters were also calculated from the Lineweaver-Burk double reciprocal plot (1/ velocity [Vo] versus 1/substrate concentration [S]) (inset of Figure 4.7),44 and compared with that for previously reported peroxidase-mimetic nanoparticles

(Table 4.1). Km is an indicator of enzyme affinity towards its substrate and a lower Km indicates the stronger affinity between enzymes and substrates. The apparent Km value for Au-

NPFe2O3NC with TMB was lower than HRP, suggesting that the nanocubes have a higher 4 affinity with TMB in compare to the HRP Moreover, Km value with both TMB and H2O2 is also higher than that of non-porous Fe3O4 NPs. The enhanced peroxidase-activity of our nanocubes at room temperature can be related to the highly porous ferric oxide moiety with high surface area (exposure of more Fe(III) ions) and large pore volume that facilitates the increased mass transfer as well as enhances the kinetics of the reaction. Additionally, as outlined earlier for PB-Fe2O3, nanoporous structure of

Au-NPFe2O3NC could be beneficial for increasing electron transfer from the top of the valence- 52 bond of Fe2O3 to the lowest unoccupied molecular orbital (LUMO) of H2O2. Moreover, transfer of lone-pairs electron density (charge transfer) from the amino group of TMB to vacant 3+ d-orbital of Fe may also enhance the electron density and mobility of Au-NPFe2O3NC nanocubes.8

190

Figure 4.7: Steady-state kinetic analyses using Michaelis-Menten model (main panel) and

Lineweaver-Burk model (inset panel) for the Au-NPFe2O3NC nanocubes by varying concentration of (a) H2O2 (0.01 to 1.1 M) and (b) TMB (0.01 to 1.0 mM) with fixed amount of

(a) TMB (800 µM) and (b) H2O2 (700 mM).

191

Table 4.1: The comparison of the kinetic parameters of Au-NPFe2O3NC and reported nanomaterials.

-8 Catalyst Substrate Km / mM Vmax / 10 Temperature References M s-1

Au-NPFe2O3NC TMB 0.0429 5.882 25 °C This work

H2O2 138.5 4.770 HRP TMB 0.434b 10 40 °C 4

H2O2 3.70 8.71 4 Fe3O4 TMB 0.098 3.44 40 °C

H2O2 154 9.78 57 Fe3O4@Pt TMB 0.147 4.11 40 °C

H2O2 702.6 71.36 12 ZnFe2O4 MNPs TMB 0.85 13.31 40 °C

H2O2 1.66 7.74 58 Co3O4 NPs TMB 0.037 6.27 40 °C

H2O2 140.07 12.1 7 PB-Fe2O3 TMB 0.009 12.3 40 °C

H2O2 15 22.8 12 Eu2O2S TMB 0.140 6.83 40 °C

H2O2 142.34 44.88 CuZnFeS NCs TMB 2.2 39 32 °C 45

H2O2 0.07 0.56

192

4.3.3. Colourimetric and amperometric detection of autoantibody Our assay for the detection of p53-specific autoantibody using peroxidase-mimetic Au-

NPFe2O3NC nanocubes is schematically represented in Figure 4.8.

Figure 4.8: Schematic representation of the assay for the detection of tumour-associated plasma (and serum) p53 autoantibody. Extravidin modified screen-printed carbon electrode was functionalized with biotinylated p53. Serum/plasma samples containing p53-specific autoantibody was then incubated onto the electrode surface followed by the incubation with

IgG/Au-NPFe2O3NC nanocatalysts. The surface-attached Au-NPFe2O3NC nanocatalysts catalyzed the oxidation of TMB in the presence of H2O2 and produced a blue-colored complex product (naked-eye), which turned yellow after the addition of an acid to the reaction media. The level of p53 autoantibody was detected via measuring the intensity (UV-visible) and amperometric current generated by the yellow product.

Here, p53 antigen was used to selectively recognise and capture the p53-specific autoantibody present in serum and plasma samples. In this proof-of-concept assay, we have chosen p53 autoantibody, because a mutation in TP53 proteins is an early indicator of cancer and also found to be present in almost 80% of all carcinogenesis.34, 53 In response to the expression and mutation of this protein, the corresponding amount of p53-tumour antigen- associated autoantibodies (TAAb) are generated, which has been considered as a potential early diagnostic biomarker for ovarian (type-II) cancer.9 In our assay, first, neutravidin-modified

193

screen-printed carbon electrode (SPCE) was modified with biotinylated p53 antigen through standard biotin-avidin chemistry. Bovine serum albumin (BSA) was then used to block the unreacted sites to reduce non-specific bindings of biomolecules followed by the incubation of serum or plasma samples containing target p53 autoantibody. As protein has a strong affinity towards the gold surface, we functionalized Au-NPFe2O3NC with α-human IgG with nanocube 54-55 to form IgG/Au-NPFe2O3NC nanocatalysts. These nanocatalysts were then added to the electrode surface to form immunocomplex with the target p53 autoantibody. To achieve the readout signals, the electrode surface was incubated with the freshly prepared TMB/H2O2 solution. The nanocatalysts initiate the oxidation of TMB, thereby producing a blue-coloured charged-transfer complex (one electron). After the addition of stop solution, the blue coloured product was converted to a stable, electroactive yellow coloured (diimine) product. The naked- eye observation of the generation of blue and subsequent yellow colours demonstrated the presence of p53 positive autoantibody present in analytes. The intensity of the TMBOx is directly linked to the number of nanocubes as well as the amount of p53 autoantibody present onto the electrode surface. A qualitative evaluation was observed by naked-eyes. Colourimetric readout (semi-quantitative) was conducted by measuring the absorbance of diimine product at 452 nm. However, the diimine product is electroactive, which allows the further quantification of autoantibody by chronoamperometry.

To evaluate the assay specificity and functionality, we performed our assay in both p53 positive and negative samples. Diluted serum samples from commercial ELISA kit (ELISA- kit, Dianova GmbH, Germany) containing p53 autoantibody was used as a positive sample and serum samples without p53 autoantibody was taken as a negative sample (cat. no. DIA 0302I). The p53 autoantibody concentration in the diluted human plasma sample is 14 U/mL, where 1.0 U represents the p53 binding activity of 100 µL undiluted calibrator. We found that positive samples containing 7.0 U/mL (1:1 Dilution) of p53 autoantibody gave an intense blue colour, while the stoichiometric amount of negative samples remained colourless. As can be seen in Figure 4.9a, a 10-times higher absorbance (abs@452 nm) was observed for a positive sample compared to that of the negative sample (0.616 versus 0.0526). Chronoamperometric measurement of the positive sample was found to be 16-times higher than that of negative one -2 (36 versus 2.116 µA cm ) (Figure 4.9b). Control experiment without IgG/Au-NPFe2O3NC nanocatalysts gave negligible responses in both absorbance (Abs@452 nm = 0.0526) and subsequent amperometric (1.9 µA cm-2) measurements. This can be explained by the fact that in the absence of nanocatalysts, TMB (in presence of H2O2) oxidation reaction does not occur

194

thereby resulting negligible response. In another control experiment, we used only phosphate buffer instead of serum with p53 autoantibody sample. This experiment resulted in a slightly higher response than that of the control without nanocatalysts (Abs@452 nm. 0.0625 versus 0.0526, and current 3.63 versus 1.9 µA cm-2). This may be due to a level of nonspecific interaction of nanocatalysts with surface-attached p53 antigen. All these control experiments clearly demonstrated the good level of specificity of our assay towards the detection of p53 autoantibody from serum samples with negligible background response.

Figure 4.9: Mean responses of (a) absorbance and (b) steady-state amperometric current obtained for the assay with one positive (presence of p53 autoantibodies in serum) with three negative control samples (no target represents PBS instead of positive serum; negative control represents serum without p53 autoantibodies, and no secondary antibody represents the no IgG/

Au-NPFe2O3NC. Insets, corresponding photos for the naked eye evaluation and i-t curves.

To evaluate the sensitivity of our assay, a series of diluted positive samples obtained via serial dilution (1:1 to 1:80; 7 U/mL to 0.0875 U/mL) was tested. We observed an increasing trend in both the absorbance and current response with the increasing concentration of target autoantibody (Figure 4.10). This is because the higher amount of target p53 autoantibody could bound an increasing amount of nanocatalysts on the electrode surface which subsequently enhance the nanocatalysts mediated oxidation of TMB (in presence of H2O2) system. As shown in Figure 4.10a, the colourimetric responses increase linearly with the increasing concentration of serum samples, and the linear regression equation was estimated to be y = 0.04741x – 0.0226, with the correlation coefficient (r2) of 0.9684. The detection limit (LOD) was estimated by the corresponding signal-to-noise ratio of 2.5 and was found to be 0.12 U/mL for the colorimetric readout. On the other hand, the linear regression equation for amperometric

195

readout was estimated as y = 5.201x – 1.4849, with the r2 of 0.9961, confirming the relatively better sensitivity (LOD of 0.08 U/mL, Figure 5b) in comparison to the colourimetric readout. A relative standard deviation (%RSD) of three different sensors for both the colourimetric and electrochemical readouts was estimated to be <5.0%, suggesting the good reproducibility of electrode surface modification, isolation of p53 autoantibody from serum samples, incubation of nanocatalysts onto the p53 autoantibody-attached electrode surface and nanocatalysts induced signal transduction protocols. This level of LOD and reproducibility of our colourimetric and electrochemical assay over a wide range of serum concentration clearly demonstrates that the peroxidase-like activity of Au-NPFe2O3NC nanocatalysts is sensitive and specific enough to detect autoantibody in p53 positive serum samples. Moreover, LOD for both readout systems is better than that of the conventional p53-ELISA kit (0.08 versus ~0.3 U/mL). These LODs are also better than that of a recently reported Halotag-fusion protein modification based electrochemical platform (0.08 versus 0.34 U/mL).38 Therefore, our methods can detect a much lower concentration of p53 autoantibody than above-reported methods. It is also important to note that the sensitivity of our recently reported HRP-based method21 is slightly better than that of the current method (i.e., 0.02 versus 0.08 U/mL) than that of the current method. However, our current method avoids the use of expensive HRP-based reaction system.

Figure 4.10: Concentration-dependent curve for p53 autoantibody standards provided in the commercial p53 autoantibody ELISA kit. Mean responses of (a) absorbance and (b) steady- state amperometric currents obtained for the designated concentration of standard samples. Inset shows the corresponding linear regression curves.

196

We further checked the applicability of our novel platform for the analysis of complex human plasma samples. The plasma samples were obtained from women with epithelial ovarian cancer high-grade serous subtype (stage I and III, P3 and P4 respectively) and controls (P1 and P2) (benign). Ovarian cancer is one of the leading causes of cancer-related death of a woman from gynecologic malignancy.9 It has been reported that, in response to the overexpression or mutation of protein (TP53) in ovarian cancers, p53-antigen-specific autoantibodies are generated.56 TP53 mutation occurs early in high-grade ovarian cancer and strongly attendant with the p53 autoantibodies.35 Thus, the development of an early diagnosis platform for ovarian cancers via p53-autoantibody analysis can potentially decrease the disease burden as well as increase the overall survival. In this study, all clinical samples were diluted (1:100) prior to performing the assay. As projected, both colourimetric and electrochemical methods response were higher in stage III patients (P4) sample than stage I (P3) (Figure 4.11). However, the assay can also differentiate the stage I response from the two non-cancerous healthy controls. Both of these healthy controls (P1 and P2) produced negligible signal suggesting the absence of p53 autoantibody. These clinical data showed a very good inter-assay reproducibility (RSD <5%, for n = 3) for the analysis of differential expression pattern of p53 autoantibodies in different stages of ovarian cancers.

Figure 4.11: Mean responses of (a) absorbance and (b) steady-state amperometric current obtained p53-specific autoantibodies present in plasma samples obtained from patients with epithelial ovarian cancer high-grade serous subtype (EOCHGS, P4 = stage III and P3 = stage I) and their non-cancerous healthy patients (benign, P1 and 2).

197

The naked-eye discrimination of the autoantibodies described here holds huge potential for the development of a user-friendly and inexpensive bioassay in resource-limited settings, where sophisticated scientific equipment is unavailable. In particular, this approach can be exploited as a rapid first-pass screening (yes/no) tool to detect clinically relevant autoantibodies in large population followed by more accurate and sensitive quantification of autoantibody via amperometric readout. The use of disposable SPE-Au successfully eliminates the need for a time-consuming electrode cleaning process typically used in conventional disk electrodes. The assay also replaces natural enzymes for TMB oxidation, and thus reduces the cost, handling and storage facilities generally required for natural enzymes. Overall, the assay platform is relatively inexpensive and portable (i.e., use of disposable and inexpensive SPE; replaces of relatively expensive HRP; avoids tedious cleaning of conventional disk electrodes). Importantly, the application of this method is not limited to autoantibody detection, it could potentially be applied as an ideal alternative for conventional ELISA assays for the detection of many other clinically relevant protein biomarkers by changing the relevant antibodies in the antibody functionalisation steps of the assay. Taken together these benefits, we expect that this peroxidase-like activity of Au-NPFe2O3NC nanocubes and their subsequent translation to p53 autoantibody detection assay may have wide application in human cancers or chronic disease.

4.4. Conclusions

We have introduced the peroxidase mimetics of a new class of Au-NPFe2O3NC nanocubes.

We have shown that Au-NPFe2O3NC resulted in the enhanced peroxidase-like activity and followed the Michaelis-Menten and Lineweaver-Burk models for the enzyme catalysed

TMB/H2O2 reaction at room temperature (25 °C). The enhanced peroxidase-like activity was mainly due to the large surface of the Au-NPFe2O3NC nanocubes (i.e., highly porous) that facilitated the binding of an increased amount of positively charged TMB and hence the

TMB/H2O2 reaction. The presence of 2% AuNPs within the nanocube framework also contributed towards the catalysis of the TMB/H2O2 reaction. This intrinsic feature was further used to develop a new proof-of-concept platform for autoantibody detection body fluids samples using both colourimetric and electrochemical readout. This platform successfully detected the p53 autoantibodies in diluted serum and a small cohort of patients’ plasma samples with high sensitivity and specificity.

198

4.5. References 1. Urbanova, V.; Magro, M.; Gedanken, A.; Baratella, D.; Vianello, F.; Zboril, R., Nanocrystalline iron oxides, composites, and related materials as a platform for electrochemical, magnetic, and chemical biosensors. Chemistry of Materials 2014, 26 (23), 6653-6673. 2. Pankhurst, Q.; Thanh, N.; Jones, S.; Dobson, J., Progress in applications of magnetic nanoparticles in biomedicine. Journal of Physics D: Applied Physics 2009, 42 (22), 224001. 3. Nguyen, N.-T.; Wu, Z., Micromixers—a review. Journal of Micromechanics and Microengineering 2004, 15 (2), R1. 4. Gao, L.; Zhuang, J.; Nie, L.; Zhang, J.; Zhang, Y.; Gu, N.; Wang, T.; Feng, J.; Yang, D.; Perrett, S., Intrinsic peroxidase-like activity of ferromagnetic nanoparticles. Nature Nanotechnology 2007, 2 (9), 577. 5. Wei, H.; Wang, E., Nanomaterials with enzyme-like characteristics (nanozymes): next- generation artificial enzymes. Chemical Society Reviews 2013, 42 (14), 6060-6093. 6. Liu, Y.; Yu, F., Substrate-specific modifications on magnetic iron oxide nanoparticles as an artificial peroxidase for improving sensitivity in glucose detection. Nanotechnology 2011, 22 (14), 145704. 7. Dutta, A. K.; Maji, S. K.; Srivastava, D. N.; Mondal, A.; Biswas, P.; Paul, P.; Adhikary,

B., Peroxidase-like activity and amperometric sensing of hydrogen peroxide by Fe2O3

and Prussian Blue-modified Fe2O3 nanoparticles. Journal of Molecular Catalysis A: Chemical 2012, 360, 71-77. 8. Xing, Z.; Tian, J.; Asiri, A. M.; Qusti, A. H.; Al-Youbi, A. O.; Sun, X., Two-

dimensional hybrid mesoporous Fe2O3–graphene nanostructures: a highly active and reusable peroxidase mimetic toward rapid, highly sensitive optical detection of glucose. Biosensors and Bioelectronics 2014, 52, 452-457. 9. Pandey, P. C.; Pandey, A. K.; Chauhan, D. S., Nanocomposite of Prussian blue based sensor for l-cysteine: Synergetic effect of nanostructured gold and palladium on electrocatalysis. Electrochimica acta 2012, 74, 23-31. 10. Kim, M. I.; Kim, M. S.; Woo, M.-A.; Ye, Y.; Kang, K. S.; Lee, J.; Park, H. G., Highly efficient colorimetric detection of target cancer cells utilizing superior catalytic activity of graphene oxide–magnetic-platinum nanohybrids. Nanoscale 2014, 6 (3), 1529-1536.

199

11. Su, L.; Feng, J.; Zhou, X.; Ren, C.; Li, H.; Chen, X., Colorimetric detection of urine glucose based ZnFe2O4 magnetic nanoparticles. Analytical Chemistry 2012, 84 (13), 5753-5758. 12. Ghosh, A. B.; Saha, N.; Sarkar, A.; Dutta, A. K.; Biswas, P.; Nag, K.; Adhikary, B.,

Morphological tuning of Eu2O2S nanoparticles, manifestation of peroxidase-like activity and glucose assay use. New Journal of Chemistry 2016, 40 (2), 1595-1604. 13. Chen, W.; Chen, J.; Liu, A. L.; Wang, L. M.; Li, G. W.; Lin, X. H., Peroxidase‐like activity of cupric oxide nanoparticle. ChemCatChem 2011, 3 (7), 1151-1154. 14. Wang, G.-L.; Xu, X.-F.; Qiu, L.; Dong, Y.-M.; Li, Z.-J.; Zhang, C., Dual responsive enzyme mimicking activity of AgX (X= Cl, Br, I) nanoparticles and its application for cancer cell detection. ACS applied Materials & Interfaces 2014, 6 (9), 6434-6442. 15. Kim, M.; Kim, M. S.; Kweon, S. H.; Jeong, S.; Kang, M. H.; Kim, M. I.; Lee, J.; Doh, J., Simple and Sensitive Point‐of‐Care Bioassay System Based on Hierarchically Structured Enzyme‐Mimetic Nanoparticles. Advanced Healthcare Materials 2015, 4 (9), 1311-1316. 16. Song, Y.; Wang, X.; Zhao, C.; Qu, K.; Ren, J.; Qu, X., Label‐free colorimetric detection of single nucleotide polymorphism by using single‐walled carbon nanotube intrinsic peroxidase‐like activity. Chemistry–A European Journal 2010, 16 (12), 3617-3621. 17. Cui, R.; Han, Z.; Zhu, J. J., Helical carbon nanotubes: intrinsic peroxidase catalytic activity and its application for biocatalysis and biosensing. Chemistry–A European Journal 2011, 17 (34), 9377-9384. 18. Shi, W.; Wang, Q.; Long, Y.; Cheng, Z.; Chen, S.; Zheng, H.; Huang, Y., Carbon nanodots as peroxidase mimetics and their applications to glucose detection. Chemical Communications 2011, 47 (23), 6695-6697. 19. Wang, S.; Chen, W.; Liu, A. L.; Hong, L.; Deng, H. H.; Lin, X. H., Comparison of the peroxidase‐like activity of unmodified, amino‐modified, and citrate‐capped gold nanoparticles. ChemPhysChem 2012, 13 (5), 1199-1204. 20. Zheng, T.; Zhang, Q.; Feng, S.; Zhu, J.-J.; Wang, Q.; Wang, H., Robust nonenzymatic hybrid nanoelectrocatalysts for signal amplification toward ultrasensitive electrochemical cytosensing. Journal of the American Chemical Society 2014, 136 (6), 2288-2291. 21. Yadav, S.; Masud, M. K.; Islam, M. N.; Gopalan, V.; Lam, A. K.-y.; Tanaka, S.; Nguyen, N.-T.; Al Hossain, M. S.; Li, C.; Yamauchi, M. Y., Gold-loaded nanoporous

200

iron oxide nanocubes: a novel dispersible capture agent for tumor-associated autoantibody analysis in serum. Nanoscale 2017, 9 (25), 8805-8814. 22. Masud, M. K.; Islam, M. N.; Haque, M. H.; Tanaka, S.; Gopalan, V.; Alici, G.; Nguyen, N.-T.; Lam, A. K.; Hossain, M. S. A.; Yamauchi, Y., Gold-loaded nanoporous superparamagnetic nanocubes for catalytic signal amplification in detecting miRNA. Chemical Communications 2017, 53 (58), 8231-8234. 23. Koo, K. M.; Sina, A. A.; Carrascosa, L. G.; Shiddiky, M. J.; Trau, M., DNA–bare gold affinity interactions: mechanism and applications in biosensing. Analytical Methods 2015, 7 (17), 7042-7054. 24. Islam, M. N.; Gopalan, V.; Haque, M. H.; Masud, M. K.; Al Hossain, M. S.; Yamauchi, Y.; Nguyen, N.-T.; Lam, A. K.-Y.; Shiddiky, M. J., A PCR-free electrochemical method for messenger RNA detection in cancer tissue samples. Biosensors and Bioelectronics 2017, 98, 227-233. 25. Haque, M. H.; Gopalan, V.; Islam, M. N.; Masud, M. K.; Bhattacharjee, R.; Al Hossain, M. S.; Nguyen, N.-T.; Lam, A. K.; Shiddiky, M. J., Quantification of gene-specific DNA methylation in oesophageal cancer via electrochemistry. Analytica Chimica Acta 2017, 976, 84-93. 26. Hossain, T.; Mahmudunnabi, G.; Masud, M. K.; Islam, M. N.; Ooi, L.; Konstantinov, K.; Al Hossain, M. S.; Martinac, B.; Alici, G.; Nguyen, N.-T., Electrochemical biosensing strategies for DNA methylation analysis. Biosensors and Bioelectronics 2017, 94, 63-73. 27. Pedersen, J.; Gentry-Maharaj, A.; Fourkala, E.; Dawnay, A.; Burnell, M.; Zaikin, A.; Pedersen, A. E.; Jacobs, I.; Menon, U.; Wandall, H., Early detection of cancer in the general population: a blinded case–control study of p53 autoantibodies in colorectal cancer. British Journal of Cancer 2013, 108 (1), 107. 28. Barderas, R.; Villar-Vázquez, R.; Fernández-Aceñero, M. J.; Babel, I.; Peláez-García, A.; Torres, S.; Casal, J. I., Sporadic colon cancer murine models demonstrate the value of autoantibody detection for preclinical cancer diagnosis. Scientific Reports 2013, 3, 2938. 29. Cho-Chung, Y. S., Autoantibody biomarkers in the detection of cancer. Biochimica et Biophysica Acta (BBA)-Molecular Basis of Disease 2006, 1762 (6), 587-591. 30. Chapman, C.; Murray, A.; Chakrabarti, J.; Thorpe, A.; Woolston, C.; Sahin, U.; Barnes, A.; Robertson, J., Autoantibodies in breast cancer: their use as an aid to early diagnosis. Annals of oncology 2007, 18 (5), 868-873. 201

31. Soler, M.; Estevez, M.-C.; Villar-Vazquez, R.; Casal, J. I.; Lechuga, L. M., Label-free nanoplasmonic sensing of tumor-associate autoantibodies for early diagnosis of colorectal cancer. Analytica Chimica Acta 2016, 930, 31-38. 32. Anderson, K. S.; Cramer, D. W.; Sibani, S.; Wallstrom, G.; Wong, J.; Park, J.; Qiu, J.; Vitonis, A.; LaBaer, J., Autoantibody signature for the serologic detection of ovarian cancer. Journal of Proteome Research 2014, 14 (1), 578-586. 33. Qiu, J.; Choi, G.; Li, L.; Wang, H.; Pitteri, S. J.; Pereira-Faca, S. R.; Krasnoselsky, A. L.; Randolph, T. W.; Omenn, G. S.; Edelstein, C., Occurrence of autoantibodies to annexin I, 14-3-3 theta and LAMR1 in prediagnostic lung cancer sera. Journal of Clinical Oncology 2008, 26 (31), 5060. 34. Soussi, T., p53 Antibodies in the sera of patients with various types of cancer: a review. Cancer Research 2000, 60 (7), 1777-1788. 35. Katchman, B. A.; Chowell, D.; Wallstrom, G.; Vitonis, A. F.; LaBaer, J.; Cramer, D. W.; Anderson, K. S., Autoantibody biomarkers for the detection of serous ovarian cancer. Gynecologic Oncology 2017, 146 (1), 129-136. 36. Ramachandran, S.; Fu, E.; Lutz, B.; Yager, P., Long-term dry storage of an enzyme- based reagent system for ELISA in point-of-care devices. Analyst 2014, 139 (6), 1456- 1462. 37. Yan, G.; Xing, D.; Tan, S.; Chen, Q., Rapid and sensitive immunomagnetic- electrochemiluminescent detection of p53 antibodies in human serum. Journal of Immunological Methods 2004, 288 (1-2), 47-54. 38. Garranzo-Asensio, M.; Guzman-Aranguez, A.; Povés, C.; Fernández-Aceñero, M. a. J. s.; Torrente-Rodríguez, R. M.; Ruiz-Valdepeñas Montiel, V. c.; Domínguez, G.; Frutos, L. S.; Rodríguez, N.; Villalba, M., Toward liquid biopsy: determination of the humoral immune response in cancer patients using halotag fusion protein-modified electrochemical bioplatforms. Analytical Chemistry 2016, 88 (24), 12339-12345. 39. Bard, A. J., Faulkner, L. R., Electrochemical Methods. Fundamentais and Applications John Wiley and Sons 1980. 40. Shiddiky, M. J.; Torriero, A. A.; Zhao, C.; Burgar, I.; Kennedy, G.; Bond, A. M., Nonadditivity of faradaic currents and modification of capacitance currents in the voltammetry of mixtures of ferrocene and the cobaltocenium cation in protic and aprotic ionic liquids. Journal of the American Chemical Society 2009, 131 (23), 7976- 7989.

202

4-/3- 41. Winkler, K., The kinetics of electron transfer in Fe(CN)6 redox system on platinum standard-size and ultramicroelectrodes. Journal of Electroanalytical Chemistry 1995, 388 (1-2), 151-159. 42. Das, J.; Aziz, M. A.; Yang, H., A nanocatalyst-based assay for proteins: DNA-free ultrasensitive electrochemical detection using catalytic reduction of p-nitrophenol by gold-nanoparticle labels. Journal of the American Chemical Society 2006, 128 (50), 16022-16023. 43. Lehninger, A. L.; Nelson, D. L.; Cox, M. M.; Cox, M. M., Lehninger principles of biochemistry. Macmillan: 2005. 44. Lineweaver, H.; Burk, D., The determination of enzyme dissociation constants. Journal of the American chemical society 1934, 56 (3), 658-666. 45. Josephy, P. D.; Eling, T.; Mason, R. P., The horseradish peroxidase-catalyzed oxidation of 3, 5, 3', 5'-tetramethylbenzidine. Free radical and charge-transfer complex intermediates. Journal of Biological Chemistry 1982, 257 (7), 3669-3675. 46. Mansour, A. M., Photocatalytic degradation of methylene blue with hematite nanoparticles synthesized by thermal decomposition of fluoroquinolones oxalato–iron (III) complexes. RSC Advances 2015, 5 (76), 62052-62061. 47. Chen, Z.; Yin, J.-J.; Zhou, Y.-T.; Zhang, Y.; Song, L.; Song, M.; Hu, S.; Gu, N., Dual enzyme-like activities of iron oxide nanoparticles and their implication for diminishing cytotoxicity. Acs Nano 2012, 6 (5), 4001-4012. 48. Dalui, A.; Pradhan, B.; Thupakula, U.; Khan, A. H.; Kumar, G. S.; Ghosh, T.; Satpati, B.; Acharya, S., Insight into the mechanism revealing the peroxidase mimetic catalytic activity of quaternary CuZnFeS nanocrystals: colorimetric biosensing of hydrogen peroxide and glucose. Nanoscale 2015, 7 (19), 9062-9074. 49. Gurol, M. D.; Lin, S.-S., Hydrogen peroxide/iron oxide-induced catalytic oxidation of organic compounds. Journal of Advanced Oxidation Technologies 2002, 5 (2), 147- 154.

50. Wei, H.; Wang, E., Fe3O4 magnetic nanoparticles as peroxidase mimetics and their

applications in H2O2 and glucose detection. Analytical Chemistry 2008, 80 (6), 2250- 2254. 51. Karaseva, E.; Losev, Y. P.; Metelitsa, D., Peroxidase-catalyzed Oxidation of 3, 3", 5, 5"-Tetramethylbenzidine in the Presence of 2, 4-Dinitrosoresorcinol and Polydisulfide Derivatives of Resorcinol and 2, 4-Dinitrosoresorcinol. Russian Journal of Bioorganic Chemistry 2002, 28 (2), 128-135. 203

52. Show, B.; Mukherjee, N.; Mondal, A., α-Fe2O3 nanospheres: facile synthesis and highly efficient photo-degradation of organic dyes and surface activation by nano-Pt for enhanced methanol sensing. RSC Advances 2016, 6 (79), 75347-75358. 53. Tsai-Turton, M.; Santillan, A.; Lu, D.; Bristow, R. E.; Chan, K. C.; Shih, I.-M.; Roden, R. B., p53 autoantibodies, cytokine levels and ovarian carcinogenesis. Gynecologic Oncology 2009, 114 (1), 12-17. 54. Lacerda, S. H. D. P.; Park, J. J.; Meuse, C.; Pristinski, D.; Becker, M. L.; Karim, A.; Douglas, J. F., Interaction of gold nanoparticles with common human blood proteins. ACS Nano 2009, 4 (1), 365-379. 55. Yadav, S.; Carrascosa, L. G.; Sina, A. A.; Shiddiky, M. J.; Hill, M. M.; Trau, M., Electrochemical detection of protein glycosylation using lectin and protein–gold affinity interactions. Analyst 2016, 141 (8), 2356-2361. 56. Anderson, K. S.; Wong, J.; Vitonis, A.; Crum, C. P.; Sluss, P. M.; LaBaer, J.; Cramer, D., p53 autoantibodies as potential detection and prognostic biomarkers in serous ovarian cancer. Cancer Epidemiology and Prevention Biomarkers 2010, 19 (3), 859- 868.

57. Ma, M.; Xie, J.; Zhang, Y.; Chen, Z.; Gu, N., Fe3O4@Pt nanoparticles with enhanced peroxidase-like catalytic activity. Materials Letters 2013, 105, 36-39.

58. Mu, J.; Wang, Y.; Zhao, M.; Zhang, L., Intrinsic peroxidase-like activity and catalase-

like activity of Co3O4 nanoparticles. Chemical Communications 2012, 48, 2540-2542.

204

Chapter 5

Gold-loaded nanoporous superparamagnetic nanocubes for catalytic signal amplification in detecting miRNA*

*Sections of this chapter are based on Masud, M.K., et al., Chemical Communications 2017, 53 (58), 8231-8234.

205

5. Gold-loaded nanoporous superparamagnetic nanocubes for catalytic signal amplification in detecting miRNA

5.1. Introduction Nanostructured magnetic nanoparticles (NPs) have attracted an immense interest for a broad range of applications in the areas of catalysis, nanotechnology and biotechnology.1 They have unique magnetic characteristics (e.g., superparamagnetism, high values of saturation magnetization, easy control by small magnetic fields), biochemical characteristics (e.g., nontoxicity, biodegradability, biocompatibility), intrinsic enzyme mimicking activity, low cost of synthesis, and ability to catalyse redox reaction of various organic and inorganic compounds.2 Introducing pores into the NPs offer a high surface area relative to volume that makes it highly capable for uptaking and releasing biological guest molecules.3-4 This porous metal structure possesses enhanced catalytic capacity as they maximise surface dependent mass transport as compared to that of bulk materials of the same mass. Besides, porous NPs offer several advantages in catalysis including stabilization of particles from sintering, an expedition of cascade reaction by placing catalytic functionality in sequential compartments, and enhancement of the selectivity of catalysis by molecular sieving. Composite nanomaterials also attract increasing attention because of their combined physicochemical properties and potential for catalysis and biosensing applications.5 They have more superior characteristics than the monometallic frameworks counterparts. For instance, iron oxides (Fe3O4 or γ-Fe2O3) containing gold nanoparticle (AuNPs) exhibits combined advantages and serendipitous 6-8 properties of both Fe3O4 and AuNPs. The combination of high surface area, conductivity, thermal/chemical stability and superparamagnetism of Fe2O3 with bio-favourable physicochemical properties of AuNPs (i.e. affinity interaction of DNA/RNA with gold) makes gold-loaded nanoporous Fe2O3 nanocubes (Au@NPFe2O3NC) extremely suitable for developing biosensors for a wide range for molecular biomarkers including micro RNA (miRNA). miRNAs are small non-coding RNAs with 21-24 nucleotides which can regulate gene expressions by modulating its downstream proteins. Recent studies have also confirmed the use of miRNAs as biomarkers for diagnosis and prognosis of many diseases including cancer.9- 10 Over the past decades, many conventional approaches such as northern blotting, microarrays, in-situ hybridisation and quantitative real-time PCR (qRT-PCR) have been used for the analysis of miRNA. Each of these methods has its own advantages and disadvantages. For examples, qRT-PCR can give a quantification of miRNA but has a very low throughput.

206

Additionally, most of these methods use PCR amplification, which are still affected by amplification bias, fluorescent labelling and rely on complicated and expensive protocols.11 As an alternative to these methods, a large number of biosensors have been developed based on hybridization, oligonucleotides labelled with enzymes and redox indicator.12-14 Despite the superior analytical performance, these sensors often involve complex and tedious amplification processes, expensive biomaterials, time-consuming and laborious procedures.15 Therefore, development of a simple, inexpensive and rapid miRNA biosensor would be highly beneficial for miRNA-based molecular diagnostics of patients with chronic diseases.

In this study, we synthesised and characterized a new class of Au@NPFe2O3NC. The electrocatalytic activity of this nanocube was used to develop a miRNA biosensor. The sensor was tested in cell lines and tissue samples obtained from patients with oesophageal squamous cell carcinoma (ESSC).

5.2. Experimental

5.2.1. Materials and Instrumentations Unless otherwise stated, the reagents and chemicals used for the conducting experiments were of analytical grade. Polyvinylpyrrrolidone (PVP) and potassium hexacyanoferrate (III) were purchased from Nacalai Tesque and Merck KGaA, Germany respectively. Reagent grade hexaammineruthenium(III) chloride (RuHex), phosphate buffer saline (PBS) tablet (0.01M phosphate buffer, 0.0027M potassium chloride and 0.137M sodium chloride, pH 7.4 at 25oC) were purchased from Sigma-Aldrich (Australia). Analytical grade hydrochloric acid (HCl) was purchased from Chem-supply (Australia). Tris was obtained from VWR Life science (Australia), glassy carbon electrode (GCE) was purchased from CH instrument (USA). Screen- printed carbon electrode (SPCE) with a three-electrode system printed on a ceramic substrate (length 34 × width 10 × height 5 mm) (DRP-150) from Dropsens (Spain). In the three-electrode system, working (4 mm diameter), counter and reference electrodes were carbon, platinum and silver-modified. All chemicals and reagents were used as received without additional purification. UltrapureTM DNase/RNase-free distilled water (Invitrogen, Australia) was used throughout the experiments. Oligonucleotides were acquired from Integrated Technologies, USA and sequences are shown in table S1. Scanning electron microscope (SEM) images were taken with a Hitachi S-4800 scanning microscope with the accelerating voltage of 10 kV. Wide-angle powder X-ray diffraction (XRD) patterns were obtained with a Rigaku RINT 2500X diffractometer using

207

monochromated Cu Kα radiation (40 kV, 40 mA) at a scanning rate of 0.5 ° min−1. The elemental chemical analysis of the nanocubes was performed by X-ray photoelectron spectroscopy (XPS, PHI Quantera SXM, ULVAC-PHI Inc., Japan). All samples were degassed in a vacuum before carrying out the measurements. All electrochemical measurements were performed with a CHI650 electrochemical workstation. (CH Instrument, USA). Cyclic voltammetry (CV) and chronoamperometry experiments were done in a single-compartment cell with a 3-mL volume. A conventional three-electrode system, comprising a bare or modified

GCE, a platinum auxiliary electrode, and an Ag/AgCl3 1.0 M NaCl reference electrode (CH Instrument, Inc. USA), was used for the measurement of electrocatalytic activity. Chronocoulometry (CC) measurements were carried out using 80 µL volume on SPCE between 0 and -500mV, 25 ms pulse width and 2 ms sample interval. A temperature and time control ultrasonic water bath (Soniclean, Australia) was applied for the dispersion of Au@NPFe2O3NC nano-hybrid before applying to the electrode surface.

Table 5.1: Oligonucleotide sequences (miR-21) Oligos 5 ́-Sequences-3 ́ Biotinylated miR-21 capture TGA CCG ACC CAG TGA GGA AGT TTT CTC T/ 3Bio probe Synthetic miR-21 sequence AGA GAA AAC UUC ACU GGG UCG GUC A

5.2.2. Electrochemical measurement of catalytic activity A GCE was polished using 0.3 and 0.05 mm alumina slurry (CH Instrument, Inc. USA) followed by rinsing with an adequate amount of water. After successive sonication with nitric acid and water, the electrode was again rinsed thoroughly using DI water, allowed it to dry at room temperature. A mirror surface was formed. To assess the electrocatalytic activity of

Au@NPFe2O3NC, 5μg of a colloidal suspension of Au@NPFe2O3NC were drop-dried onto the surface of the clean GCE electrode. The electrocatalytic activity Au@NPFe2O3NC towards the reduction of RuHex was studied using cyclic voltammetric technique at room temperature with the conventional three-electrode system using Au@NPFe2O3NC-modified GCE as the working electrode. The chronoamperometric experiment was also carried at -0.25V versus Ag/AgCl at optimum condition. The current response due to the successive addition of RuHex (10 – 1100

208

app μM) was monitored. The apparent Michaelis-Menten constant (Km ) of 16 GCE/Au@NPFe2O3NC can be determined from the Michaelis-Menten equation;

I max [S] I  app … … … (1) K m  [S]

In this equation, I is the steady-state current, Imax is the maximum current measured under the app condition of enzyme saturation, [S] is the substrate concentration, and Km is the Michaelis- Menten constant, which is equivalent to the substrate concentration at the conversion rate is app app half of Imax. Km is the indicator of enzyme or catalyst affinity to substrates. A high Km indicates week affinity while a low value suggests a high affinity. The rearrangement of the Michaelis-Menten equation gives the electrochemical version of Lineweaver–Burk equation,17 app which also widely used to determine electrocatalytic enzyme kinetics terms Km and Imax.

1 K app 1 1  m  … … … (2) I I max [S] I max

5.2.3. Determination of the surface area of the electrodes The effective surface areas of both GCE and SPCE were determined by the measurement of the peak current obtained as a function of scan rate under cyclic voltammetric conditions for 3- the one-electron reduction of [Fe(CN)6] [2.0 mM in PBS (0.5 M KCl)] and by using the Randles-Sevcik equation (eq 3),18-19

5 3/ 2 1/ 2 1/ 2 ip  (2.6910 ) n AD C ...... (3)

3+ 2+ where, ip is the peak current (A), n is the number of electrons transferred (Fe → Fe , n = 1), A is the effective area of the electrode (cm2), D is the diffusion coefficient of [Fe(CN)6]3- (taken to be 7.60 × 10-5cm2s-1), C is the concentration (mol cm-3), ν is the scan rate (Vs-1).

5.2.4. Preparation of the miRNA recognition interface Before the adsorption of miRNA, SPCE was washed and cleaned by rinsing with an excess amount of miliQ water. To attach Au@NPFe2O3NC to SPCE surface, the electrode was positioned on a permanent magnet so that the surface is centred to the magnet and 5µg of

Au@NPFe2O3NC was employed onto the electrode surface (Figure 5.11). The

Au@NPFe2O3NC was allowed to attach onto the surface for 45 min. The electrode was then washed with 10 mM PBS to remove unattached or loosely attached particles from the electrode surface.

209

5.2.5. RNA Extraction from cell lines and tissue samples Two oesophageal squamous cell carcinoma cell lines (HKESC-1 and HKESC-4) were used for this study. These cells were cultured in minimum essential medium alpha (MEMα growth medium, Gibco (ThermFisher scientific, Waltham, MA, USA) medium with non-essential amino acids and supplemented with 10% fetal bovine serum (FBS, Gibco), 100 µg/mL penicillin (Gibco) and 100 units/mL streptomycin (Gibco) in a humidified cell culture incubator containing 5% CO2 at 37 °C. Four matched fresh oesophageal squamous cell carcinoma (ESCC) and non-neoplastic tissues were snap-frozen and sectioned into 10µm slices using a cryostat (Leica CM 1850 UV, Wetzlar, Germany). Tissue sections were stained with haematoxylin and eosin for RNA extraction. Ethical approval was obtained from the Griffith University Human research ethics committee for the use of ESCC tissues (GU Ref Nos: MED/19/08/HREC). Total RNA was isolated and purified from all tissue samples following the manufacturer’s recommendations of all prep DNA/RNA mini kit (Qiagen, Hilden, NRW, Germany). Briefly, tissue samples were suspended in 0.01M PBS following a digestion step to remove the debris, protein and DNA in the solution via proteinase and DNase enzymes, respectively. The digested proteins and DNA were then removed by the centrifugation of the solution in a spin column. The purified RNA was eluted from the column using 100 µL of elution buffer. To evaluate the quality and quantity of RNA, agarose gel (1.5%) electrophoresis and Nanodrop spectrophotometric analysis (BioLab, Ipswich, MA, USA) using 260:280 ratio was performed. The concentration of RNA was noted in ng/µl and stored at -80OC until assayed.

5.2.6. Isolation of target miRNA Target miRNA was captured by hybridizing with magnetic beads functionalized complementary capture probe followed by magnetic isolation and heat release of miRNA in accordance with our previous method.20 Briefly, 10µL of commercial streptavidin-labelled magnetic beads (Dynabeads® MyOne™ Streptavidin C1, Invitrogen, Australia) was washed with binding and washing (B and W) solution followed by 20 min incubation with an equal volume of 10 µM biotinylated capture probes. The functionalized beads were washed and re-suspended in the 10 µL 5x SSC (saline sodium citrate) buffer. 10µL of target miRNA (pre-adjusted 50 ng total RNA from tissue samples to 10µL with RNase-free water or various concentration of synthetic miRNA) was then mixed with beads functionalized capture probe. After 20 min of incubation and washing with the (B and W) solution, the miRNA attached 210

beads were isolated with an external magnet and resuspended in 9 µL of RNase free water. The resuspended miRNA mixture was heated for 2 min at 95 oC, immediately attached the beads with magnet and supernatant containing the desired miRNA was collected. Before applying miRNA on to Au@NPFe2O3NC-modified electrode, the miRNA was diluted two times with 5XSSC buffer.

5.2.7. Electrochemical detection of adsorbed microRNA

For detecting miRNA, 5µg of Au@NPFe2O3NC was added on the SPCE. The electrode was then positioned on a permanent magnet (Figure 5.11). 4 μL of target miRNA sample was then incubated onto the magnetically attached Au@NPFe2O3NC/SPCE surface for 30 minutes followed by PBS washing. The electrode was then incubated with 7 μL of 50 μM Ruhex so that positively charged Ru3+ can bound with the negatively charged phosphate backbone of adsorbed miRNA. The electrode was then washed with PBS before performing CC measurement. CC measurement was then carried out in 10mM Tris buffer (pH 7.4, 80 μL) with a potential step of 5 mV and a pulse width of 250 ms, and a sample interval of 2 msec. Using

CC, the amount of miRNA adsorbed onto the Au@NPFe2O3NC/SPCE surface was then calculated from the number of cationic redox molecules (RuHex) electrostatically associated with the surface-attached anionic phosphate backbone of miRNA. The total charge Q at a time t can be expressed by the integrated Cottrell equation.21-22

1/ 2 * 2nFA C 1/ 2 Q  0 0 t  Q  nFA … … … (4)  1/ 2 dl 0 where, n is the number of electrons involved in electrode reaction, F is Faraday constant 2 2 (C/equivalent), A is the electrode area (cm ), Do is the diffusion coefficient (cm /s), Co* is the

2 bulk concentration (mol/cm ), 0 is representing the amount of RuHex-confined near electrode surface and nFA 0 (kwon as surface excess) is the charge obtained by adsorbed miRNA. CC curves were constructed by plotting the charge flowing through the RNA-attached electrode 1/2 1/2 versus square-root of time (t /s ) in the presence and absence RuHex. Q and Qdl were estimated from the intercept of these two curves at t = 0. Therefore, Q represents the total charge comprising both Faradic and non-Faradic (capacitive) charges. Hence, the corresponding charge of RuHex (electrostatically bound to surface-confined RNA) can be calculated as,

QRNA  Q  Qdl … … … (5)

211

And the saturated surface density of RuHex could be used to calculate surface attached miRNA using the following equation;

RNA  QRNAN A / nFAz / m … … … (6) where, n is the number of electrons involved in the reaction (n = 1), A is the working electrode area, NA is the Avogadro’s number, m is the number of nucleotides in the RNA, and z is the charge of redox molecules (for RuHex, z = 3). For Figure 5.12B, using the equation (5) and (6), the surface density of miR-21 on the electrode surface was calculated to be 8.52 × 1013 and 1.6 × 1013 moleculescm-2 for 1.0 µM and 100 fM of miRNA respectively.

5.2.8. Quantitative reverse-transcription polymerase chain reaction (RT-qPCR) cDNA conversion was first carried out using miScript Reverse Transcription kit (Qiagen, Germany) as previously described.23-24 Each cDNA sample of 30 ng/µL was allocated and stored at -20 ºC for RT-qPCR analysis. The level miR-21 expression was amplified using the primers: forward, 5′- CGGCGGTAGCTTATCAGACTGA-3′ and reverse, 5′- GTGCAGGGTCCGAGGT-3′. Primer pairs were purchased from Integrated DNA Technologies (USA). RT-qPCR was achieved in a total volume of 10 µL reaction mixture comprising 5 µL of 2xSensiMix SYBR No-ROX master mix (Bioline, UK), 1 µL of each 1 µmole/µL primer, 1 µL of cDNA at 30 ng/µL and 2 µL of Nuclease-free water. Thermal cycling was initiated with a first denaturation step at 95 °C for 10 minutes followed by 40 cycles of 95 °C for 15 seconds (denaturation), 60°C for 30 seconds (annealing), and 72°C for 30 seconds (extension). Expression levels were normalized against the endogenous U6 control gene, which was amplified in the same run and following the same procedure described above. The sequences of the U6 primers were as follows: forward, 5'- GCTTCGGCAGCACATATACTAAAAT-3' and reverse, 5'- GTGCAGGGTCCGAGGT-3'. Assays were accomplished in triplicate to verify the results and a non-template control was included in all the experiment. Data analysis of miR-21 expression was performed as previously reported.23

212

5.3. Result and Discussion

5.3.1. Synthesis and Characterization of Au@NPFe2O3NC

Porous iron oxide nanocubes (NPFe2O3NC) were prepared from Prussian blue (PB) nanocubes (NC) via calcination of PB at 250 °C following our previous report.4, 25 Concisely, 6.0 g of PVP

(polyvinylpyrrolidone) (K30) and 264 mg of K3[Fe(CN)6].3H2O were dissolved in 80 mL 0.01 M HCl solution followed by the magnetic stirring for 30 min to produce a clear yellow solution. The obtained solution was then heated at 80 °C for 30 h in an electronic furnace, and the produced precipitates were collected by centrifugation. PB NCs of 80 nm sized were obtained after 24 hr hours drying at room temperature. The obtained PB NCs are in a cubic shape which is a typical PB material (Figure 5.1A). The prepared PB NCs show the same face-centred cubic diffraction patterns that of the bulk PB crystals (JCPDS card 73-0687) (Figure 5.1C). The prepared PB NCs was highly pure as there were no peaks derived from impurities in XRD pattern. In the parent PB crystals, iron atoms are separated by cyano-bridges, which facilitates the favourable conditions for Fe2O3 particles during the calcination process. To prepare nanoporous iron oxide, 50 mg of obtained PB powders was taken in a melting pot and heated in an electronic furnace to achieve complete thermal decomposition. The powders were then allowed to cool inside the furnace. The morphology of the prepared PB-derived iron oxide remains as NCs, but their sizes were slightly reduced and surface roughness was increased (Figure 5.1B).

Figure 5.1: (A-B) SEM images of (A) PB nanocubes and (B) the calcined PB nanocubes. (C- D) Wide-angle XRD patterns of (C) PB nanocubes and (D) Au-loaded nanoporous iron oxide nanocubes.

213

For loading AuNPs on to the porous nanocube, 250 mg of iron oxide nanocubes were dispersed in sodium citrate solution followed by the addition of 3 mL of 10 mM HAuCl4 solution. The mixed solution was then incubated under ice-water bath until its temperature was stable. Then, sodium borohydride solution as a reducing agent was quickly added into the above solution under vigorous stirring. After reacting for 10 min, the product was washed and collected by successive centrifugation. After deposition of Au NPs, uniformly sized Au NPs are distributed on the surface of nanoporous iron oxide NCs (Figure 5.2A). The loading amount of Au nanoparticles is around 2 wt% in the product (Au@NPFe2O3NC) (Figure 5.2B). The

XRD pattern shows the diffraction peaks derived from Au, α-Fe2O3, and γ-Fe2O3 (Figure 5.1D).

Figure 5.2: (A) Elemental mapping images (O, Fe, and Au), and (B) EDX spectrum of

Au@NPFe2O3NC.

This sample is found to be superparamagnetic from the complete reversibility of the M- H curve recorded at room temperature (300 K). The S-shaped hysteresis loops are shown in Figure 5.3 with the negligible coercive field (Hc) are a typical characteristic of superparamagnetic nanoparticles.26-28 The saturation magnetisation (Ms) reported in this study -1 is 16 emu g at 300 K for Au@NPFe2O3NC. Due to this sufficient Ms value, the samples can be easily collected by a neodymium magnet. With the loading amount of Au nanoparticles, the Ms value is decreased because of the non-magnetic property of Au.

214

Figure 5.3: Magnetization curve measured at 300 K for Au@NPFe2O3NC

5.3.2 Electrocatalytic Activity of Au@NPFe2O3NC

To assess the electrocatalytic activity of Au@NPFe2O3NC, the Au@NPFe2O3NC-modified glassy carbon electrode (GCE) was used as a working electrode. The cyclic voltammetric (CV) measurements were carried out in the presence of Ru(NH3)6Cl3 (RuHex), a redox label widely 29 used in electrochemical biosensing. Well-defined cathodic (Epc) and anodic (Epa) peaks for 3+/2+ the [Ru(NH3)6] sytem were obtained at -225 mV and -165 mV (vs Ag/AgCl) at the unmodified GCE (GCE/bare), indicating one-electron reversible process (E = 60 mV, Figure

5.4A). The Au@NPFe2O3NC-modified GCE (GCE/Au@NPFe2O3NC) offered significantly enhanced cathodic (ipc), and anodic (ipa) peak currents compared to that of the GCE/bare -2 electrode (Figure 5.4B). Notably, ipc increased approximately four-times (4.6 vs 20.1 µAcm )

- with Epc shifted by  -55 mV, whereas ipa increased approximately two-times (2.3 vs 5.2 µAcm

2 ) with Epa shifted by  -30 mV. These data indicate that Au@NPFe2O3NC catalysed both the oxidation and reduction of RuHex, while the rate of the reduction of RuHex is greater than that of the oxidation process.

215

Figure 5.4: (A) CVs obtained at an unmodified GCE (top) and Au@NPFe2O3NC- modified GCE (bottom) in 50µM RuHex (scan rate, 50 mVs-1). (B) Comparison of these two CVs.

The effect of the pH and temperature of the electrolyte for the electrocatalytic activity of Au@NPFe2O3NC was also studied. An enhanced catalytic response was found in acidic pH and at a higher temperature (Figure 5.5 and 5.6). However, due to the physiological conditions, pH 7.0 and room temperature (~25 oC) were selected as optimal conditions for all subsequent experiments.

Figure 5.5: Cyclic voltammograms at GCE/Bare (left) and GCE/Au@NPFe2O3NC (right) at designated pH from 3 to 11 in the presence of 50 µM RuHex (0.01M PBS, pH-7, scan rate 50 mVs-1).

216

o o o Figure 5.6: Cathodic peak currents of GCE/Au@NPFe2O3NC at 25 C, 37 C and 50 C in the presence of 50 µM RuHex (0.01 M PBS, pH-7, scan rate = 50 mVs-1). Inset, corresponding cyclic voltammogram. To examine the charge transport mechanism, we recorded CVs of both GCE/bare and -1 GCE/Au@NPFe2O3NC as a function of scan rates (10 – 1500 mVs ). As shown in Figure -1 5.7A, the currents ipc and ipa increase with increasing scan rate from 10 to 1500 mVs , indicating that the Au@NPFe2O3NC retained its electrocatalytic activity within the studied scan rates. Figure 5.7B shows a linear relationship between ipc and ipa with the square root of 1/2 the scan rate (ν ) for both the unmodified and Au@NPFe2O3NC-modified GCE, suggesting the electrocatalytic redox reactions of ReHex at the GCE/Au@NPFe2O3NC electrode occurred

18 1/2 mainly through the diffusion-controlled process. Notably, the curve of ipc and ipa versus v for the GCE/Au@NPFe2O3NC electrode showed a steeper slope than that of the unmodified

GCE, further verifying the catalytic activity of Au@NPFe2O3NC towards the redox reaction of RuHex (Figure 5.7B).

217

Figure 5.7: (A) Cyclic voltammograms obtained at GCE/Bare (top, left) and

GCE/Au@NPFe2O3NC (top, right) electrodes at different scan rate (50µM RuHex, 0.01M 1/2 PBS, pH 7.0). (B) Corresponding curves for ipc and ipa (current density) as a function of ν .

We also found that the catalytic ipc of RuHex at the GCE/Au@NPFe2O3NC electrode increases with increasing RuHex concentration (Figure 5.8). To further understand the electrocatalytic activity, chronoamperometric (CA) responses were recorded at the

GCE/Au@NPFe2O3NC electrode upon the successive addition of RuHex (Figure 5.9), where the CA response first increased steeply and then moved along the saturation. The calibration curve (Figure 5.9B) follows the typical Michaelis-Menten equation for enzyme catalysis.16 The app apparent Michaelis-Menten constant (Km ) can be obtained from the electrochemical version of Lineweaver-Burk model (inset of Figure 5.9B), and it was estimated to be 0.539 mM. This value is significantly low, suggesting the higher affinity of Au@NPFe2O3NC to RuHex, further 17 verifying electrocatalytic activity of Au@NPFe2O3NC towards RuHex. The electrocatalytic activity of this materials could be related to its smaller size, enhanced surface area, and superior 2, 5 electron transfer ability of porous Fe2O3 metal centre.

218

Figure 5.8: Cyclic voltammograms of GCE/Au@NPFe2O3NC upon successive addition of RuHex (a-0, b-25, c-50, d-100, and e-200 µM) to the 0.01M PBS (pH-7, scan rate = 50mVs-1).

Figure 5.9: (A) Amperometric responses of GCE/Au@NPFe2O3NC with the successive addition of RuHex solution (10 to 1100µM) into the 0.01M PBS (pH-7); (B) the corresponding calibration plot. Inset of Figure B: Lineweaver-Burk Model. 3+/2+ We subsequently studied the [Ru(NH3)6] sytem using NPFe2O3NC before and after deposition of AuNPs into NPFe2O3NC. We found that 2% AuNPs loaded Au@NPFe2O3NC generated approximately 2-times lower ipc in compare to that of NPFe2O3NC (20.11 vs 39.82 µAcm-2, Figure 5.10). This findings suggest that deposition of AuNPs may reduce the surface dependent mass transport as well as nanocube mediated electron transfer, and hence reduces 5, 30 the resultant ipc. As our assay design requires the gold-DNA affinity interaction for adsorbing target miRNA, we synthesised porous Fe2O3 which favours the loading of AuNPs to achieve Au@NPFe2O3NC as a proof-of-concept particle framework.

219

Figure 5.10: Cathodic peak currents obtained by bare GCE, Au@NPFe2O3NC-modified-GCE and NPFe2O3NC-modified GCE at room temperature, in presence of 50μM RuHex (0.01M PBS, pH-7, scan rate = 50mVs-1). Inset, corresponding cyclic voltammogram.

5.3.3. microRNA Detection

The electrocatalytic activity of the Au@NPFe2O3NC can be used as a signal amplifying label for the detection of miR-21, a potential biomarker for detecting cancer in patients with oesophageal squamous cell carcinoma (ESSC).31-35 miR-21 is usually upregulated in tissues, plasma or serum samples from patients with ESCC and significantly linked with the poor OS (overall survival) in patients with ESCC.36 Figure 5.11 represents the outline of our assay. We initially extracted the total RNA from the tissue samples of ESCC using a commercial extraction kit. To capture specific miR-21 RNA present in this sample, we designed a biotinylated-capture probe and incubated it with the extracted sample. The target miRNA was hybridized with the biotinylated probe. The hybridized dsRNA was then magnetically separated and purified by streptavidin-modified dynabead based protocol. The captured miRNAs were heat released, separated and purified by another magnetic separation step. The isolated and purified miRNA was directly adsorbed onto the Au@NPFe2O3NC-modified screen-printed carbon electrode (SPCE/Au@NPFe2O3NC) via RNA-gold affinity interaction, which follows conventional physisorption and chemisorption mechanism. This involves the direct interaction of nitrogen atoms of nucleobase ring’s with gold and partial contribution from

220

the exocyclic amino group and charge transfer between the aromatic ring and gold surface.37- 38 The adsorbed miR-21 were then detected by the chronocoulometric (CC) charge 3+ interrogation in the presence of [Ru(NH3)6] , which act as a signalling molecule that stoichiometrically binds to the anionic phosphate backbone of miRNA, and indicates the amount of miRNA adsorbed on the electrode surface.29

Figure 5.11: Assay principle. miRNA was extracted from target cell lines or tissue samples. After magnetic isolation and purification, the target miR-21 was adsorbed onto the

Au@NPFe2O3NC attached SPCE. An enhance electrochemical signals wer generated by the 3+/2+ CC interrogation of miRNA-bound Ru(NH3)6] complexes. Inset, typical CC signals (charge density vs. t1/2) showing the miRNA adsorbed SPCE/NC produces higher charge compare to an unmodified SPCE.

To check the assay functionality and specificity, we performed our assay with the same amount of starting synthetic miRNA (10nM) using Au@NPFe2O3NC-modified and unmodified-SPCE (Figure 5.12A). As expected, the charge density (4.5 μCcm-2) for the miR-

21-attached (without Au@NPFe2O3NC) SPCE (i.e., SPCE/miR-21) was very similar to that of the bare (without Au@NPFe2O3NC and miR-21) SPCE (i.e., SPCE/bare). A slightly higher -2 response (4.5 versus 8.5 μCcm ) was estimated for the Au@NPFe2O3NC-modified SPCE (i.e., SPCE/NC/buffer, NOT), indicating a low level of adsorption of RuHex onto the surface of the SPCE/NC electrode. To evaluate the specificity of the capture probes for isolating miR-21 targets, we have also performed our assay using a noncomplementary miR-107 (i.e., SPCE/NC/miR-107) and compared the data against target miR-21 at the same starting

221

concentration. We found that charges from experiments involving miR-107 were similar to the SPCE/NC electrode, indicating that our assay is not affected by the isolation of nonspecific RNAs present in the sample. Most importantly, the CC signal from the detection of target miR- 21 was about 4-fold higher (8.79 vs 29.51 μCcm-2) than that of the nonspecific miR-107. These experiments demonstrated the good specificity of our assay in isolating miR-21 RNA for miRNA-gold adsorption and subsequent electrocatalytic detection.

Figure 5.12: (A) Charge density for the SPCE/Bare, SPCE/miR-21, SPCE/NC/miR-21 (Qdl), SPCE/NC/buffer (NoT), SPCE/NC/miR-107 (Wrong target) and SPCE/NC/miR-21 (Q) electrodes. The concentration of miR-21 and miR-107 was 10 nM. (B) Typical CC curves for the (c-j) 100 fM-1.0 μM of synthetic miRNA. Curves a, and b are for the Qdl and NOT respectively. (C) Charge density-concentration profiles. Inset, corresponding calibration plot. (D) Charge density obtained for eight tissue samples derived from ESCC patients. The concentration of RuHex is 50 µM. Each data point represents the average of three independent trails, and error bars represent the standard deviation of measurements (%RSD = <5%, for n = 3).

222

To evaluate the assay sensitivity, We tested the designated concentration of synthetic target miR-21 ranging from 100 fM to 1.0 µM (Figure 5.12B,C). The increasing level of QRNA was noted with increasing concentration of miRNA. This is attributed to the increased amount of miR-21, which was isolated and thus adsorbed onto the SPCE/ Au@NPFe2O3NC surface (the surface density were estimated to be 8.52 × 1013 and 1.60 × 1013 molecules/cm2 for 1.0 µM and 100 fM of miR-21 respectively. An increased amount of adsorbed miR-21 bind with 3+ an increased amount of [Ru(NH3)6] and thereby generating a higher QRNA. The linear regression equation was estimated to be y (charge density, µCcm-2) = 3.375 (amount of miR- 2 21) – 2.31, with a correlation coefficient (R ) of 0.9961. The level of generated QRNA indicates that the minimum detectable miR-21 concentration is 100 fM. The high sensitivity of the assay can be related to; (i) the large exposed surface area of AuNPs within the Au@NPFe2O3NC adsorbs more miRNA through the RNA-gold affinity interaction, thus larger amount of 3+ [Ru(NH3)6] ions bind on the miRNA-confined surface and (ii) electrocatalytic signal 3+/2+ enhancement of [Ru(NH3)6] system by Au@NPFe2O3NC. The sensitivity of our method is similar or slightly better than those reported in previous methods (Table 5.2).22, 39-43 However, it is important to note that our method simplifies the assay design by avoiding the complex chemistries underlying each step of the sensor fabrication (i.e., cleaning of the electrode surface, the formation of the self-assembled monolayer, hybridization of the target with electrode-attached probe). Our assay also offers other advantages, such as (i) enhancement of the detection sensitivity through the catalytic activity of porous framework of Au@NPFe2O3NC (ii) provides a mean for efficient removal of non-specific species and improves the isolation purity and efficiency using repetitive magnetic bead-based isolation and purification steps, (iii) disposable SPCE based electrochemical detection offer the high translational potential of the assay at relatively low cost (AUD 4 per electrode).

223

Figure 5.13: CC charge generated by the extracted miR-21 from two HKESC-1 and HKESC- 4 cell lines.

After establishing the sensitivity and specificity of the assay, we have performed clinical validation of the assay by analysing miR-21 levels in patient-derived human ESCC cell lines, HKESC-1 and HKESC-4.44-45 As shown in Figure 5.13, with starting total RNA concentration of 10ng/mL, miR-21 levels were expressed at varying levels in these two cell lines. The expression of miR-21 in HKCSE-4 cells was found to be higher compared to HKCSE-1 cells (RT-qPCR validation data supported this results, see Figure 5.14). We further challenged our assay in-vivo by profiling the miR-21 expression in cancer and matched non- cancer tissue samples from patients diagnosed with ESSC. The ESSC patients (n=4) with primary tumours (denoted as T) and the adjacent non-neoplastic mucosae (denoted as N) from the same patients (matched) were used to detect miR-21 expression levels. Similar to cell lines, all paired cancer and non-cancer tissue samples from these patients showed varied levels of miR-21 expression. With the same amount of total RNA, our assay successfully detected high expression of miR-21 in all four patients with ESCC (P1(T), P2(T), P3(T) and P4)(T) compared to their non-neoplastic counterparts (P1(N), P2 (N), P3 (N) and P4(N). High expression of miR- 21 reported having oncogenic effects in ESCC cells leading to increased cell proliferation and growth properties.

224

Figure 5.14: RT-qPCR validation of miR-21 expression levels in the (A) two ESCC cell lines (B) four tumor tissue samples obtained from the patients with ESCC.

Notably, the conventional RT-qPCR study supported these data, and has noted approximately 3.0-, 1.5-, and 1.5-fold higher miR-21 expression in P1, P2 and P4 samples (Figure 5.14B). The overexpression of miR-21 noted in this study are in agreement with previous findings on miR-21 expression.31, 34-35 Moreover, %RSD for n = 3, in quantifying the level of miR-21 in these clinical samples was found to be <5%. These data clearly indicated that CC signals generated by our assay were able to quantify the different degree of miR-21 level in ESCC tissue samples. Thus the method has potential implications for tracking the growth and progression of ESCC.

225

Table 5.2: Comparative Analytical performance of electrochemical sensors for microRNA Assay Target LOD Comparison of LOD Remarks Ref (approx.)

Photo-electrochemical biosensing miR-7f 34 fM ~3 times lower (100 This method uses cumbersome ssDNA-Cds QDs labelling and 39 platform based on DNA–CdS quantum fM versus 34 fM) adopted DNase I amplifications. dots (QDs) sensitized SWCNTs

Electrical silicon nanowires (SiNW) let-7b 1fM 10 times 1ower (100 Detection is based on tedious, time consuming PNA 40 biosensor based on peptide nucleic acid and fM versus 1 fM) functionalization of SiNW followed by hybridization with target (PNA) let-7c miRNA. Moreover, the sensor is not reusable as chemical treatment and heat denaturation (∼95 ◦C) may damage SiNW properties.

Electrochemical detection based on miR- 10 fM ~3 times lower (100 This techniques utilizes E. Coli Poly(A) extension reaction, 41 Poly(A) Extensions 107 fM versus 10 fM) required extra incubation time and heating, which may cause target miRNA degradation.

Chronocoulometric detection based on miR- 100 fM Similar (100 fM This platform uses complicated RCA amplification and usually it 22 rolling circle 143 versus 100 fM) is not capable of amplifying a satisfactory length of nucleic acids. Amplification (RCA)

Differential pulse voltammetry using an miR- 2 nM ~1 x104 times higher In this methods, labelling of miRNA with Os(VI)bipy may 42 redox complex of osmium (VI) and 2,2 261 and (100 fM versus 10 increase the risk of contamination or sample degradation. ′-bipyridine miR- nM) 522 Amperometry using Pd nanoparticles as miR- 1.87 pM ~60 times higher (100 This method involves multi-steps electrode surface modifications 46 enhancer and linker 155 fM versus 1.87 pM) and based on tedious Thi as alinker.

Chronoamperometry based on magnetic miR- 7.0 pM ~20 times higher (100 It relies on enzymatic treatment, required biotinylation of target 47 bead based capture 222 fM versus 7.0 pM) RNA, and quantification of miRNA is carried out via p- aminophenol followed by enzyme kinetics (indirect way).

226

Chronocoulometric detection based on miR-21 100 fM This work Unlike to the traditional affinity-based detection, this assay This electrocatalytic nanoporous exploits magnetic nanoparticle-based intimate mixing, separation work superparamagnetic nanocubes. and purification of miRNA which reduce the matrix effects of the biological samples. It avoids conventional hybridization chemistries and need of expensive electrochemical tags or labelling or enzymatic amplifications.

227

5.4. Conclusion we have demonstrated the electrocatalytic activity of Au@NPFe2O3NC to develop an entirely new and sensitive electrochemical assay for non-enzymatic and amplification-free detection of miRNA. The applicability of our assay has been successfully tested and validated in cancer cell lines and a small cohort of tissue samples of patients with ESCC. We envisage that our method is not limited to miRNA detection, it could be further applied to detect a wide variety of RNA based biomarkers for other human cancers and chronic diseases.

5.5. References 1. Beveridge, J. S.; Stephens, J. R.; Williams, M. E., The use of magnetic nanoparticles in analytical chemistry. Annual Review of Analytical Chemistry 2011, 4, 251-273. 2. Urbanova, V.; Magro, M.; Gedanken, A.; Baratella, D.; Vianello, F.; Zboril, R., Nanocrystalline iron oxides, composites, and related materials as a platform for electrochemical, magnetic, and chemical biosensors. Chemistry of Materials 2014, 26 (23), 6653-6673. 3. Zakaria, M. B.; Hu, M.; Hayashi, N.; Tsujimoto, Y.; Ishihara, S.; Imura, M.; Suzuki, N.; Huang, Y. Y.; Sakka, Y.; Ariga, K., Thermal conversion of hollow Prussian blue nanoparticles into nanoporous iron oxides with crystallized hematite phase. European Journal of Inorganic Chemistry 2014, 2014 (7), 1137-1141. 4. Zakaria, M. B.; Belik, A. A.; Liu, C. H.; Hsieh, H. Y.; Liao, Y. T.; Malgras, V.; Yamauchi, Y.; Wu, K. C. W., Prussian Blue Derived Nanoporous Iron Oxides as Anticancer Drug Carriers for Magnetic‐Guided Chemotherapy. Chemistry–An Asian Journal 2015, 10 (7), 1457-1462. 5. Leung, K. C.-F.; Xuan, S.; Zhu, X.; Wang, D.; Chak, C.-P.; Lee, S.-F.; Ho, W. K.- W.; Chung, B. C.-T., Gold and iron oxide hybrid nanocomposite materials. Chemical Society Reviews 2012, 41 (5), 1911-1928. 6. Zhuo, Y.; Yuan, P.-X.; Yuan, R.; Chai, Y.-Q.; Hong, C.-L., Bienzyme functionalized three-layer composite magnetic nanoparticles for electrochemical immunosensors. Biomaterials 2009, 30 (12), 2284-2290. 7. Zhao, J.; Zhang, Y.; Li, H.; Wen, Y.; Fan, X.; Lin, F.; Tan, L.; Yao, S., Ultrasensitive electrochemical aptasensor for thrombin based on the amplification of aptamer–AuNPs–HRP conjugates. Biosensors and Bioelectronics 2011, 26 (5), 2297-2303. 228

8. Liu, B.; Cui, Y.; Tang, D.; Yang, H.; Chen, G., Au (III)-assisted core–shell iron oxide@ poly (o-phenylenediamine) nanostructures for ultrasensitive electrochemical aptasensors based on DNase I-catalyzed target recycling. Chemical Communications 2012, 48 (20), 2624-2626. 9. Dong, H.; Lei, J.; Ding, L.; Wen, Y.; Ju, H.; Zhang, X., MicroRNA: function, detection, and bioanalysis. Chemical Reviews 2013, 113 (8), 6207-6233. 10. Lin, S.; Gregory, R. I., MicroRNA biogenesis pathways in cancer. Nature Reviews Cancer 2015, 15 (6), 321-333. 11. Islam, M. N.; Masud, M. K.; Haque, M. H.; Hossain, M. S. A.; Yamauchi, Y.; Nguyen, N. T.; Shiddiky, M. J., RNA biomarkers: diagnostic and prognostic potentials and recent developments of electrochemical biosensors. Small Methods 2017, 1 (7), 1700131. 12. Lusi, E.; Passamano, M.; Guarascio, P.; Scarpa, A.; Schiavo, L., Innovative electrochemical approach for an early detection of microRNAs. Analytical Chemistry 2009, 81 (7), 2819-2822. 13. Xia, N.; Wang, X.; Deng, D.; Wang, G.; Zhai, H.; Li, S.-J., Label-free electrochemical sensor for MicroRNAs detection with ferroceneboronic acids as redox probes. International Journal of Electrochemical Science 2013, 8 (7), 9714- 9722. 14. Wu, S.; Chen, H.; Zuo, Z.; Wang, M.; Luo, R.; Xu, H., A simple electrochemical biosensor for rapid detection of microRNA based on base stacking technology and enzyme amplification. Int. J. Electrochem. Sci 2015, 10, 3848-3858. 15. Hamidi-Asl, E.; Palchetti, I.; Hasheminejad, E.; Mascini, M., A review on the electrochemical biosensors for determination of microRNAs. Talanta 2013, 115, 74- 83. 16. Lehninger, A. L.; Nelson, D. L.; Cox, M. M.; Cox, M. M., Lehninger principles of biochemistry. Macmillan: 2005. 17. Lineweaver, H.; Burk, D., The determination of enzyme dissociation constants. Journal of the American chemical society 1934, 56 (3), 658-666. 18. Bard, A. J., LR Faulkner Electrochemical Methods. Fundamentais and Applications John Wiley and Sons 1980. 19. Shiddiky, M. J.; Torriero, A. A.; Zhao, C.; Burgar, I.; Kennedy, G.; Bond, A. M., Nonadditivity of faradaic currents and modification of capacitance currents in the voltammetry of mixtures of ferrocene and the cobaltocenium cation in protic and 229

aprotic ionic liquids. Journal of the American Chemical Society 2009, 131 (23), 7976-7989. 20. Koo, K. M.; Carrascosa, L. G.; Shiddiky, M. J.; Trau, M., Amplification-free detection of gene fusions in prostate cancer urinary samples using mrna–gold affinity interactions. Analytical Chemistry 2016, 88 (13), 6781-6788. 21. Steel, A. B.; Herne, T. M.; Tarlov, M. J., Electrochemical quantitation of DNA immobilized on gold. Analytical Chemistry 1998, 70 (22), 4670-4677. 22. Yao, B.; Liu, Y.; Tabata, M.; Zhu, H.; Miyahara, Y., Sensitive detection of microRNA by chronocoulometry and rolling circle amplification on a gold electrode. Chemical Communications 2014, 50 (68), 9704-9706. 23. Haque, M. H.; Gopalan, V.; Yadav, S.; Islam, M. N.; Eftekhari, E.; Li, Q.; Carrascosa, L. G.; Nguyen, N.-T.; Lam, A. K.; Shiddiky, M. J., Detection of regional DNA methylation using DNA-graphene affinity interactions. Biosensors and Bioelectronics 2017, 87, 615-621. 24. Gopalan, V.; Pillai, S.; Ebrahimi, F.; Salajegheh, A.; Lam, T. C.; Le, T. K.; Langsford, N.; Ho, Y. H.; Smith, R. A.; Lam, A. K. Y., Regulation of microRNA‐ 1288 in colorectal cancer: Altered expression and its clinicopathological significance. Molecular Carcinogenesis 2014, 53 (S1), E36-E44. 25. Yadav, S.; Masud, M. K.; Islam, M. N.; Gopalan, V.; Lam, A. K.-y.; Tanaka, S.; Nguyen, N.-T.; Al Hossain, M. S.; Li, C.; Yamauchi, M. Y., Gold-loaded nanoporous iron oxide nanocubes: a novel dispersible capture agent for tumor- associated autoantibody analysis in serum. Nanoscale 2017, 9 (25), 8805-8814. 26. Mustapić, M.; Horvat, J.; Hossain, M. S.; Sun, Z.; Skoko, Ž.; Mitchell, D. R.; Dou, S. X., Novel synthesis of superparamagnetic Ni–Co–B nanoparticles and their effect on superconductor properties of MgB2. Acta Materialia 2014, 70, 298-306. 27. Zysler, R.; Fiorani, D.; Testa, A.; Suber, L.; Agostinelli, E.; Godinho, M., Size dependence of the spin-flop transition in hematite nanoparticles. Physical Review B 2003, 68 (21), 212408. 28. Cardillo, D.; Tehei, M.; Hossain, M. S.; Islam, M. M.; Bogusz, K.; Shi, D.; Mitchell, D.; Lerch, M.; Rosenfeld, A.; Corde, S. p., Synthesis-dependent surface defects and morphology of hematite nanoparticles and their effect on cytotoxicity in vitro. ACS Applied Materials & Interfaces 2016, 8 (9), 5867-5876. 29. Zhang, J.; Song, S.; Zhang, L.; Wang, L.; Wu, H.; Pan, D.; Fan, C., Sequence- specific detection of femtomolar DNA via a chronocoulometric DNA sensor (CDS): 230

Effects of nanoparticle-mediated amplification and nanoscale control of DNA assembly at electrodes. Journal of the American Chemical Society 2006, 128 (26), 8575-8580. 30. Cui, H.-Z.; Guo, Y.; Wang, X.; Jia, C.-J.; Si, R., Gold-iron oxide catalyst for CO oxidation: effect of support structure. Catalysts 2016, 6 (3), 37. 31. Winther, M.; Alsner, J.; Tramm, T.; Baeksgaard, L.; Holtved, E.; Nordsmark, M., Evaluation of miR-21 and miR-375 as prognostic biomarkers in esophageal cancer. Acta Oncologica 2015, 54 (9), 1582-1591. 32. Mori, Y.; Ishiguro, H.; Kuwabara, Y.; Kimura, M.; Mitsui, A.; Ogawa, R.; Katada, T.; Harata, K.; Tanaka, T.; Shiozaki, M., MicroRNA-21 induces cell proliferation and invasion in esophageal squamous cell carcinoma. Molecular Medicine Reports 2009, 2 (2), 235-239. 33. Tanaka, Y.; Kamohara, H.; Kinoshita, K.; Kurashige, J.; Ishimoto, T.; Iwatsuki, M.; Watanabe, M.; Baba, H., Clinical impact of serum exosomal microRNA‐21 as a clinical biomarker in human esophageal squamous cell carcinoma. Cancer 2013, 119 (6), 1159-1167. 34. Nouraee, N.; Van Roosbroeck, K.; Vasei, M.; Semnani, S.; Samaei, N. M.; Naghshvar, F.; Omidi, A. A.; Calin, G. A.; Mowla, S. J., Expression, tissue distribution and function of miR-21 in esophageal squamous cell carcinoma. PloS one 2013, 8 (9), e73009. 35. Komatsu, S.; Ichikawa, D.; Kawaguchi, T.; Miyamae, M.; Okajima, W.; Ohashi, T.; Imamura, T.; Kiuchi, J.; Konishi, H.; Shiozaki, A., Circulating miR-21 as an independent predictive biomarker for chemoresistance in esophageal squamous cell carcinoma. American Journal of Cancer Research 2016, 6 (7), 1511. 36. Kumarswamy, R.; Volkmann, I.; Thum, T., Regulation and function of miRNA-21 in health and disease. RNA Biology 2011, 8 (5), 706-713. 37. Sina, A. A. I.; Howell, S.; Carrascosa, L. G.; Rauf, S.; Shiddiky, M. J.; Trau, M., eMethylsorb: electrochemical quantification of DNA methylation at CpG resolution using DNA–gold affinity interactions. Chemical Communications 2014, 50 (86), 13153-13156. 38. Koo, K. M.; Sina, A. A.; Carrascosa, L. G.; Shiddiky, M. J.; Trau, M., DNA–bare gold affinity interactions: mechanism and applications in biosensing. Analytical Methods 2015, 7 (17), 7042-7054.

231

39. Cao, H.; Liu, S.; Tu, W.; Bao, J.; Dai, Z., A carbon nanotube/quantum dot based photoelectrochemical biosensing platform for the direct detection of microRNAs. Chemical Communications 2014, 50 (87), 13315-13318. 40. Zhang, G.-J.; Chua, J. H.; Chee, R.-E.; Agarwal, A.; Wong, S. M., Label-free direct detection of MiRNAs with silicon nanowire biosensors. Biosensors and Bioelectronics 2009, 24 (8), 2504-2508. 41. Koo, K. M.; Carrascosa, L. G.; Shiddiky, M. J.; Trau, M., Poly (A) extensions of miRNAs for amplification-free electrochemical detection on screen-printed gold electrodes. Analytical Chemistry 2016, 88 (4), 2000-2005. 42. Bartosik, M.; Trefulka, M.; Hrstka, R.; Vojtesek, B.; Palecek, E., Os (VI) bipy-based electrochemical assay for detection of specific microRNAs as potential cancer biomarkers. Electrochemistry Communications 2013, 33, 55-58. 43. Wen, Y.; Liu, G.; Pei, H.; Li, L.; Xu, Q.; Liang, W.; Li, Y.; Xu, L.; Ren, S.; Fan, C., DNA nanostructure-based ultrasensitive electrochemical microRNA biosensor. Methods 2013, 64 (3), 276-282. 44. Hu, Y.-C.; Lam, K. Y.; Wan, T. S.; Fang, W.-G.; Ma, E. S.; Chan, L. C.; Srivastava, G., Establishment and characterization of HKESC-1, a new cancer cell line from human esophageal squamous cell carcinoma. Cancer Genetics and Cytogenetics 2000, 118 (2), 112-120. 45. Cheung, L. C.; Tang, J. C.; Lee, P.; Hu, L.; Guan, X.-Y.; Tang, W.; Srivastava, G.; Wong, J.; Luk, J. M.; Law, S., Establishment and characterization of a new xenograft-derived human esophageal squamous cell carcinoma cell line HKESC-4 of Chinese origin. Cancer Genetics and Cytogenetics 2007, 178 (1), 17-25. 46. Chen, A.; Ma, S.; Zhuo, Y.; Chai, Y.; Yuan, R., In situ electrochemical generation of electrochemiluminescent silver naonoclusters on target-cycling synchronized rolling circle amplification platform for microRNA detection. Analytical Chemistry 2016, 88 (6), 3203-3210. 47. Bettazzi, F.; Hamid-Asl, E.; Esposito, C. L.; Quintavalle, C.; Formisano, N.; Laschi, S.; Catuogno, S.; Iaboni, M.; Marrazza, G.; Mascini, M.; Cerchia, L.; De Franciscis, V.; Condorelli, G.; Palchetti, I., Electrochemical detection of miRNA-222 by use of a magnetic bead-based bioassay. Analytical and Bioanalytical Chemistry 2013, 405 (2), 1025-1034.

232

Chapter 6

Electrochemical Detection of Motor Neuron Disease (MND) Derived Exosomal miRNA at Attomolar Sensitivity*

*Sections of this chapter are based on Masud, M.K., et al., Journal of Material Chemistry B, 2020 (submitted).

233

6. Electrochemical Detection of Motor Neuron Disease (MND) Derived Exosomal miRNA at Attomolar Sensitivity

6.1. Introduction Motor neuron disease (MND) is a progressive and fatal neurodegenerative disorder, leading to the loss of upper and lower motor neurons and eventually death within 2-3 years of illness onset.1-2 MND results from the relapse of lower motor neurons in the spinal cord and/or the brainstem and upper motor neurones in the motor cortex, or both.3-4 It ultimately leads to progressive loss of the use of their limbs or causes paralysis, i.e. it affects the ability to speak, move, breathe and swallow. Among the four clinical phenotypes of MND (amyotrophic lateral sclerosis, isolated bulbar palsy, progressive muscular atrophy and primary lateral sclerosis), amyotrophic lateral sclerosis (ALS) is the most common type representing 70% of the cases occurring 2-3 per 100,000 people worldwide in a year.5 Recent studies have shown that mutations in the genes encoding the RNA-binding proteins FUS/TLS [ALS6 locus]6 and TARDBP/TDP43 [ALS10 locus]7 recommend the critical roles of regulatory RNA in the pathogenesis of ALS.8 Interestingly, these disease-associated RNA-binding proteins were recognized in neuronal RNA granules and with microRNA (miRNA)-associated complexes.9 To date, several studies have shown that miRNAs are differentially expressed in ALS patients when compared to controls in a variety of biofluids, including CSF, and in the blood-derived components plasma and serum and therefore considered as potential biomarkers for ALS.

MicroRNAs (miRNAs) are a large group of small (~ 22 nucleotides) non-coding single- strand RNAs that play essential roles in gene expression via post-transcriptional regulation. They generally bind the 3´-untranslated region of mRNAs, leading to the gene silencing through mRNA cleavage, translational repression and adenylation.10-11 Dysregulation of these highly conserved regulatory RNAs can potentially impact on the progression and prognosis of the disease and have gained immense interest in recent time as diagnostic biomarkers, especially in cancer.8 Non-coding RNAs are also reported as profusely expressed in the central nervous system (CNS), and aberrancies in miRNA expression patterns have been described in several neurodegenerative diseases.12 However, the roles of individual miRNAs are not fully understood in neurodegenerative disease (probably due to the nervous system complexity and technical difficulties), but several reports have provided evidence that miRNAs play significant roles in neurodegenerative disorders, including miR338-3p/miR-106/miR-451 in ALS.13-14 For instance, miR-338-3p is stated to control several molecular pathways and contribute to ALS

234

pathological processes through apoptosis, neurodegeneration, and/or glutamate clearance.15 Furthermore, miR-338-3p expression was noticeably upregulated in ALS patient leukocytes, serum, CSF and spinal cord in comparison to that of healthy controls, identifying it as an attractive prognostic circulating biomarker of ALS disease onset.16 Nevertheless, circulating miRNA suffers from digestion by RNAase and other environmental damage, therefore encapsulated miRNA such as exosomal (the lipid-bilayer protects them from RNase degradation, providing a stable source of miRNAs) or apoptotic miRNAs could be highly informative for early diagnosis.10, 17 Currently, miRNA detection mostly relies on conventional nucleic acid detection assays such as quantitative reverse transcription PCR (RT-qPCR), microarrays, Northern blot, and RNA-sequencing. Despite being reliable in laboratory settings, these conventional techniques are expensive and not suitable for the resource-poor and decentralized settings.10-11 Electrochemical assays, on the contrary, have shown more potential for clinical application due to their inherent advantages of being inexpensive, simple, rapid, and miniaturized. Most of the electrochemical sensors for miRNA however still rely on multiple sensor fabrication steps, some sorts of enzymatic amplification ( e.g. isothermal transcription mediation amplification (TMA),18 or reverse transcription-PCR (RT-PCR)19 and target RNA modification (e. g., polyadenylation, labelling) which could destabilize RNA and complicate the assay protocol.10 Furthermore, having rigorous target selectivity and faster analysis time, many of these sensors lack additional signal enhancement steps, thereby failing to achieve the sensitivity levels required for the analysis of miRNA in clinical samples.20

With the latest advancement in nanotechnology, nanostructured materials with superparamagnetic behaviour, and biocompatibility are currently transitioning to a new paradigm of applications in the fields of biosensing, where they are used in developing novel methods and devices for diagnosis and monitoring of specific diseases via detecting levels of disease-specific biomolecules.11, 21 They exhibit advantages in molecular diagnostics, particularly in disease diagnosis applications by breaking down the barrier for structural miniaturization of diagnostic platforms, accelerating the signal transduction and hence boosting the sensitivity of the analysis. Their electrocatalytic properties can be exploited to espouse many novel transduction schemes.21-23 Recently, we have reported a new class of gold- loaded superparamagnetic ferric oxide nanocubes (AuNP-Fe2O3NC), which exhibits multiple functionalities, e.g. enhanced catalytic activity toward the common electroactive molecules,21 direct adsorption aptitude for a large number of nucleic acids (DNA, RNA) through gold−nucleic acid affinity interactions,23 magnetic dispersible capture vehicles,24 and

235

peroxidase mimetic activity (as nanozymes).25-26 However, the catalytic activity of the nanocubes towards the methylene blue (a readily engaged nucleic acid redox marker) redox reaction and the synergistic effect on cascade electrocatalysis of nanocube surface-bound miRNA interrogation with methylene blue in the presence and absence of ferricyanide is not yet reported. The multi coupling (three-mode chemical amplification) system comprising electrocatalytic nanostructures, methylene blue and ferricyanide redox couples, have the potential for detecting ultra-low amounts of miRNA present in neurodegenerative disease (e.g. ALS) samples.

Herein, we report the electrocatalytic properties of the AuNP-Fe2O3NC nanocube towards the reduction of methylene blue for the development of ultrasensitive exosomal miRNA detection platform obtained from exosome extracted from preconditioned media of motor neurons (ALS). The exosome extracted and magnetically purified miRNAs were directly adsorbed onto the AuNP-Fe2O3NC modified (magnetically bound onto a screen-printed carbon electrode-SPCE) through gold-RNA affinity interaction. The level of miRNAs was quantified by chronocoulometric (CC) charge interrogation in the presence of methylene blue (MB), electrostatically attached with the guanine bases of surface-bound target miRNA. The signal was further enhanced with the coupling of a higher amount of the ferri/ferrocyanide 3-/4- 3-/4- ([Fe(CN)6] ) system (i. e., MB/[Fe(CN)6] ). We considered miR-338-3p as a model target to test the applicability of our assay both in synthetic and in ALS motor neuron -derived exosomal miRNA samples, which was reported to have a strong correlation with the progression of MND. The multi-enhancement steps facilitate our assay to accomplish a detection limit of 100 aM with good reproducibility (% RSD = <5%, for n = 3).

236

6.2 Experimental

6.2.1. Materials and Instrumentation Reagent grade methylene blue (MB), phosphate buffered saline (PBS) tablet (0.01 M diphosphate buffer, 0.0027 M potassium chloride and 0.137 M sodium chloride, pH 7.4 at 25 oC) and cyclic AMP were purchased from Sigma Life Science (Australia). Analytical grade hydrochloric acid (HCl) was purchased from Chem-supply (Australia). N-2, B27, GlutaMAX, MEM non-essential amino acids supplements, accutase, collagen, laminin, fibronectin and neurobasal media were purchased from Life technologies, (Australia). Human FGF-2, glial cell line-derived neurotrophic factor and purmorphamine were obtained from miltenyi biotec (Australia). Tris was obtained from VWR Life science (Australia), glassy carbon electrode (GCE) was purchased from CH instrument (USA). Screen-printed carbon electrode (SPCE) with a three-electrode system printed on a ceramic substrate (length 34 × width 10 × height 5 mm) (DRP-150) from Dropsens (Spain). In the three-electrode system, working (4 mm diameter), counter and reference electrodes were carbon, platinum, and silver-modified. All chemical and reagent were used as received without additional purification. Oligonucleotides were acquired from Integrated Technologies, USA and sequences are shown in Table 6.11.

Table 6.1: Oligonucleotide sequences (miR-338-3p) Oligos 5 ́-Sequences-3 ́ Biotinylated miR-338-3p capture CAA CAA AAT CAC TGA TGC TGG A/3Biotin/ probe Synthetic miR-338-3p sequence UCC AGC AUC AGU GAU UUU GUU G

Electrochemical measurements have carried out using a CHI650 electrochemical workstation. (CH instrument, USA). Cyclic voltammetry (CV) and chronocoulometry (CC) performed in a single-compartment cell with a 3-mL volume and using in SPCE. A conventional three-electrode system, comprising a bare or modified GCE, a platinum auxiliary electrode, and an Ag/AgCl3 1.0 M NaCl reference electrode (CH instrument, Inc. USA), was used for the measurement of electrocatalytic activity. The CV was measured between -200 and 600 mV unless stated otherwise. CC was carried out between 0 and -500mV, 25 ms pulse width, and 2 ms sample interval. A temperature and time control ultrasonic water bath

(Soniclean, Australia) was applied for the dispersion of AuNP-Fe2O3NC nano-hybrid before applying to the electrode surface.

237

6.2.2. Electrocatalytic activity A GCE (3 mm diameter) was polished using 0.3 and 0.05 mm alumina slurry (CH Instrument, Inc. USA) followed by rinsing with an adequate amount of water. After successive sonication with ethanol and water, the electrode was again rinsed thoroughly using DI water, allowed it to dry at room temperature and gave a mirror surface. To assess the electrocatalytic activity of

AuNP-Fe2O3NC, 7 μg (1 μg/μL) of a colloidal suspension of AuNP-Fe2O3NC were drop-dried onto the surface of the clean GCE electrode and allowed to dry. The cyclic voltammetry was then performed at room temperature with the conventional three-electrode system using AuNP-

Fe2O3NC-modified GCE as the working electrode.

6.2.3. Cell culture and cell culture media collection All experimental protocols were approved by the University of Wollongong Human Research Ethics Committees. The methods were carried out by the guidelines as set out in the National Statement on Ethical Conduct in Research Involving Humans, and informed consent was obtained from all donors. Skin samples were collected from two healthy individuals and three ALS patients, and human feeder-free iPSCs were generated as previously described.27-28 The iPSCs were confirmed pluripotent by Pluritest and were karyotyped to verify the lack of chromosomal changes during reprogramming. The iPSCs were cultured on Matrigel (Corning) coated tissue culture plates in TeSR-E8 (Stem Cell Technologies) at 37 °C, 5% CO2 in a humidified incubator.

Differentiation of iPSCs to motor neurons was performed as previously described.29-30 Using cell scrapers, iPSC colonies were detached from the plate and cultured in a non-tissue culture plate for 4 days in neural induction media (DMEM/F12 media supplemented with 1% N-2, 0.4% B27, 1% GlutaMAX, 1% MEM non-essential amino acids) to form embryoid bodies. Embryoid bodies were then collected and plated onto tissue culture plates coated with 20 ug/mL of laminin in neural induction media. Half media changes were performed every other day, and 2.5 ng/mL FGF-2 was added to the media once radially organised structures (neural rosettes) were observed. After approximately 1 week, neural rosettes were isolated and dissociated to single cells using 1x accutase, plated onto tissue culture plates coated with 0.1 mg/mL collagen I (Life Technologies), 20 µg/mL laminin and 20 µg/mL fibronectin at a density of approximately 4000 cells/cm2. Half media changes were performed every other day with neuronal precursor cell media (DMEM/F12 supplemented with 1% N-2, 0.5% B27, 1%

238

Glutamax, 1% MEM non-essential amino acids and 10 ng/mL FGF-2) for 1 week. Cells were then passaged and plated onto tissue culture plates coated with 20 μg/mL laminin and 10 μg/mL fibronectin. Half media changes were performed every other day for 4 days in neuronal precursor cell media. The remaining cells were treated with an increasing concentration of retinoic acid (Sigma Aldrich) over 3 days (0.1 µM on day 1, 0.2 µM on day 2, 0.3 µM on day 3) and on the third day, 2 µM purmorphamine was also included in the media. Cells were then grown in motor neuron precursor cell media (neurobasal media supplemented with 0.5% N2, 1% Glutamax, 1% MEM non-essential amino acids, 5 ng/mL FGF-2, 0.1 µm retinoic acid (Sigma Aldrich) and 2 µM puramorphamine for 1 week with media changes performed every other day. Finally, motor neurons were matured over a period of 4 weeks in motor neuron media (neurobasal media supplemented with 0.5% N-2, 0.2% B27, 1% Glutamax, 1% MEM non-essential amino acids, 10 ng/mL glial cell line-derived neurotrophic factor, 10 ng/mL brain-derived neurotrophic factor, 10 ng/mL insulin-like growth factor 1 (STEM CELL Technologies) and 10 ng/mL cyclic AMP. Half media changes were performed every other day. At 2 and 4 weeks of motor neuron differentiation, the conditioned media was collected and stored at 4 °C until exosome isolation for subsequent miRNA analysis.

6.2.4. Exosome isolation Total exosome obtained from motor neuron preconditioned media using total exosome isolation reagent (Invitrogen) following manufacturers guidelines. In brief, 2mL of cell culture media was centrifuged at 2000×g for 30 minutes to remove cell or debris. The supernatant was then transferred to a new tube followed by the addition of 1mL of total exosome isolation reagent (Invitrogen). The mixture was incubated at 4°C for overnight and finally centrifuged at 10,000×g for 1 hour at 4°C. After that, the supernatant was discarded, and the pellet was resuspended in 50µL of 1×PBS buffer. The resuspended exosome was then stored at -20°C for the isolation of small RNA present in exosome.

6.2.5. Isolation of exosomal (total) small RNA The RNA was extracted from exosomes using the total exosome RNA & protein isolation kit (Life Technologies, Australia) as per the manufacturer’s instructions. Briefly, the exosome was diluted with 10 mM of 1×PBS to had a total volume of 200 µL followed by the addition of the same amount of 2×denaturing solution. To lyse the exosomes, acid-phenol: chloroform was added and vortexed vigorously for a minute. They were then centrifuged at 12,000 g for 5 minutes at the room-temperature to split up the mixture into aqueous (upper) and organic

239

(lower) phases. The upper phase was then subsequently treated with ethanol to dissolve and remove larger RNA and DNA present in the sample. The loop of small miRNA was eluted from filter cartridge and stored -20˚Ϲ for further analysis.

6.2.6. Probe hybridization and magnetic isolation of target miRNA (miR-338-3p) Target miRNA was captured by hybridizing with a capture probe followed by biotinylated magnetic beads functionalization and magnetic isolation and heat release of miRNA by our previous method.21-22, 31-32 Briefly, for probe hybridization 10µL of exosomal small RNA, were mixed with 10 μL of 10 μM biotinylated capture probes. The mixture was heated at 65°C for two minutes and placed on a thermo-mixture under stirring for 1 hour at room temperature. For magnetic isolation, 10µL of commercial streptavidin-labelled magnetic beads (Dynabeads® MyOne™ Streptavidin C1, Invitrogen, Australia) was washed with 2×B&W solution and resuspended in 10 μL of 2× B&W buffer. The resuspended dynabeads were then added to the prepared capture probe-exosomal small RNA hybrid, mixed thoroughly, and incubated on a thermos-mixer for 30 min at room temperature to allow dynabeads labelling of capture probes via biotin-streptavidin interactions. After the incubation and washing with a B&W solution, the miRNA attached beads were isolated with an external magnet and resuspended in 9 µL of RNase free water. The resuspended miRNA-hybrid mixture was heated for 2 min at 95oC, and the bead was immediately attached with a magnet and supernatant containing the desired miRNA was collected. Before applying miRNA on to AuNP-Fe2O3NC-modified electrode, the miRNA was diluted two times with 5×SSC buffer.

6.2.7. Electrochemical detection of adsorbed microRNA CC readouts were obtained in 40 mM Tris-HCl buffer (pH 7.4) in absence and presence of 2 µM MB with a potential step of 5 mV and pulse width of 250 ms, and a sample interval of 2 minutes. For detecting miRNA, 5 µL of AuNP-Fe2O3NC (1 mg/mL) was added on the SPCE. The electrode was then positioned on a permanent magnet. 5 μL of target miRNA sample were then incubated onto the magnetically attached AuNP-Fe2O3NC/SPCE surface for 30 minutes followed by PBS washing. The electrode was then incubated with 7 μL of 2 μM MB so that positively charged MB+ can bound with the negatively charged phosphate backbone of adsorbed miRNA. The electrode was then washed with PBS before performing CC measurement. CC measurement was then carried out in 40 mM Tris-HCl buffer (pH 7.4) and 3- 4 mM [Fe(CN)6] in 40 mM Tris-HCl buffer (pH 7.4). Using CC, the amount of miRNA adsorbed onto the AuNP-Fe2O3NC/SPCE surface was then calculated from the number of

240

cationic redox molecules (MB) electrostatically associated with the surface-attached anionic phosphate backbone of miRNA. The total charge Q at a time t can be expressed by the integrated Cottrell equation,33 2nFA1/ 2C* Q  0 0 t1/ 2  Q  nFA … … … … … (2)  1/ 2 dl 0 where n is the number of electrons involved in electrode reaction, F is Faraday constant 2 2 (C/equivalent), A is the electrode area (cm ), Do is the diffusion coefficient (cm /s), Co* is the

2 bulk concentration (mol/cm ), 0 is representing the amount of MB-confined near electrode surface and nFA 0 (known as surface excess) is the charge obtained by adsorbed miRNA. CC curves were constructed by plotting the charge flowing through the RNA-attached electrode 1/2 1/2 versus square-root of time (t /s ) in the presence and absence MB. Q and Qdl were estimated from the intercept of these two curves at t = 0. Therefore, Q represents the total charge comprising both Faradic and non-Faradic (capacitive) charges. Hence, the corresponding charge of MB (electrostatically bound to surface-confined RNA) can be calculated as,

QRNA = Q - Qdl … … … … … (3) And the saturated surface density of MB could be used to calculate surface attached miRNA using the following equation;

RNA  (QRNA NA/nFA)(z/m) … … … … … (4) where n is the number of electrons involved in the reaction (n = 1), A is the working electrode area, NA is the Avogadro’s number, m is the number of nucleotides in the RNA, and z is the charge of redox molecules (for MB, z = 1).

6.2.8. Statistical analysis Two sample comparisons were tested for statistical significance using a two-way ANOVA. P- values < 0.05 were considered statistically significant.

6.2.9. qRT-PCR validation The extracted small RNA was converted to cDNA using the MiScript reverse transcription kit (Qiagen, Hilden, NRW, Germany) following a published protocol.21 The cDNA samples were aliquoted at 30 ng µL−1 and stored at -20°C until further use for RT-qPCR analysis. The expression level of miRNA-338-3p was quantified using RT-qPCR. The reaction mixture was in a total volume of 10 µL comprising 5.0 µL of a 2× SensiMix SYBR No-ROX master mix

241

(Bioline, London, UK), 1.0 µL each of 1.0 µmol µL−1 primer, 1.0 µL of cDNA (30 ng µL−1 ) and 2.0 µL of nuclease-free water. Thermal cycling was initiated with a first denaturation step at 95°C for 10 min followed by 40 cycles of 95°C for 15 s (denaturation), 57°C for 30 s (annealing), and 72°C for 30 s (extension). Assays were accomplished in triplicate to verify the results and a non-template control was included in all the experiments.

6.3. Result and discussion

6.3.1. Electrocatalytic activity of nanocubes towards Methylene Blue (MB) To achieve ultrasensitive detection, nanostructured materials should have high catalytic activity, conductivity, biocompatibility and a synergetic effect among them to accelerate the signal transduction towards the electrocatalytic reduction of target-bound or intercalated redox 11 molecules. To assess the electro-catalytic activity of AuNP-Fe2O3NC, AuNP-Fe2O3NC modified GCE was directly used as a working electrode for electrochemical detection of electroactive redox marker MB in 0.01M PBS (pH 7.0). It has been shown from Figure 6.1a,

AuNPs-Fe2O3NC modified-GCE (GCE/AuNP-Fe2O3NC) displayed a pair of well-defined redox peaks in 50 μM MB solution at -241 mV and -272 mV (vs. Ag/AgCl), indicating a two- electron redox process of MB (ΔE = 31 mV, Figure 6.1a. The GCE/AuNP-Fe2O3NC) offered significantly enhanced cathodic (ipc), and anodic (ipa) peak currents in comparison with the _2 GCE/bare electrode. Notably, ipc increased approximately 2.8-times (7.50 vs. 18.5 µA cm ) _2 with Epc shifted by -19 mV, whereas ipa increased approximately 2.8-times (7.5 vs. 21 µA cm ) with Epa moved by around -27 mV. The very similar current response was also obtained from differential pulse voltammetry (DPV) measurements (23.2 vs. 71.5 µA cm_2; Figure 6.1b).

These data indicate that AuNP-Fe2O3NC catalysed both the oxidation and reduction process of MB.

242

Figure 6.1: Electrocatalytic activity of Au-Fe2O3NC in MB. Comparison of the (a) CVs and

(b) DPV obtained at an unmodified GCE and AuNP-Fe2O3NC-modified GCE in 50 µM MB -1 (scan rate, 50 mVs ). (c) CV obtained at GCE/AuNP-Fe2O3NC-electrodes at different scan rate (50μM MB, 0.01M PBS, pH 7.0); (d) corresponding curves for ipc and ipa (current density) as a function of ν1/2.

Regarding the electrochemical mechanism of nanocube catalysis (diffusion or adsorption controlled) that take place on the GCE electrode surface, we performed CV measurements of both bare and GCE/AuNP-Fe2O3NC modified GCE as a function of different -1 scan rate (ν). Both ipc and ipa are proportional at the scan rates values (10-1500 mVs ) and 1/2 showed a linear relationship with ν for both the bare and AuNP-Fe2O3NC modified GCE, which indicates the electrode process is diffusion-controlled (Figure. 6.1c and d)33 and the 1/2 -1 2 equations can be expressed as, ipc (µA) = 0.9404 ν (mVs ) ̶ 2.7240, r = 0.97 for bare GCE 1/2 -1 2 (Figure 6.2b and c) and ipa (µA) = -0.6443 ν (mVs ) + 3.5644, r = 0.9714 for AuNP- 1/2 Fe2O3NC modified GCE (Figure 6.2e and f). Moreover, the plot of ipa and ipc versus ν for

AuNP-Fe2O3NC modified GCE (Figure 6.2e and f) showed a steeper slope than that of the bare

GCE (Figure 6.2b and b), verifying the catalytic performance of AuNP-Fe2O3NC towards the 1/2 redox reaction of MB. However, the slope for ipc versus ν was exhibited steeper than that of

243

1/2, ipa versus ν suggesting the AuNP-Fe2O3NC has higher catalytic activity towards reduction of MB than that of oxidation.

Figure 6.2: (a) Represents the modification of a GCE by AuNPs-Fe2O3NC for interrogating catalytic activity of NCs towards the redox process of MB; (b) and (c) depicts the calibration curve of square root of potential (scan rate) vs. current density obtained from scan rate study of bare GCE towards reduction and oxidation process of MB respectively. The CV curve obtained from the interrogation of different scan rate ranging from 10 to 1500 mVs-1 using a bare GCE is figured in d. Figure e and f represents the calibration curve of the square root of potential (scan rate) vs. current density obtained from scan rate study of AuNPs-Fe2O3NC- modified GCE towards reduction and oxidation process of MB respectively.

To further examine the performance of and AuNP-Fe2O3NC, we conducted the CV measurement at different concentration of MB solution (0 to 200 µM) of the MB (Figure 6.3). With increasing strength of the MB, the observed currents increase gradually, reaching saturation at 150 µM level of MB, revealing that the porous structure provides enormous surface to accelerate the redox reactions (Figure 6.3).

244

Figure 6.3: (a) Amperometric responses of AuNP-Fe2O3NC-modified GCE with the successive addition of MB solution (0 to 200 µM) into 0.01 M PBS (pH-7); (b) the corresponding calibration plot.

6.3.2. Electrocatalytic detection of miRNA

Our novel assay for miRNA biosensor based on electrocatalytic AuNP-Fe2O3NC and redox cycling schematically depicted in Figure 6.4. To demonstrate the working principle of the developed sensor, we initially extracted the total RNA from the exosomes obtained from preconditioned media of motor neurons (healthy controls and ALS patients). To capture specific miRNA, we designed and applied a biotinylated capture probe and had adopted magnetic bead-based isolation. Target miRNA samples were screened out through magnetic isolation and extracted by heating release from Magneti bead-DNA hybrid (through the breakdown of dsDNA to magnetic bead attached capture probe and target miRNA. The miRNA was directly adsorbed on to the AuNP-Fe2O3NC attached SPCE. The adsorbed miRNA were then detected by CC interrogation in the presence of an electroactive redox marker MB. It has been shown that MB can bind with DNA sequences in at least three different manners; i) specific binding between MB and guanine bases,34 ii) intercalation of MB in the DNA double helix,35 and iii) electrostatic interaction between anionic DNA and cationic MB.36 Here, MB cations act as a signalling molecule that stoichiometrically binds to the guanine bases of miRNA and quantitatively indicates the amount of miRNA localized at the electrode surface.

In this method, the nanocomposite (AuNP-Fe2O3NC) offers more sensitive detection using catalytic properties of mesoporous Fe2O3 and higher adsorption of miRNA onto coated gold NPs, and this proof-of-concept method can detect the pico-molar level of miRNA.21 However, its urgent demand for detecting ultra-low such as attomolar level detection of clinically relevant miRNA as some clinical samples may contain a very tiny amount of miRNAs.

245

Figure 6.4: Schematic representation of the assay. Exosomal RNA was isolated from exosome that derived from preconditioned media of motor neurons (ALS). Target miRNA was hybridized with capture probe, and biotinylated magnetic beads were then added to the hybrid for magnetic isolation and purification. After magnetic purification, target miRNA were released from magnetic beads and allowed to adsorb onto the gold NPs of AuNP-

Fe2O3NC/SPCE surface for electrochemical detection. The presence of adsorbed miRNA interact with positively charged MB redox molecules, and the amount of charge intercalation 3- is measured by CC readout in the presence of 4mM [Fe(CN)6] in 40mM Tris-HCl buffer (pH 7.4). In the electrocatalytic process, electrons flow from the electrode surface to intercalated MB+ in an RNA-mediated reaction. The reduced form of MB+, leucomethylene blue (LB+), in turn, reduces solution-borne ferricyanide so that more electrons can flow to MB+ and the catalytic cycle continues.

To generate a high electrocatalytic signal amplification, the MB system was coupled to 3- 3- [Fe(CN)6] system (shown in equation 1 and 2). [Fe(CN)6] in the solution-phase further 3- triggers the electrocatalytic reduction of MB. This is because as the [Fe(CN)6] is a relatively stronger oxidant, it oxidized MB for the regeneration of leuco-methylene blue (LMB) from MB allowing multiple turnovers of MB resulting in a drastic increase in the signal.37 It is also important to note that, previously Boon et al. has shown that the negatively charged ferricyanide gave no electrochemical response at the DNA-modified electrode; however in presence tiny amount (micro-molar) of MB resulted in the voltammetric peak currents near

246

those expected for the reduction of ferricyanide at the diffusion-controlled limit. Therefore, LMB release from the miRNA attached-surface appears to be a key element in the overall 3- 38 mediated reduction of [Fe(CN)6] . Thus, the amount of CC charge generated by MB and 3- [Fe(CN)6] system should have a clear correlation with the concentration of miRNA. MB+ + 2e- LB+ (1) + 3- - + LB + [Fe(CN)6] [Fe(CN)6]4 + MB (2)

6.3.3. Assay Specificity To check the assay functionality and specificity, the sequence of miR-338-3p had been cast- off as a target miRNA, and a series of control samples were interrogated. As can be seen from Figure 6.5, the bare SPCE generated a very tiny amount of charge as there was no electroactive

AuNP-Fe2O3NC or miRNA was absorbed for MB intercalation. The control samples (PBS -2 were used instead of miRNA) generated 5.2 µC cm , whereas the Qdl (absence of MB) generated 4.6 µC cm-2 of current. The almost double current response of control than bare SPCE is probably coming from the electrocatalytic effect AuNP-Fe2O3NC attached to SPCE surface.

The slight reduction of current response of Qdl than control sample is probably due to the blockage of some catalytic site by adsorbed miRNA, but no charge from miRNA was observed as because in the absence of miRNA no interaction had been occurring and hence no current associated from miRNA. A very similar response was attributed when non-complementary (to target miRNA thus capture probe) sequence was chosen instead of target miR-338-3p. This is because no miR-338-3p were isolated and hence no adsorption of miRNA onto the surface of

AuNP-Fe2O3NC. However, a minuscule increase of current is due to the very tiny amount MB - adsorbed onto AuNP-Fe2O3NC surface. A substantial amount of current (4.6 vs. 16.1 µC cm 2) had resulted when the same amount (10 pM) of miR-338-3p was interrogated onto the sensor surface, presenting the specificity of the sensor. This is because miR-338-3p specific-capture probe isolated the significant amount of target miRNA and allowed to absorb onto the sensor, and hence MB intercalation generated the current response. Surprisingly boosted amount of 3- current has been obtained when the sensor was integrated with [Fe(CN)6] system. With the same amount of starting miRNA (10 pM), the redox cycling system-generated almost 3 times of higher current responses (45.16 vs. 16.1 16.1 µC cm-2), which is due to the multiple turnovers 3- of MB by stronger oxidant [Fe(CN)6] , demonstrating the high specificity of our assay.

247

Figure 6.5: Assay specificity; CC (inset) showing the charge density obtained after assay was performed on different target sequences (SPCE/miR-338-3p, SPCE/NC/ miR-338-3p (control), SPCE/NC , SPCE/NC/miR-338-3p (wrong target), SPCE/NC/miR-338-3p/MB in 40mM Tris- 3- HCl buffer (pH 7.4) and SPCE/NC/miR-338-3p/MB in 4mM [Fe(CN)6] in 40mM Tris-HCl buffer (pH 7.4) for redox cycle. Concentration of miR-338-3p and miR-338-3p were 10 pM

(NC: AuNP-Fe2O3NC).

We envisage that along with high loading capacity and electrocatalytic activity of 3- AuNP-Fe2O3NC (towards MB, and/or MB/[Fe(CN)6] ) a few others features attributed this high specificity of our assay; (i) magnetic bead-based purifications provides purified targets miRNA for the sensor surface, (ii) the direct adsorption of miRNA on to the transduction surface revoke the tedious surface chemistry for nucleic acid immobilization, (iii) the clinical (plasma or serum) or spiked samples are spatially separated from the transduction surface, thereby avoids the adsorption of unwanted-species (e.g. proteins, non-targeted nucleic acids).

6.3.4. Assay sensitivity To evaluate the assay sensitivity, synthetic target miR-338-3p of different concentration 3- ranging from 1µM to 10 pM were initially detected without [Fe(CN)6] redox cycling. It has been observed that the charge generated by MB intercalation was augmented with increasing concentration of miRNA. This was attributed to the higher amount of miRNA was isolated and thus adsorbed on to the Au NPs of nanocube-attached SPCE surface. An increased amount of adsorbed miRNA contains a relatively higher amount of negatively charged phosphate

248

backbone, thus binding with a greater number of positively charged MB intercalator and thereby, generating higher charge (Figure 6.6a and c). The level of engendered redox charge clearly designates that our inexpensive nanocube attached SPCE can detect 10 fM miRNA with higher sensitivity. As stated earlier, with the increasing demand for detecting the ultra-low level of disease-specific miRNA, the electrochemical techniques have integrated more than one signal amplification step in a single assay platform.

Figure 6.6: Improved assay sensitivity; CC curves showing amount of charge generated to different concentration of starting miR-338-3p targets (a) before (a; a-control; b to g-10 fM to 3- 1 nM) and (b) after (b; a-control; b to I -100 aM to 1 nM) coupling with [Fe(CN)6] (redox cycle). The figure c and d show the analogous bar diagrams (inset; linear calibration plots) 3 before (and after coupling with [Fe(CN)6] respectively.

3- To achieve the ultra-low level of detection, we have coupled our assay with [Fe(CN)6] redox 3- cycling. The MB/[Fe(CN)6] redox cycling resulted in boosted charge amplification, which results in the detection of 100 aM level of miRNA (Figure 6.6b and d). This is because, MB first reduced (to LB+) at the electrode surface upon hybridization/intercalation with the target 3- + + miRNA sequence, which was further oxidized by [Fe(CN)6] and LB back to MB in the diffusion layer. This redox cycling ensuing in the sharp upsurge in the electron flux produces 249

an enhanced CC readout. The limit of detection (LOD) was estimated to be 100 aM (signal to noise ratio 3.0), and the response is clearly distinguishable from the control. It is noteworthy to state that prior to redox cycle (only MB), the assay can detect up to 10 fM, which was improved to 100 aM (hundred times more sensitive) after redox cycling (electro-catalytic cycle 3- between MB and [Fe(CN)6] ) (Figure 6.6). Though there is some nanotechnology (nanowire, carbon nanotube), based methods have been reported with similar detection sensitivity or even lower range of miRNA,11 but our methods offer several advantages. The three steps amplifications; (i) AuNP-Fe2O3NC-modified SPCE electrode provides electrocatalytic signal amplification (through the catalytic redox reaction of MB),21 (ii) specific binding capacity of redox intercalator MB with surface-attached target miRNA, provides precise signals,35 and (iii) 3- the electrocatalytic redox cycling (MB/[Fe(CN)6] ) boosted the catalytic signal amplification.37 Besides, our sensor design is relatively simple (non-enzymatic, external magnet-based AuNP-Fe2O3NC attachment, direct adsorption of target miRNAs, electrocatalytic signal amplification), inexpensive (reusable AuNP-Fe2O3NC-modified SPCE, easy synthesis of AuNP-Fe2O3NC, enzymatic amplification or labelling free.

6.3.5. Analysis of miR-338-3p in exosomes from ALS patient and healthy control motor neurons To examine whether this nanostructures based assay could be applied to clinical (patient) samples of ALS, we interrogated our method in exosomal RNA sample extracted from preconditioned media of motor neurons (ALS). Exosomes are nanovesicles consisting of different functional proteins and genetic materials, e.g. mRNA and miRNA, which are secreted from many types of cells.39 Exosomes work as an intercellular communication vesicle, and they guide axonal development, modulate synaptic activity in the nervous system and help regenerate peripheral nerve tissues.40 Xin et al. testified that exosomes derived from MSC promote neurogenesis, neurite remodelling, and functional recovery after stroke. Furthermore, they demonstrated that the transfer of miR-133b from mesenchymal stem cells (MSCs) to neurons and astrocytes via MSC-derived exosomes promotes the neurological recovery.41 miRNA is also known to affect cell growth and direct differentiation of stem cells into many types of cells, including neurons.42

250

Figure 6.7: Assay performance on clinical samples; (a) Electrochemical signal obtained from sample C1, C2, and P1 to P3 in week 2 and week 4 intervals, where C1 and C2 represent the motor neuron samples from healthy patients and P1, P2 and P3 represent motor neuron samples from ALS patients. (b) RT-qPCR validation of exosomal miR-338-3p expression levels in the two healthy control samples and three ALS samples over the two and four weeks of motor neuron differentiation. Below: Corresponding p values obtained using two-way ANOVA comparing the control (healthy) samples and ALS patient’s samples.

In this proof of assay, we have chosen mir-338-3p, which was extracted from exosomes obtained from preconditioned media of motor neurons. It has been reported that miR-338-3p is over-expressed in blood, CFS, serum and spinal cord from sporadic ALS patients and has been considered as a potential biomarker for ALS diagnosis.15, 43 Herein, we have analysed two healthy patient samples (named control; C1 and C2) and three ALS samples (P1, P2 and P3) in 2 and 4 weeks of motor neuron differentiation. As can be from Figure 6.7a, our assay enabled detection of miR-338-3p from healthy and patient motor neuron exosomes. The data showed that the miR-338-3p levels were expressed differently in different samples (Figure 6.7a). With the same amount of starting cell culture media, our method detected the lower expression of miR-338-3p in one of the ALS samples (P1) compared to those of two controls and two ALS samples. Figure 6.7b represents the adjusted p values obtained using two-way ANOVA comparing the control (healthy) samples and ALS patient samples. We validated our proof-of- assay performance with a standard RT-qPCR method (Figure 6.7), which shows consistent results with the electrochemical measurements. However, control C1 and ALS sample P3 shows no significant difference in expression of miR-338-3p. In future studies, we will address

251

these differences and evaluate the effects of time during differentiation with miR-338-3p as well as the combination of multiple miRNAs on ALS diagnosis.

6.4. Conclusions In the current study, we have designed a nanocomposite, studied its catalytic properties and applied successfully to develop a simple, inexpensive method for detecting the level of miRNA expression based on adsorption affinity interaction of miRNA nucleobases with gold. We directly adsorbed miRNA onto the AuNP-Fe2O3NC attached on SPCE and amount of miRNA adsorbed was then quantified via CC interrogation of the RNA-bound MB complexes. The method is robust, fast (<2.5 hr), sensitive and potentially applicable to a wide variety of miRNA detection. Most notably, our developed assay can successfully quantify miRNA level from motor neuron exosomes. This method offers some distinct advantages- (i) catalytic activity of highly porous framework of AuNP-Fe2O3NC significantly enhances the detection sensitivity,

(ii) direct adsorption of target miRNA on a AuNP-Fe2O3NC/SPCE surface (it substantially simplifies the overall sensing approach by avoiding the use of complicated chemistries underlying each step of the conventional recognition and transduction layers based iRNA biosensors), (iii) use of magnetic bead-based isolation and purification steps reduces the matrix effect as non-targets species can be removed via magnetic washing and enhances the isolation purity and efficiency, (iv) disposable screen-printed electrode for portable detection of serum mi-RNA at a relatively low cost (AUD$3 per electrode), and (v) electrochemical detection that can complement with the portable and inexpensive detection platform, and thus highly potential to translate the method into a simple and affordable screening of miRNA in clinical samples. Importantly, the assay can detect ultra-low amount (100 aM) of miRNA present in the motor neuronal exosomes. We foretell that the assay could also be radially extended to other miRNA marker detection through the further design of nanostructures and adaption of capture probe sequences.

252

6.5. References 1. Traxinger, K.; Kelly, C.; Johnson, B. A.; Lyles, R. H.; Glass, J. D., Prognosis and epidemiology of amyotrophic lateral sclerosis: analysis of a clinic population, 1997– 2011. Neurology: Clinical Practice 2013, 3 (4), 313-320. 2. Kiernan, M. C., Motor neuron disease in 2017: Progress towards therapy in motor neuron disease. Nature Reviews Neurology 2018, 14 (2), 65. 3. Sathasivam, S., Motor neurone disease: clinical features, diagnosis, diagnostic pitfalls and prognostic markers. Singapore Med J 2010, 51 (5), 367-373. 4. Puls, I.; Jonnakuty, C.; LaMonte, B. H.; Holzbaur, E. L.; Tokito, M.; Mann, E.; Floeter, M. K.; Bidus, K.; Drayna, D.; Oh, S. J., Mutant dynactin in motor neuron disease. Nature Genetics 2003, 33 (4), 455. 5. Dharmadasa, T.; Henderson, R. D.; Talman, P. S.; Macdonell, R. A.; Mathers, S.; Schultz, D. W.; Needham, M.; Zoing, M.; Vucic, S.; Kiernan, M. C., Motor neurone disease: progress and challenges. Medical Journal of Australia 2017, 206 (8), 357-362. 6. Kwiatkowski, T. J.; Bosco, D.; Leclerc, A.; Tamrazian, E.; Vanderburg, C.; Russ, C.; Davis, A.; Gilchrist, J.; Kasarskis, E.; Munsat, T., Mutations in the FUS/TLS gene on chromosome 16 cause familial amyotrophic lateral sclerosis. Science 2009, 323 (5918), 1205-1208. 7. Kabashi, E.; Valdmanis, P. N.; Dion, P.; Spiegelman, D.; McConkey, B. J.; Velde, C. V.; Bouchard, J.-P.; Lacomblez, L.; Pochigaeva, K.; Salachas, F., TARDBP mutations in individuals with sporadic and familial amyotrophic lateral sclerosis. Nature Genetics 2008, 40 (5), 572. 8. Haramati, S.; Chapnik, E.; Sztainberg, Y.; Eilam, R.; Zwang, R.; Gershoni, N.; McGlinn, E.; Heiser, P. W.; Wills, A.-M.; Wirguin, I., miRNA malfunction causes spinal motor neuron disease. Proceedings of the National Academy of Sciences 2010, 107 (29), 13111-13116. 9. Buratti, E.; De Conti, L.; Stuani, C.; Romano, M.; Baralle, M.; Baralle, F., Nuclear factor TDP‐43 can affect selected microRNA levels. The FEBS journal 2010, 277 (10), 2268-2281. 10. Islam, M. N.; Masud, M. K.; Haque, M. H.; Hossain, M. S. A.; Yamauchi, Y.; Nguyen, N. T.; Shiddiky, M. J., RNA biomarkers: diagnostic and prognostic potentials and recent developments of electrochemical biosensors. Small Methods 2017, 1 (7), 1700131.

253

11. Masud, M. K.; Umer, M.; Hossain, M. S. A.; Yamauchi, Y.; Nguyen, N.-T.; Shiddiky, M. J., Nanoarchitecture frameworks for electrochemical miRNA detection. Trends in Biochemical Sciences 2019. 12. Quan, Z.; Zheng, D.; Qing, H., Regulatory roles of long non-coding RNAs in the central nervous system and associated neurodegenerative diseases. Frontiers in Cellular Neuroscience 2017, 11, 175. 13. Maggio, S.; Ceccaroli, P.; Polidori, E.; Cioccoloni, A.; Stocchi, V.; Guescini, M., Signal Exchange through Extracellular Vesicles in Neuromuscular Junction Establishment and Maintenance: From Physiology to Pathology. International Journal of Molecular Sciences 2019, 20 (11), 2804. 14. Takahashi, I.; Hama, Y.; Matsushima, M.; Hirotani, M.; Kano, T.; Hohzen, H.; Yabe, I.; Utsumi, J.; Sasaki, H., Identification of plasma microRNAs as a biomarker of sporadic Amyotrophic Lateral Sclerosis. Molecular Brain 2015, 8 (1), 67. 15. De Felice, B.; Annunziata, A.; Fiorentino, G.; Borra, M.; Biffali, E.; Coppola, C.; Cotrufo, R.; Brettschneider, J.; Giordana, M. L.; Dalmay, T., miR-338-3p is over- expressed in blood, CFS, serum and spinal cord from sporadic amyotrophic lateral sclerosis patients. Neurogenetics 2014, 15 (4), 243-253. 16. De Felice, B.; Guida, M.; Guida, M.; Coppola, C.; De Mieri, G.; Cotrufo, R., A miRNA signature in leukocytes from sporadic amyotrophic lateral sclerosis. Gene 2012, 508 (1), 35-40. 17. Boriachek, K.; Islam, M. N.; Möller, A.; Salomon, C.; Nguyen, N. T.; Hossain, M. S. A.; Yamauchi, Y.; Shiddiky, M. J., Biological functions and current advances in isolation and detection strategies for exosome nanovesicles. Small 2018, 14 (6), 1702153. 18. Zhao, Y.; Chen, F.; Li, Q.; Wang, L.; Fan, C., Isothermal amplification of nucleic acids. Chemical Reviews 2015, 115 (22), 12491-12545. 19. Shi, R.; Chiang, V. L., Facile means for quantifying microRNA expression by real-time PCR. Biotechniques 2005, 39 (4), 519-525. 20. Kilic, T.; Erdem, A.; Ozsoz, M.; Carrara, S., microRNA biosensors: opportunities and challenges among conventional and commercially available techniques. Biosensors and Bioelectronics 2018, 99, 525-546. 21. Masud, M. K.; Islam, M. N.; Haque, M. H.; Tanaka, S.; Gopalan, V.; Alici, G.; Nguyen, N.-T.; Lam, A. K.; Hossain, M. S. A.; Yamauchi, Y., Gold-loaded nanoporous

254

superparamagnetic nanocubes for catalytic signal amplification in detecting miRNA. Chemical Communications 2017, 53 (58), 8231-8234. 22. Islam, M. N.; Gorgannezhad, L.; Masud, M. K.; Tanaka, S.; Hossain, M. S. A.; Yamauchi, Y.; Nguyen, N. T.; Shiddiky, M. J., Graphene‐Oxide‐Loaded Superparamagnetic Iron Oxide Nanoparticles for Ultrasensitive Electrocatalytic Detection of MicroRNA. ChemElectroChem 2018, 5 (17), 2488-2495. 23. Islam, M. N.; Masud, M. K.; Nguyen, N.-T.; Gopalan, V.; Alamri, H. R.; Alothman, Z. A.; Al Hossain, M. S.; Yamauchi, Y.; Lamd, A. K.; Shiddiky, M. J., Gold-loaded nanoporous ferric oxide nanocubes for electrocatalytic detection of microRNA at attomolar level. Biosensors and Bioelectronics 2018, 101, 275-281. 24. Yadav, S.; Masud, M. K.; Islam, M. N.; Gopalan, V.; Lam, A. K.-y.; Tanaka, S.; Nguyen, N.-T.; Al Hossain, M. S.; Li, C.; Yamauchi, M. Y., Gold-loaded nanoporous iron oxide nanocubes: a novel dispersible capture agent for tumor-associated autoantibody analysis in serum. Nanoscale 2017, 9 (25), 8805-8814. 25. Masud, M. K.; Yadav, S.; Islam, M. N.; Nguyen, N.-T.; Salomon, C.; Kline, R.; Alamri, H. R.; Alothman, Z. A.; Yamauchi, Y.; Hossain, M. S. A., Gold-loaded nanoporous ferric oxide nanocubes with peroxidase-mimicking activity for electrocatalytic and colorimetric detection of autoantibody. Analytical Chemistry 2017, 89 (20), 11005- 11013. 26. Boriachek, K.; Masud, M. K.; Palma, C.; Phan, H.-P.; Yamauchi, Y.; Hossain, M. S. A.; Nguyen, N.-T.; Salomon, C.; Shiddiky, M. J., Avoiding Pre-Isolation Step in Exosome Analysis: Direct Isolation and Sensitive Detection of Exosomes Using Gold- Loaded Nanoporous Ferric Oxide Nanozymes. Analytical Chemistry 2019, 91 (6), 3827-3834. 27. Engel, M.; Balez, R.; Muñoz, S. S.; Cabral-da-Silva, M. C.; Stevens, C. H.; Bax, M.; Do-Ha, D.; Sidhu, K.; Sachdev, P.; Ooi, L., Viral-free generation and characterization of a human induced pluripotent stem cell line from dermal fibroblasts. Stem Cell Research 2018, 32, 135-138. 28. Muñoz, S. S.; Balez, R.; Berg, T.; Engel, M.; Bax, M.; Do-Ha, D.; Stevens, C. H.; Greenough, M.; Bush, A.; Ooi, L., Generation and characterization of human induced pluripotent stem cell lines from a familial Alzheimer's disease PSEN1 A246E patient and a non-demented family member bearing wild-type PSEN1. Stem Cell Research 2018, 31, 227-230.

255

29. Zeineddine, R.; Pundavela, J. F.; Corcoran, L.; Stewart, E. M.; Do-Ha, D.; Bax, M.; Guillemin, G.; Vine, K. L.; Hatters, D. M.; Ecroyd, H., SOD1 protein aggregates stimulate macropinocytosis in neurons to facilitate their propagation. Molecular Neurodegeneration 2015, 10 (1), 57. 30. Bilican, B.; Livesey, M. R.; Haghi, G.; Qiu, J.; Burr, K.; Siller, R.; Hardingham, G. E.; Wyllie, D. J.; Chandran, S., Physiological normoxia and absence of EGF is required for the long-term propagation of anterior neural precursors from human pluripotent cells. PLoS One 2014, 9 (1), e85932. 31. Koo, K. M.; Carrascosa, L. G.; Shiddiky, M. J.; Trau, M., Amplification-free detection of gene fusions in prostate cancer urinary samples using mrna–gold affinity interactions. Analytical Chemistry 2016, 88 (13), 6781-6788. 32. Koo, K. M.; Carrascosa, L. G.; Shiddiky, M. J.; Trau, M., Poly (A) extensions of miRNAs for amplification-free electrochemical detection on screen-printed gold electrodes. Analytical Chemistry 2016, 88 (4), 2000-2005. 33. Steel, A. B.; Herne, T. M.; Tarlov, M. J., Electrochemical quantitation of DNA immobilized on gold. Analytical Chemistry 1998, 70 (22), 4670-4677. 34. Tuite, E.; Norden, B., Sequence-specific interactions of methylene blue with polynucleotides and DNA: a spectroscopic study. Journal of the American Chemical Society 1994, 116 (17), 7548-7556. 35. Liu, M.; Luo, C.; Peng, H., Electrochemical DNA sensor based on methylene blue functionalized polythiophene as a hybridization indicator. Talanta 2012, 88, 216-221. 36. Wang, Y.; Zhou, A., Spectroscopic studies on the binding of methylene blue with DNA by means of cyclodextrin supramolecular systems. Journal of Photochemistry and Photobiology A: Chemistry 2007, 190 (1), 121-127. 37. Boon, E. M.; Ceres, D. M.; Drummond, T. G.; Hill, M. G.; Barton, J. K., Mutation detection by electrocatalysis at DNA-modified electrodes. Nature Biotechnology 2000, 18 (10), 1096. 38. Boon, E. M.; Barton, J. K.; Bhagat, V.; Nersissian, M.; Wang, W.; Hill, M. G., Reduction of ferricyanide by methylene blue at a DNA-modified rotating-disk electrode. Langmuir 2003, 19 (22), 9255-9259. 39. Takeda, Y. S.; Xu, Q., Neuronal differentiation of human mesenchymal stem cells using exosomes derived from differentiating neuronal cells. PLoS One 2015, 10 (8), e0135111.

256

40. Lai, C. P.-K.; Breakefield, X. O., Role of exosomes/microvesicles in the nervous system and use in emerging therapies. Frontiers in Physiology 2012, 3, 228. 41. Xin, H.; Li, Y.; Buller, B.; Katakowski, M.; Zhang, Y.; Wang, X.; Shang, X.; Zhang, Z. G.; Chopp, M., Exosome‐mediated transfer of miR‐133b from multipotent mesenchymal stromal cells to neural cells contributes to neurite outgrowth. Stem Cells 2012, 30 (7), 1556-1564. 42. Loohuis, N. O.; Kos, A.; Martens, G.; Van Bokhoven, H.; Kasri, N. N.; Aschrafi, A., MicroRNA networks direct neuronal development and plasticity. Cellular and Molecular Life Sciences 2012, 69 (1), 89-102. 43. Ricci, C.; Marzocchi, C.; Battistini, S., MicroRNAs as biomarkers in amyotrophic lateral sclerosis. Cells 2018, 7 (11), 219.

257

Chapter 7

Mesoporous Gold Biosensor for Electrochemical Detection of MicroRNA at Attomolar Level*

*Sections of this chapter are based on Masud, M.K., et al., ACS Applied Materials and Interfaces, 2020 (submitted).

258

7. Mesoporous Gold Biosensor for Electrochemical Detection of MicroRNA at Attomolar Level

7.1 Introduction MicroRNAs (miRNAs) are short (19-25 nucleotides in length), endogeneous, noncoding RNA molecules, presently gaining enormous interest as a cancer biomarker. miRNAs bind to the 3′ untranslated region (3'-UTR) of targeted messenger RNAs (mRNAs) to inhibit translation, functioning as oncogenes and tumour suppressors, thereby regulate post-transcriptional gene expression and remodelling of the epigenome (e.g., DNA methylation and histone modification).1-3 They play crucial roles in cancer initiation and progression through their altered expression levels, single-nucleotide polymorphism, copy number variations and mutations. Moreover, miRNAs are available and stable in easily accessible bodily fluids such as urine, saliva, and blood, presenting them as minimally invasive or invasive biomarkers for diagnosis and prognosis of various diseases including cancer.4-5 The prominent functional insights, therefore, triggered widespread development of advanced strategies for quantitative detection of miRNAs in clinical practice. RNA sequencing (RNA-seq), microarrays, quantitative real-time PCR (qRT-PCR) have widely been utilized for the analysis of miRNA.2, 6-7 Despite their reliability, good sensitivity and specificity, they are yet suffering from PCR amplification bias, fluorescent labelling, lean on complicated and upscale protocols.7 Moreover, these methods are entirely confined in a sophisticated laboratory with the requirement of expensive instrumentation. Substitute for these techniques, a wide variety of biosensors has been emerged based on hybridisation, oligonucleotides labelled with different enzymes and redox indicator. Most of these sensors generally involve complicated and monotonous amplification procedures, affluent biomolecules, time-consuming and complicated procedures.3 It is noteworthy to mention that regardless of these advances, the miRNA detection technologies have yet to accomplish their translation into routine clinical practice specifically in resource-limited settings, somewhat due to the lack of sensitive and specific analysis in a portable, inexpensive point-of-care detection system. Electrochemical strategies have been shown to offer rapid, sensitive and accurate miRNA detection exclusive of any preceding amplification process.8-9 However, the transduction surface needs to be functionalized with a probe DNA or aptamer, require to employ signalling probe, different redox moiety to get high sensitivity. One of the effective ways to replace that cumbersome functionalization and signal amplification is to integrate

259

engineered nanostructure that could function as both the signal transducer as well as a signal amplifier.3,8 Advancement of nanoarchitonics offers a wide range of size- and shape-controlled mesoporous/nanoporous structure for designing sensitive biosensors.1, 10 Despite the broad applicability of mesoporous metals nanostructure in many applications including batteries, fuel cells, solar cells, chemical sensors and photonic devices, mesoporous films have appealed much attention for obtaining advanced biodevices such protein microarray, bio-photoelectrode, encapsulation of cells, biosensor.11-12 In biosensor, they can be integrated as an electrical signal transducer, like a matrix or a selective capturing platform for proteins, nucleic acids, enzymes and cells, and as a biointerface within an ordered porous structure for obtaining a high selectivity towards harvesting and diffusion.3,11 Having small pore size, high surface-to-volume ratio, biocompatibility, and intrinsic electrochemical (EC) properties, mesoporous metallic nanostructures can overcome the barriers of structural miniaturization of diagnostics, steering to the possibility of developing inexpensive, sensitive portable devices.8, 13-14 Mesoporous metal architecture with electro-conductive framework results in superior signal transduction by providing pore-induced high surface area for uptaking of biological molecules and redox species, hybridisation moiety for the efficient and faster analyte (probe or target) binding, and electrocatalytic cascade signal amplification.14-16 Moreover, mesoporous structure (as electrode material) offers several advantages for improving the performance of electrodes; i) it has dramatically increased the electrochemically active surfaces, ii) it reduces the amplitude of the noise and achieved the high signal-to-noise ratio by lowering the impedance of the electrode, and iii) it can reduce the electrode size, which enables placing of multiple electrodes in a high- density array for advanced spatial resolution.17 Au has attained immense attention in biosensing due to its inherent bio-favourability, conductivity, chemical and physical stability and predominantly bio-inspired unique surface chemistry.18-19 The use of planar Au surfaces or Au nanoparticles (NPs) holds great promise for high-throughput electrochemical or optical detection of biomolecules on a single device or microfluidic devices for multiplexed detection with nominal sample supplies. Mirkin and Rothberg's groups have revealed that DNA can be directly adsorbed on the Au surfaces and this interaction is DNA sequence-dependent which could occur through the typical chemisorption and physisorption mechanism.20-21 This comprises direct interaction of nitrogen atoms of DNA nucleobase rings with Au as well as the partial contribution from the exocyclic amino group and charge transfer between the aromatic ring and Au surface.22 Our target in this

260

study is to realise the ultrasensitive detection of miRNA using a mesoporous Au electrode (MPGE). Although mesoporous metals have been prepared by soft- and hard-templating approaches, MPGE with well-defined pores has not been achieved due to the difficulty in controlling the crystal growth of Au. To realise this preparation, stable block copolymers with long and rigid hydrophobic chains have aroused more interest.23-24 Herein, we detail a new class of MPGE fabrication for biosensor by using block (di) polymeric-micelles (polystyrene-block- polyoxyethylene; PS-b-PEO) and demonstrate amplification and label-free ultrasensitive detection of microRNA which have not been realized yet. Our highly active MPGE shows excellent potential towards direct adsorption of miRNA and succeeding electrochemical interrogation (through 3-/4- differential pulse voltammetry, DPV) of adsorbed miRNA in the presence of [Fe(CN)6] redox system. The MPGE-based single-step strategy has attained a wide dynamic linear range (100 aM to 1 nM) with an ultra-low limit detection of 100 aM without any form of amplification or surface-modification steps.

Figure 7.1: (a) Schematic representation of the preparation of mesoporous Au electrode (MPGE) via electrodeposition of gold (III)-containing polymeric (block) micelles. The polymeric micelles are formed by dissolving polystyrene-b-poly(ethylene oxide)) (PS-b-PEO) diblock copolymer in THF followed by the addition of ethanol and aqueous gold (III) chloride

261

solution; and. (b) Schematic representation of electrochemical detection of miRNA, where target miR-9-2 are isolated and purified by using a complementary probe-bound dynabead followed by the adsorption at MPGE surface and DPV interrogation in the presence of 3- 4- [Fe(CN)6] / redox system. The magnetically isolated and purified miRNA (miR-9-2) are directly adsorbed onto the MPGE via RNA-Au affinity, which follows conventional physisorption and chemisorption mechanism. Using the DPV technique in the presence of 3-/4- [Fe(CN)6] redox system, we can precisely estimate the exact concentration of miR-9-2.

7.2. Experimental

7.2.1. Reagents and materials

Unless otherwise stated, the reagents and chemicals used for the conducting experiments were of analytical grade. Gold(III) chloride, tetrahydrofuran (THF), potassium hexacyanoferrate(II), potassium hexacyanoferrate(III) and phosphate buffer saline (PBS) tablet (0.01 M phosphate buffer, 0.0027 M potassium chloride and 0.137 M sodium chloride, pH 7.4 at 25 oC) were purchased from Sigma-Aldrich (Australia). Polystyrene–Poly(ethylene oxide) (PS-b-PEO) diblock Copolymer PS-b-PEO were obtained from Polymer Source Inc. (USA). Tris was purchased from VWR Life science (Australia). All chemicals and reagents were used as received without additional purification. UltrapureTM DNase/RNase-free distilled water (Invitrogen, Australia) was used throughout the experiments. Oligonucleotides were acquired from Integrated Technologies, USA and sequences are shown in table 7.1.

Table 7.1: Oligonucleotide sequences (miR-9-2) Oligos 5 ́-Sequences-3 ́

Biotinylated miR-9-2 capture TCA TAC AGC TAG ATA ACC AAA GA/3Biotin/ probe

Synthetic miR-9-2 sequence UCU UUG GUU AUC UAG CUG UAU GA

7.2.2 Instrumentations Scanning electron microscope (SEM) images were taken with a JEOL JSM-7800F scanning microscope with the accelerating voltage of 10 kV. The elemental chemical analysis of the nanocubes was performed by X-ray photoelectron spectroscopy (Kratos Axis Ultra XPS,

262

CMM, The University of Queensland). All samples were degassed in a vacuum before carrying out the measurements. Small-angle neutron scattering (SANS) measurements were performed at the Australian Nuclear Science and Technology Organisation on the Quokka instrument. Neutrons of 5 Å wavelength were used with the sample to detector distances of 8 and 1.3 m. Samples were corrected for background measurements and placed on an absolute scale using standard procedures. All electrochemical measurements were performed with a CHI650 electrochemical workstation. (CH Instrument, USA). Cyclic voltammetry (CV) and chronoamperometry experiments were conducted in a single-compartment cell with a 3-mL volume. A conventional three-electrode system, comprising MPGE, a platinum auxiliary electrode, and an Ag/AgCl3 1.0 M NaCl reference electrode (CH Instrument, Inc. USA), was used for the measurement of electrocatalytic activity. Differential pulse voltammetric (DPV) experiments were recorded at −0.1 to −0.5 V with a pulse amplitude of 50 mV and a pulse width of 50 ms in 10 mM PBS solution containing 2 mM [K3Fe(CN)6] and 2 mM [K4Fe(CN)6] electrolyte solution.

7.2.3 Synthesis and characterisation of the mesoporous gold electrode (MPGE) The MPGE were prepared by electrodeposition in the presence of the self-assembled polymeric micelles containing gold(III) (Au3+) ions as electrolyte onto a gold (sputtered) substrate. The PS (18,000)-b-PEO (7,500) polymeric micelles were acted as pore directing agent and Au3+ as a precursor. The electrolyte solutions were prepared by dissolving 10 mg of PS-b-PEO in 3 mL of tetrahydrofuran (THF) at 40 °C. After adding 1 mL of ethanol, aqueous HAuCl4 (40 mM) solution were slowly incorporated under constant stirring. 2.5 mL deionized H2O was then added to form the spherical micelles followed by the gentle stirring for 30 min at room temperature until complete dissolution of the Au precursors. The successive addition of aqueous HAuCl4 enabled the formation of spherical micelles of PS-b-PEO based on the less solubility of PS core in the water (Figure 7.1a). To understand the formation of the polymeric micelle, FT-IR were recorded during micelle formations and Au ion complexation (Figure 7.2). PS-b-PEO (THF) exhibits characteristics peaks at 3025.50 and 2919.09 cm-1 for aliphatic chain; at 1603.72, 1458.33, 1494.55, 725.11 and 692.00 cm-1 (phenyl ring) for PS and at 1080.64 cm-1 (-C-O-C-) for PEO moiety. Upon the complexation with gold ion, all peaks are experienced a negative shift. A new broad peak arises at 3369.08 cm-1 (hydroxyl group absorption maxima), which probably due to the formation of hydrogen bonding between PEO and gold ions (Figure 7.2).

263

Figure 7.2: FT-IR spectra during PS-b-PEO micellization with Au.

To further confirm the complexation, shape and structure of the micelles, Small-angle neutron scattering (SANS) was employed using the Bilby instrument at ANSTO.25 The instrument was used in velocity selector mode using neutrons of wavelength 6 Å with detectors positioned at 16 m (rear), 2.5 m (horizontal curtains) and 1.5 m (vertical curtains). Data were reduced and put on an absolute scale relative to the direct beam using Mantid26 and then solvent subtracted and modelled using the Igor Pro macros from NIST.27 The micelles containing PS- b-PEO was measured at room temperature under two conditions; (i) the block polymer dissolved in THF and deuterated ethanol, and (ii) aqua-HAuCl4 was added in the polymeric solution (THF and deuterated ethanol). Small-angle neutron scattering data are presented for both samples in the figure below. The scattering data from (i) are consistent with the sample staying dispersed in the monomeric form, and can be adequately fit with the Debye function representing a linear polymer chain in dilute solution (Figure 7.3a). A radius of gyration of 4 nm was extracted from the fit to the Debye function of the data in Figure 7.3a. Upon the addition of aqua-HAuCl4, the small-angle scattering data show the presence of significantly larger particles (spherical micelles) are formed in solution as the polymer self-assembles (Figure 7.3b). The data presented for (ii) in figure b can also be adequately fit with a polydisperse sphere with the radius gyration ~ 13.5 nm (Figure 7.3b).

264

Figure 7.3: Small-angle neutron scattering (SANS) patterns of two types of polymeric micelle solutions with corresponding debye fits. Sample 1 (a) was prepared by mixing PS-b-PEO in THF followed by the addition of ethanol; sample (b) was obtained by mixing PS-b-PEO in

THF followed by the addition of ethanol and HAuCl4 solution. The as-prepared micelle solution was used as precursor solution (electrolyte) directly used for electrodeposition. The electrochemical deposition was carried out using a conventional three-electrode system at a constant applied potential of −0.5 V (vs Ag/AgCl) for 1000 s without stirring by using an electrochemical workstation. The electrodeposition was performed in a standard three-electrode cell system with a gold-coated silicon wafer substrate, typical area of 0.09 cm2 (0.3 cm × 0.3 cm), as the working electrode, Ag/AgCl (saturated KCl) as the reference electrode, and a platinum wire as the counter electrode. The optimal electrodeposition of Au was carried out at room temperature at a constant potential of –0.5 V (vs. Ag/AgCl) for 1,000 s without stirring. During the electrodeposition, a stable current was detected corresponding to the Au reduction (Figure 7.4).

265

Figure 7.4. Amperometric (i-t) plots for deposition of MPGE under a typical electrolyte containing PS18000-b-PEO7500 micelles, THF, ethanol and gold (III) solution.

After the Au deposition, it is crucial to remove altogether the polymer from the substrate, and it was removed by dissolving the substrate in THF at 40oC followed by rinsing with deionized water. To check the complete removal of the polymer we have recorded the IR- spectra before and after treating the substrate with THF. The corresponding peaks for PS-b- PEO are disappeared after the treatment with hot THF (Figure 7.5). The complete removal of the polymeric template gives rise to the MPGE. The uniformly distributed mesopores onto the top surface are seen in the scanning electron microscope (SEM) image (Figure 7.6).

Figure 7.5. FT-IR spectra before and after removal of PS-b-PEO template during the preparation of MPGE.

266

The SEM image displays the spherical pores of around 20 ± 5 nm with concave exteriors around the mesopores. X-ray photoelectronic spectroscopy (XPS) was conducted to know the Au oxidation state and surface chemistry (Figure 7.7). A doublet at 87.7 and 84.0 eV were obtained from the high-resolution scan of Au4f which are separated by 3.7 eV probably because of spin-orbit coupling. This characteristically represents the presence of Au(0) species in MPGE. Our block-polymeric template approach is highly advantageous over sol-gel-based fabrication methods (as it follows solvent evaporation for pre-forming ordered mesophases). Electrochemical deposition enables facile and precise control of Au crystal growth, controlled growth rate, fine-tuning of the electrode surface and pore geometry by simply adjusting the potential and size of block polymers.

Figure 7.6: The SEM image of the top-surface of MPGE

267

Figure 7.7: The high-resolution XPS spectra of Au-4f.

7.2.4. Calculation of electrochemical surface area (ECSA) and roughness factor The electrochemical surface area of MPGE was calculated using the charge associated with the reduction of gold oxide by integration as follows (this charge is directly proportional to the active surface area of mesoporus gold); 28

Charge (Q) = Area (cm2) / scan rate (Vs-1)

-2 -2 ECSA = Q (µC) / Qref (μCcm ), where Qref = 390 μCcm is a calibration factor for a polycrystalline gold electrode.29

The roughness factor (Rf) was calculated using the following formula;

Rf = ECSA (cm2) / Geometrical area (cm2)

7.2.5. Hybridization and magnetic isolation of target miRNA (miR-9-2) The target-miRNA is isolated and purified from the spiked sample as shown in Figure 7.8 following previously published protocol.30-31 Briefly, the spiked RNA (total RNA) sample adjusted to the required concentration in 5 μL of RNase-free water before mixing with 10 μL of 5× SSC buffer and 10 μL of 10 μM biotinylated capture probes. After heating the mixture at 60 oC for 2 min followed by incubation for 60 min at room temperature. For magnetic isolation of target miR-9-2, 5 μL of streptavidin-labeled Dynabeads MyOne Streptavidin C1 (Invitrogen) magnetic beads washed and resuspended in 10 μL of 2× B&W buffer. Then, the

268

resuspended magnetic beads were added to the prepared capture probes-target miRNA hybrid. After 30 min incubation, magnetic bead-hybrid structure was washed and heat up at 95 oC to release target miRNA. The magnetic bead-capture probe was stuck by an external magnet and the released target miRNA was collected within a very short time. Before applying miRNA on to MPGE, the miRNA was diluted two times with 5×SSC buffer.

Figure 7.8: Magnetic isolation and purification of target miRNA using biotinylated capture probes and streptavidin-coated magnetic beads.

7.2.6. Electrochemical detection of adsorbed target miRNA 5.0 μL (diluted in 5X SSC buffer) sample was adsorbed on the MPGE surface. After 30 min of incubation, the electrode was washed (gently) with PBS (three times). The differential pulse voltammetric (DPV) experiments were recorded at -0.1 to -0.5 V with a pulse amplitude of 50 mV and a pulse width of 50 ms in 10 mM PBS solution containing 2 mM [K3Fe(CN)6] and 2 mM [K4Fe(CN)6] electrolyte solution before and after adsorbing target miRNA onto MPGE surface. The relative DPV current changes (i.e., %iRelative, per cent difference of the DPV signals generated for captured miRNA (imiRNA) with respect to the baseline current (iBaseline)) due to the adsorption of miRNA were then measured by using the following equation;

푖퐵푎푠푒푙푖푛푒− 푖푚푖푅푁퐴 %iRelative = × 100 푖퐵푎푠푒푙푖푛푒 where iBaseline and imiRNA are current density obtained for bare electrode and electrode after miRNA adsorption respectively.

7.3. Results and discussion

The MPGE was prepared through electrochemical reduction of Au(III) species present in the micelles containing PS-b-PEO block polymers (Figure 7.1a), where the polymeric micelles work as a pore directing agent for forming uniformly sized mesopores.32 However, the size of polymeric micelles is crucial as the size of pores and geometry is dependent on the size of block polymer. Therefore, the formation of spherical micelles was characterised by using small-angle

269

neutron scattering (SANS) (Figure 7.2) and infra-red spectroscopy (Figure 7.3). The SANS data for block polymer (dissolved in THF and deuterated ethanol) are consistent with the sample staying dispersed in the monomeric form, and the Debye function fit representing a linear polymer chain in dilute solution with a radius of gyration of 4 nm. Upon the addition of aqua-HAuCl4, the SANS data shows the presence of significantly larger particles (spherical micelles) are formed in solution and Debye function fit representing a polydisperse sphere with the radius gyration ~ 13.5 nm (For details, please see experimental section). Au-sputtered silicon was used as the substrate (as a working electrode) with a common area of 0.09 cm2 (0.3 cm × 0.3 cm). The film thickness can be controlled by varying the electrodeposition time (0 s to 2000 s). Both the electrochemical surface area (ECSA) and surface roughness is increased with increasing deposition time (Figure 7.9). The ECSA of MPGE was calculated using the charge associated with the reduction of gold oxide, determined by integration, which is proportional to the real active surface area of the mesoporous gold surface.28,29 The detailed synthesis and characterization are described in the experimental section (7.2).

Figure 7.9: Electrodeposition time versus ECSAs. (a) Cyclic voltammograms (CVs) in 0.5 M

H2SO4 of MPGE with different film thicknesses obtained by conducting electrodeposition in different timescale ranging from 0 s to 2000 s ; (b) the linear relationship between the ECSAs and the deposition time (film thickness), and (c) the roughness factor in response to deposition time. Mesoporous architecture facilitates the increased surface area that accelerates the redox 8,10 3-/4- reaction and accessibility of analyte. The redox response of [Fe(CN)6] system can be 270

utilized for quantification of surface-bound oligonucleotides on Au electrodes. Typically the adsorbed nucleic acids present on the electrode surface causes Coulombic repulsion of 3-/4- [Fe(CN)6] away from the surface, resulting in the significant lowering of current compared to a bare electrode.30, 33 The relative decrease in Faradaic current (bare vs. Oligoneucleotides- adsorbed electrode; % current response change) is inversely linked to the amount of adsorbed species. To understand the responses of prepared MPGE towards the redox process of 3-/4- [Fe(CN)6] system, herein, we have assessed the electrochemical activity of both MPGE and sputtered Au electrode (SGE) by measuring cyclic voltammetry (CV). Both electrodes generate 3-/4 a pair of well-defined sharp redox peaks in 2 mM [Fe(CN)6] , presenting a reversible single- electron transfer process (Figure 7.10a). Surprisingly, the MPGE give rise to significantly enhance cathodic (ipc), and anodic (ipa) peak currents. Notably, the ipc increases approximately -2 7.8-times (0.7456 vs. 0.098 mA cm ) with the Epc shifted by 61 mV, whereas the ipa increases -2 approximately 7.3-times (0.9415 vs. 0.13 mA cm ) with the Epa shifted by 47 mV. It has been demonstrated that the MPGE efficiently boosts both the oxidation and reduction process of 3-/4- [Fe(CN)6] system, owing to more exposed Au surface through the presence of abundant pores. Moreover, the porous structure experienced a negative shifting of both Epc and Epa 3-/4- associated with the redox reaction of [Fe(CN)6] is relative to that of SGE. This suggests a higher affinity as well as the interaction of MPGE towards the redox system.34 Similar responses were obtained from the DPV measurements (Figure 7.10b), where MPGE generated approximately 7-times higher current signal than SGE (0.7682 vs. 0.1124 mAcm-2) with a negative shift of potential by 49 mV.

271

4-/3- Figure 7.10: Electrocatalytic activity of MPGE in [Fe(CN)6] redox system. (a) CV curves 3-/4- of SGE and MPGE and (b) DPV responses of SGE and MPGE in 2 mM [Fe(CN)6] solution. 3-/4- (c) CV curves at different concentrations of [Fe(CN)6] on MPGE and (d) bar-diagram of the corresponding currents (cathodic). (e) CV curves at different scan rates from 5 mVs-1 to 1000 -1 3-/4- mVs on MPGE in 2 mM [Fe(CN)6] solution and (f) summary of the corresponding current responses upon the square-root of scan rates. (g) Amperometric responses on MPGE with the 3-/4- successive addition of Fe[(CN)6] solution into the 0.01 M PBS (pH-7) and (h) the corresponding calibration plot; inset- Lineweaver-Burk Model.

272

We have performed the CV and DPV measurement of MPGEs with different thickness. In both cases, the current reached a plateau after 1000 s of electrodeposition and experienced a polynomial relationship (Figures 7.11 and 7.12). The ECSA follows a linear relationship with the film thickness (i.e., deposition time); however, the current density (DPV response in 3-/4- [Fe(CN)6] redox system) follows a polynomial relationship. Moreover, the ECSA also gives rise a polynomial pattern with the current density, which expressing the dependency of current density with the thickness (Figure 7.13). Based on these results, we select 1000 s as the optimal deposition for further investigation.

3-/4- Figure 7.11: (a) The CV’s in 2 mM [Fe(CN)6] for MPGEs prepared at different deposition time (thickness); (b) the corresponding cathodic current, which follows a polynomial relationship with the increasing film thickness.

Figure 7.12: (a) The DPV responses of MPGE with increasing film thickness; (b) the corresponding current density. The current follows the polynomial trend with the increasing deposition time.

273

Figure 7.13: (a) The representation of both the current density (black line) and ECSA’s (red line) obtained from different electrodeposited MPGE, where the film thickness (deposition) versus ECSA follows the linear relationship and the film thickness (deposition) versus current density follows polynomial relationship (a characteristic of enzymatic electrocatalysis).

To further examine the performance of MPGE, we conducted the CV measurement at 3-/4- different concentration (0.5 mM to 15 mM) of the [Fe(CN)6] system (Figure 7.10c). With the increasing strength of the redox system, the observed currents increase gradually, revealing that the porous structure provides an enormous surface to accelerate the redox reactions (Figure 7.10d). To elucidate the electrochemical mechanism involving in the MPGE towards 3-/4- [Fe(CN)6] system, we performed CV measurements as a function of different scan rate (ν)

(Figure 7.10e). Both the ipc and ipa are increasing proportionally with the increased scan rates -1 1/2 values (5-1000 mVs ) and show a linear relationship with ν for both ipa and ipc, which indicates the kinetics towards electrode surface are mainly diffusion-controlled (Figure 7.10f).35 Furthermore, chronoamperometric (CA) responses of MPGE were recorded upon the 3-/4- successive addition of [Fe(CN)6] solution (Figure 7.9g), where the CA response increases steeply. The calibration curve follows the characteristic Michaelis-Menten equation (Figure app 7.10h). The apparent Michaelis-Menten constant (Km ) can be obtained from the electrochemical version of Lineweaver-Burk model, and it is estimated to be 1.89 mM. This 3-/4 value is significantly low, suggesting the higher affinity of MPGE to [Fe(CN)6] redox system, further verifying electrochemical activity of MPGE.36 This superior activity is related to its s strong electronic field around the centre of the uniform mesopores and on the walls between the pores. In addition to this excellent activity, the MPGE is advantageous with high stability. The MPGE provides stable signal generation over the multiple cycles (30 cycles) of

274

3-/4 CV measurement using [Fe(CN)6] redox system (Figure 7.14). This stability in reaction media (PBS) and strong affinity towards the redox molecule are highly strategic for exploiting MPGE as the impeccable substrate (signal-transduction) for electrochemical biosensing.

3-/4- Figure 7.14: Stability of MPGE over several CV curves in 2 mM [Fe(CN)6] solution

To explore the functionality mesoporous Au structure for a proof-of-concept (POC) biosensing, we have utilised as prepared MPGE for miRNA detection. The assay principle for the MPGE integrated amplification-free ultra-sensitive miRNA detection is outlined in Figure 7.1b. The assay is mainly comprised of two-steps; (i) magnetic separation and purification of target miRNA sequence using target-complementary capture probe attached-with a dynabead,37 (ii) direct adsorption of purified miRNA on MPGE via RNA-Au affinity interaction through conventional physisorption and chemisorption mechanism followed by DPV interrogation.38 Herein, miR-9-2 is chosen as a target because of its potentiality as predictive and diagnostic biomarkers for carcinoma.39-40 For instance, a low level of miR-9 is significantly correlated with worse lymphatic invasion and advanced TNM stage in metastatic nasopharyngeal carcinoma.41 The magnetic isolation significantly reduces matrix effect from complex bio-system, remove debris and non-specific sequences, thereby provides the pure target miRNA for electrochemical readout. Moreover, bringing target miRNA (only) onto MPGE significantly biofouling issues generally experienced in the electrochemical sensor. The purified miRNA was then directly adsorbed on to the MPGE. The underlying principle of the direct adsorption of miRNA sequences on a clean MPGE can be explained by the well-explored nucleobases’ adsorption affinity towards the bare Au surface where miRNA bases are directly adsorbed in a sequence-dependent manner.19 The adsorbed miRNA were then detected by DPV

275

3-/4- interrogation in the presence of an electroactive [Fe(CN)6] system. It has been shown earlier 4-/3- that the [Fe(CN)6] redox system alone can be used for quantification of surface-bound nucleic acid attached at unmodified Au surface.42-43 Under the electron-transfer kinetic-based 3-/4- mechanism the coulombic repulsion between [Fe(CN)6] and negatively charged nucleic acid 3-/4- strands (low coverage) at the electrode surface generate reduced current from [Fe(CN)6] molecules. The amount of current reduction in relatives to bare is directly related to the miRNA present at the electrode surface. To select the optimum electrode thickness, initially, we have challenged the same amount of purified miRNA on to the MPGE with different thickness (prepared by different deposition; 0 s to 2000 s). It has been shown that the current responses have been increased with increasing electrode thickness, however, the polynomial fittings express that the current started to became a platue with the MPGE deposited for 1000s (Figure 7.15). Therefore we have chosen the MPGE that prepared at 1000 s electrodeposition time for the subsequent experiment for proof-of-concept assay for microRNA detection.

Figure 7.15: DPV responses after the adsorption of the same quantity of miR-9-2 (10 pM) at MPGE with different deposition time (0 - 2000 s).

To examine the superiority of MPGE over SGE for miRNA detection, we have absorbed the same amount of (10 pM) of magnetically purified miRNA sequence at both surfaces. Figure 7.16 (a and b) represent the DPV responses before and after miRNA adsorption at both Au surface. The current response for miR-9-2 at MPGE was significantly higher (0.382 vs. 0.61 mAcm-2) than SGE (Figure 7.16c). The high current from MPGE can be related to; (i) mesopores induced the large exposed surface area of MPGE, which adsorbs more miRNA 3-/4- through the RNA–Au affinity interaction, thereby more substantial amount of [Fe(CN)6]

276

redox can be diffused from the negatively charged miRNA-confined surface and (ii) 3-/4- electrocatalytic signal enhancement of [Fe(CN)6] redox system by MPGE. To visualize the adsorption of miRNA sequence at MPGE surface, we have recorded the topographic image after miRNA adsorption using SEM and atomic force microscopy (AFM) (Figure 7.17 a and b). As can be seen in Figure 7.17b, the morphology (topography) of the miRNA-MPGE electrode surface is entirely different from the bare MPGE and the presence of miRNA at MPGE detected by AFM. To evaluate the assay specificity, we have challenged the MPGE surface with no-template control (NoT) and wrong-target (non-complementary sequence; miR- 338-3p) (Figure 7.16d). It has been seen that the NoT and the non-complementary sequence generates negligible current responses (% Jr = 1.79 and 2.56 respectively), revealing the high specificity of dynabead based isolation and purification. A very tiny increment of current for miR-338-3p, probably due to the binding of very few amounts of miR-338-3p sequence with the capture probe, however, this value is much lower than the target one (% Jr = 51.2). The current response for 10 pM of target miRNA is almost 28-times and 20-times higher than both NoT and wrong target, which is due to the sequence-specific capture bring an adequate amount of target miRNA at MPGE surface, which upon DPV interrogation in the presence of redox system generate a significant reduction of current in comparison to bare.

277

Figure 7.16: (a, b,) DPV responses before and after 10 pM of miRNA adsorbed on SGE and MPGE, respectively. (c) Sensitivity Comparison; both SGE and MPGE generates a significantly higher amount of current in response to NoT (PBS is used instead of miRNA), and surprisingly MPGE generate 8-times higher current than that of SGE (0.384 vs. 0.061 mAcm-2) (d) Assay specificity where MPGE with the target miRNA generate pointedly higher responses from Bare (MPGE; without miRNA), NoT (PBS), wrong target (miR-338-3p), respectively.

Recent studies on disease diagnosis show that the miRNA levels either upregulated or downregulated in disease state (cells, plasma or exosomes) compare to the healthy source. miRNA expression also varies according to the type and stage of the disease, especially cancer. Importantly, the changes in miRNA expression are meagre compared to that of a healthy state, clearly indicating the requirement of a highly sensitive detection system. To determine the detection sensitivity using MPGE, a series of synthetic miR-9-2 with different concentration ranging from 1 nM to 100 aM were analysed. It has been observed that the charge generated by DPV interrogation was increased with increasing level of miRNA (Figure 7.17 c and d). This was attributed to the higher amount of miRNA was isolated and thus adsorbed on to the 3-/4- MPGE surface produces increased coulombic repulsion of [Fe(CN)6] system to the

278

negatively charged miRNA resulting in the reduced Faradic current and increased relative current response changes regarding the control (NoT) and bare MPGE. The linear regression equation was estimated, and the limit of detection was calculated by considering s/n ration equals to three compared to NoT. The MPGE enables a limit of detection of 100 aM with good reproducibility (relative standard deviation, RSD of < 6%, for n = 3) for detecting magnetically purified miRNA. This dynamic series of detection with attomolar sensitivity indicates the potentiality of MPGE for identifying and analysing miR-9-2 in purified miRNA from complex biological matrixes with varying level of miRNA even at an earlier stage.

Figure 7.17 (a) SEM and (b) AFM topography image miRNA-adsorbed MPGE, respectively. (c) Assay sensitivity; the DPV responses with the different concentration of synthetic target miR-9-2 ranging from 1 fM to 1 nM. (d) The corresponding bar diagram of assay sensitivity. Error bars represent the standard deviation of the measurements (% RSD = < 5%, for n = 3). Inset, corresponding calibration plot.

The detection limit of the assay is highly comparable with the recently reported electrochemical strategies for mRNA detection. For instance, our assay has realized around 9 times LOD than recently reported readout by Li et. al. though they have utilized base- mismatched catalytic hairpin assembly (CHA) amplification whereas this assay offers much simpler platform without any amplification steps.44 This assay also shows increased sensitivity

279

compared to recent time reported nanostructure-based strategies such as SPGE,34 silver nanofoam (AgNF),45 amino-graphene-modified glassy carbon electrode (GCE),464 Au- 22, 31 47 48 49 NPFe2O3/SPCE, DNA-modified Au electrode, GO-NPFe2O3/SPCE and Au-μPADs. It is also imperative to remark that there are some nanotechnology-based methods have also been reported with similar detection sensitivity or even lower range of miRNA (Details comparison are shown in Table7.2).44-51 However, these methods utilized cumbersome enzymatic amplification, multi-step cascade electrocatalysis, isothermal amplification, enzymatic displacement reaction and different redox probe. On the contrary, our methods offer straightforward, secure, non-enzymatic, label-free detection and most significantly it requires an inexpensive MPGE. We deem that the LoD of 100 aM of our assay is passable to retract the level of readily available miRNA biomarker from the clinically-pertinent concentration of target samples. We also have confidence in that several distinctive features of our assay have ascribed to this ultra-sensitivity. These are as follows;- (i) the tunable mesoporous, concave surface inside the pores, profusion in high-index facets together with the assembly of kink/steps sites preferentially facilitates the adsorption of nucleic acid as well as significant enhancement of current density towards the redox process, (ii) complementary capture probe and dynabeads- based magnetic collection and purification of target miRNA expressively cuts the matrix effect of the compound biological testers and eradicates the nonspecific targets. Furthermore, the capturing and purification steps of the target miRNA can be spatially separated from the electrode to ease the biofouling issues, (iii) the inherently sensitive DPV readout (its superior capacitive or background current elimination ability curtails the influence of charging current i.e., only Faradic current is counted) resulting in more accurate detection of the target miRNA, and (iv) PCR or any enzymatic amplification-free processes avoid amplification bias (i.e., less false-positive response). We foretell that the MPGE-based assay could also be radially able to detect any miRNA marker detection and single-step electrochemical readout can easily be translated to an automative device for clinical diagnostics.

280

Table 7.2: Comparison of nanostructure-based approaches of electrochemical detection of miRNA

MicroRNA Electrode Materials Principle Amplification strategy Dynamic range LOD Ref.

miR-21 Gold EC Base mismatched catalytic hairpin assembly 5 fM–0.1 nM 1.1 fM 44 ratiometry amplification

miR-155 amino-graphene DPV highly conductive substrate for hybridization 50 pM – 1 nM 12.5 fM 46 assembled GCE miR-21 acpcPNA/ CV AgNF redox reaction 0.2 fM – 1nM 0.2 fM 45 PPy/AgNF-Au miR-141 DNA-modified Au DPV T7 exonuclease (exo) (DNA cleavage reaction) 100 aM – 10 pM 45 aM 47 and CuNPs miR-155 Au SWV Cascade strand displacement polymerization and 10 aM – 5 pM 3.2 aM 50 polymerization–nicking isothermal synergetic reactions

48 miR-21 Au-μPADs DPV CeO2-Au@GOx amplification probe 1 fM – 1 pM 0.434 fM microRNA- Au SWV Cyclic enzymatic signal amplification (CESA) 50 aM - 50 pM 15 aM 51 196a and template-free DNA extension reaction

22 miR-21 Au-NPFe2O3NC- CC Electrocatalytic signal amplification towards 1 fM – 10 nM 1 fM modified SPCE redox marker RuHex 31 miR-107 Au-NPFe2O3NC- CC Electrocatalytic signal amplification and redox 100 aM – 1 nM 100 aM modified SPCE coupling miR-107 SPGE DPV Poly(A) Extensions of miRNAs 1 pM – 1 nM 10 fM 30 3+ 3- 48 miR-21 GO-NPFe2O3/SPCE CC [Ru(NH3)6] /[Fe(CN)6] redox system 1 fM – 1nM 1fM

miR-9-2 MPGE DPV Amplification-free 100 aM – 1 nM 100 aM This study

281

7.4. Conclusion

In summary, we have developed a novel electrode platform, which allows highly sensitive (LOD: 100 aM) detection of miRNA without any amplification or enzymatic process. Compared with the up-to-date miRNA assays with different cascade signal amplification, the synthesis of MPGE and design of this work is simple but subtle. The MPGE is highly stable and can be reused for multiple tests. The hollow pore-mediated signal amplification provides the superior adsorption sites for miRNA, catalytic effect and thereby enhances the sensitivity. Besides, the preparation of MPGE is very cost-effective with excellent repeatability and reusability. This assay offers a single-step detection after isolation of target miRNA from the sources, required less than 1 hr, providing the translation capability for designing portable miRNA sensor for clinics. We envisage that the developed method indeed provides a versatile platform for miRNA detection and may find a broad spectrum of applications in biological studies, bioanalysis and clinical diagnosis.

7.5. References

1. Rosenfeld, N.; Aharonov, R.; Meiri, E.; Rosenwald, S.; Spector, Y.; Zepeniuk, M.; Benjamin, H.; Shabes, N.; Tabak, S.; Levy, A., MicroRNAs accurately identify cancer tissue origin. Nature Biotechnology 2008, 26 (4), 462. 2. Bracken, C.P., Scott, H.S. and Goodall, G.J., A network-biology perspective of microRNA function and dysfunction in cancer. Nature Reviews Genetics, 2016, 17 (12), 719. 3. Islam, M. N.; Masud, M. K.; Haque, M. H.; Hossain, M. S. A.; Yamauchi, Y.; Nguyen, N. T.; Shiddiky, M. J., RNA biomarkers: diagnostic and prognostic potentials and recent developments of electrochemical biosensors. Small Methods 2017, 1 (7), 1700131. 4. Chin, L. J.; Slack, F. J., A truth serum for cancer—microRNAs have major potential as cancer Biomarkers. Cell Research, 2008, 18 (10), 983. 5. Jeffrey, S. S., Cancer biomarker profiling with microRNAs. Nature Biotechnology 2008, 26 (4), 400. 6. Git, A.; Dvinge, H.; Salmon-Divon, M.; Osborne, M.; Kutter, C.; Hadfield, J.; Bertone, P.; Caldas, C., Systematic comparison of microarray profiling, real-time PCR, and

282

next-generation sequencing technologies for measuring differential microRNA expression. RNA 2010, 16 (5), 991-1006. 7. Pritchard, C. C.; Cheng, H. H.; Tewari, M., MicroRNA profiling: approaches and considerations. Nature Reviews Genetics 2012, 13 (5), 358. 8. Masud, M.K.; Umer, M.; Hossain, M.S.A.; Yamauchi, Y.; Nguyen, N.T.; and Shiddiky, M.J., Nanoarchitecture frameworks for electrochemical miRNA detection. Trends in biochemical sciences, 2019, 44, 5, 433-452. 9. Labib, M.; Sargent, E. H.; Kelley, S. O., Electrochemical methods for the analysis of clinically relevant biomolecules. Chemical Reviews 2016, 116 (16), 9001-9090. 10. Soleymani, L.; Fang, Z.; Sargent, E. H.; Kelley, S. O., Programming the detection limits of biosensors through controlled nanostructuring. Nature Nanotechnology 2009, 4 (12), 844. 11. Innocenzi, P.; Malfatti, L., Mesoporous thin films: properties and applications. Chemical Society Reviews, 2013, 42 (9), 4198-4216. 12. Walcarius, A., Mesoporous materials and electrochemistry. Chemical Society Reviews, 2013, 42 (9), 4098-4140. 13. Ferrari, M., Cancer nanotechnology: opportunities and challenges. Nature Reviews Cancer 2005, 5 (3), 161. 14. Lei, J.; Ju, H., Signal amplification using functional nanomaterials for biosensing. Chemical Society Reviews 2012, 41 (6), 2122-2134. 15. Azhar, A.; Li, Y.; Cai, Z.; Zakaria, M. B.; Masud, M. K.; Hossain, M. S. A.; Kim, J.; Zhang, W.; Na, J.; Yamauchi, Y., Nanoarchitectonics: A New Materials Horizon for Prussian Blue and Its Analogues. Bulletin of the Chemical Society of Japan 2019, 92 (4), 875-904. 16. Yadav, S.; Masud, M. K.; Islam, M. N.; Gopalan, V.; Lam, A. K.; Tanaka, S.; Nguyen, N. T.; Hossain, M. S. A.; Li, C.; Yamauchi, M. Y.; Shiddiky, M. J. A., Gold-loaded nanoporous iron oxide nanocubes: a novel dispersible capture agent for tumor- associated autoantibody analysis in serum. Nanoscale 2017, 9 (25), 8805-8814. 17. Vafaiee, M.; Vossoughi, M.; Mohammadpour, R.; Sasanpour, P., Gold-plated electrode with High scratch strength for electrophysiological Recordings. Sci Rep 2019, 9 (1), 2985. 18. Wang, S.; Liu, K.; Yao, X.; Jiang, L., Bioinspired surfaces with superwettability: new insight on theory, design, and applications. Chemical Reviews 2015, 115 (16), 8230- 8293. 283

19. Koo, K. M.; Sina, A. A.; Carrascosa, L. G.; Shiddiky, M. J.; Trau, M., DNA–bare gold affinity interactions: mechanism and applications in biosensing. Analytical Methods 2015, 7 (17), 7042-7054. 20. Demers, L. M.; Östblom, M.; Zhang, H.; Jang, N.-H.; Liedberg, B.; Mirkin, C. A., Thermal desorption behavior and binding properties of DNA bases and nucleosides on gold. Journal of the American Chemical Society 2002, 124 (38), 11248-11249. 21. Li, H.; Rothberg, L., Colorimetric detection of DNA sequences based on electrostatic interactions with unmodified gold nanoparticles. Proceedings of the National Academy of Sciences 2004, 101 (39), 14036-14039. 22. Masud, M. K.; Islam, M. N.; Haque, M. H.; Tanaka, S.; Gopalan, V.; Alici, G.; Nguyen, N.-T.; Lam, A. K.; Hossain, M. S. A.; Yamauchi, Y., Gold-loaded nanoporous superparamagnetic nanocubes for catalytic signal amplification in detecting miRNA. Chemical communications 2017, 53 (58), 8231-8234. 23. Mai, Y.; Eisenberg, A., Self-assembly of block copolymers. Chemical Society Reviews 2012, 41 (18), 5969-5985. 24. Cheng, Y.-J.; Gutmann, J. S., Morphology phase diagram of ultrathin anatase TiO2 films templated by a single PS-b-PEO block copolymer. Journal of the American Chemical Society 2006, 128 (14), 4658-4674. 25. Sokolova A, Whitten AE, de Campo L, Christoforidis J, Eltobaji A, Barnes J, Darmann F, Berry A. Performance and characteristics of the BILBY time-of-flight small-angle neutron scattering instrument. Journal of Applied Crystallography 2019, 52, 1-12. 26. Arnold O, Bilheux JC, Borreguero JM, Buts A, Campbell SI, Chapon L, Doucet M, Draper N, Leal RF, Gigg MA, Lynch VE. Mantid—Data analysis and visualization package for neutron scattering and μ SR experiments. Nuclear Instruments and Methods in Physics Research Section A: Accelerators, Spectrometers, Detectors and Associated Equipment 2014, 764, 156-66. 27. Sukeri, A.; Saravia, L.P.H.; Bertotti, M., A facile electrochemical approach to fabricate a nanoporous gold film electrode and its electrocatalytic activity towards dissolved oxygen reduction. Physical Chemistry Chemical Physics, 2015, 17 (43), 28510-28514. 28. Trasatti, S.; Petrii, O.A., Real surface area measurements in electrochemistry. Pure and Applied Chemistry, 1991, 63 (5), 711-734. 29. Kline SR. Reduction and analysis of SANS and USANS data using IGOR Pro. Journal of Applied Crystallography 2006, 39 (6), 895-900.

284

30. Koo, K.M., Carrascosa, L.G., Shiddiky, M.J. and Trau, M., Amplification-free detection of gene fusions in prostate cancer urinary samples using mrna–gold affinity interactions. Analytical Chemistry, 2016, 88 (13), 6781-6788. 31. Islam, M. N.; Masud, M. K.; Nguyen, N.-T.; Gopalan, V.; Alamri, H. R.; Alothman, Z. A.; Al Hossain, M. S.; Yamauchi, Y.; Lamd, A. K.; Shiddiky, M. J., Gold-loaded nanoporous ferric oxide nanocubes for electrocatalytic detection of microRNA at attomolar level. Biosensors and Bioelectronics 2018, 101, 275-281. 32. Li, C., Dag, Ö., Dao, T.D., Nagao, T., Sakamoto, Y., Kimura, T., Terasaki, O. and Yamauchi, Y., Electrochemical synthesis of mesoporous gold films toward mesospace- stimulated optical properties. Nature Communications, 2015, 6, 6608. 33. Kelley, S.O., Boon, E.M., Barton, J.K., Jackson, N.M. and Hill, M.G., Single-base mismatch detection based on charge transduction through DNA. Nucleic Acids Research, 1999, 27 (24), 4830-4837. 34. Li, C.; Jiang, B.; Chen, H.; Imura, M.; Sang, L.; Malgras, V.; Bando, Y.; Ahamad, T.; Alshehri, S. M.; Tominaka, S., Superior electrocatalytic activity of mesoporous Au film templated from diblock copolymer micelles. Nano Research 2016, 9 (6), 1752-1762. 35. Bard, A. J.; Faulkner, L. R.; Leddy, J.; Zoski, C. G., Electrochemical methods: fundamentals and applications. Wiley New York 1980, Vol. 2. 36. Dutta, A. K.; Maji, S. K.; Srivastava, D. N.; Mondal, A.; Biswas, P.; Paul, P.; Adhikary, B., Peroxidase-like activity and amperometric sensing of hydrogen peroxide by Fe2O3 and Prussian Blue-modified Fe2O3 nanoparticles. Journal of Molecular Catalysis A: Chemical 2012, 360, 71-77. 37. Islam, M. N.; Gopalan, V.; Haque, M. H.; Masud, M. K.; Al Hossain, M. S.; Yamauchi, Y.; Nguyen, N.-T.; Lam, A. K.-Y.; Shiddiky, M. J., A PCR-free electrochemical method for messenger RNA detection in cancer tissue samples. Biosensors and Bioelectronics 2017, 98, 227-233. 38. Koo, K. M.; Carrascosa, L. G.; Shiddiky, M. J.; Trau, M., Poly (A) extensions of miRNAs for amplification-free electrochemical detection on screen-printed gold electrodes. Analytical Chemistry 2016, 88 (4), 2000-2005. 39. Hildebrandt, M. A.; Gu, J.; Lin, J.; Ye, Y.; Tan, W.; Tamboli, P.; Wood, C. G.; Wu, X., Hsa-miR-9 methylation status is associated with cancer development and metastatic recurrence in patients with clear cell renal cell carcinoma. Oncogene 2010, 29 (42), 5724-5728.

285

40. Zhou, X.; Marian, C.; Makambi, K. H.; Kosti, O.; Kallakury, B. V.; Loffredo, C. A.; Zheng, Y.-L., MicroRNA-9 as potential biomarker for breast cancer local recurrence and tumor estrogen receptor status. PloS One 2012, 7 (6), e39011. 41. Lu, J.; Xu, X.; Liu, X.; Peng, Y.; Zhang, B.; Wang, L.; Luo, H.; Peng, X.; Li, G.; Tian, W., Predictive value of miR-9 as a potential biomarker for nasopharyngeal carcinoma metastasis. British Journal of Cancer 2014, 110 (2), 392. 42. Zhang, J.; Wang, L.; Pan, D.; Song, S.; Fan, C., DNA hybridization “turns on” electrocatalysis at gold electrodes. Chemical Communications 2007, (11), 1154-1156. 43. Sina, A. A. I.; Howell, S.; Carrascosa, L. G.; Rauf, S.; Shiddiky, M. J.; Trau, M., eMethylsorb: electrochemical quantification of DNA methylation at CpG resolution using DNA–gold affinity interactions. Chemical Communications 2014, 50 (86), 13153-13156. 44. Li, X.; Dou, B.; Yuan, R.; Xiang, Y., Mismatched catalytic hairpin assembly and ratiometric strategy for highly sensitive electrochemical detection of microRNA from tumor cells. Sensors and Actuators B: Chemical 2019, 286, 191-197. 45. Kangkamano, T.; Numnuam, A.; Limbut, W.; Kanatharana, P.; Vilaivan, T.; Thavarungkul, P., Pyrrolidinyl PNA polypyrrole/silver nanofoam electrode as a novel label-free electrochemical miRNA-21 biosensor. Biosensors and Bioelectronics 2018, 102, 217-225. 46. Salimi, A.; Kavosi, B.; Navaee, A., Amine-functionalized graphene as an effective electrochemical platform toward easily miRNA hybridization detection. Measurement 2019, 143, 191-198. 47. Miao, P.; Zhang, T.; Xu, J.; Tang, Y., Electrochemical detection of miRNA combining T7 exonuclease-assisted cascade signal amplification and DNA-templated copper nanoparticles. Analytical Chemistry 2018, 90 (18), 11154-11160. 48. Islam, M. N.; Gorgannezhad, L.; Masud, M. K.; Tanaka, S.; Hossain, M. S. A.; Yamauchi, Y.; Nguyen, N. T.; Shiddiky, M. J., Graphene‐Oxide‐Loaded Superparamagnetic Iron Oxide Nanoparticles for Ultrasensitive Electrocatalytic Detection of MicroRNA. ChemElectroChem 2018, 5 (17), 2488-2495. 49. Sun, X.; Wang, H.; Jian, Y.; Lan, F.; Zhang, L.; Liu, H.; Ge, S.; Yu, J., Ultrasensitive microfluidic paper-based electrochemical/visual biosensor based on spherical-like cerium dioxide catalyst for miR-21 detection. Biosensors and Bioelectronics 2018, 105, 218-225.

286

50. Miao, P.; Jiang, Y.; Zhang, T.; Huang, Y.; Tang, Y., Electrochemical sensing of attomolar miRNA combining cascade strand displacement polymerization and reductant-mediated amplification. Chemical Communications 2018, 54 (53), 7366- 7369. 51. Guo, J.; Yuan, C.; Yan, Q.; Duan, Q.; Li, X.; Yi, G., An electrochemical biosensor for microRNA-196a detection based on cyclic enzymatic signal amplification and template-free DNA extension reaction with the adsorption of methylene blue. Biosensors and Bioelectronics 2018, 105, 103-108.

287

Chapter 8

Conclusions and future recommendations

288

8. Conclusions and Future Recommendations

8.1. Conclusions

The overall emphasis of this PhD study was to develop engineered nanostructured-based platform technology for the analysis of clinically relevant biomarkers. This study advances the integration of intrinsic properties (superparamagnetism, nanozyme activity and electrocatalytic activity) of nanostructure into the biosensor to achieve relatively simple and ultra-sensitive detection tools for clinics. This thesis explores the synthesis of ordered mesoporous nanostructure and exploits the phase-dependent activity for biosensing. Upon the selection of favourable mesoporous nanostructure, a new class of gold-loaded iron oxide nanocube was reported, which provides the good biocompatibility, enabled direct functionalization with different biomolecules (antibody, nucleic acid) thereby avoids the cumbersome biofunctionalization steps. Moreover, the nanocube shows significant nanozyme activity even at room temperature, which facilitates the room-temperature sensing of a cancer biomarker; autoantibody. In addition, the nanocube possesses intrinsic electrocatalytic activity towards the popularly used redox molecules and enabled non-enzymatic amplification-steps free electrochemical sensing of microRNA. However, these electrochemical sensors required modification of electrode with the porous nanostructure. To avoid this modification step, this thesis presented the fabrication of mesoporous gold electrode-based microRNA sensor. The microRNA sensor provides single-step electrochemical detection with an attomolar level of sensitivity without any external electrode or modification steps. Remarkably, all the detection strategies described here demonstrate great promise headed for the development of portable point-of-care diagnostics devices for clinics.

Briefly, the thesis describes the synthesis of mesoporous nanostructures and the design and development of biosensing device consist of four novel readout system for the rapid, inexpensive, straight-forward analysis of disease-specific biomarkers, i.e., autoantibody and microRNA. Considering the prevailing advantages (e.g., small size, high surface area, flexible structure, low toxicity, biocompatibility, magnetism, nanozyme and electrocatalytic activity) of nanostructured (porous) materials, we have synthesized and investigate the influence of crystal phases (α-Fe2O3 and γ-Fe2O3) of mesoporous iron oxide (IO) towards the peroxidase mimetic activity and superparamagnetism. Moreover, we have also demonstrated the applicability of these mesoporous IOs as nanozymes for detecting an easily accessible glucose

289

biomarker. Mesoporous γ-Fe2O3 shows high nanozyme activities (and magnetism) toward the catalytic oxidation of chromogenic substances, such as TMB, ABTS related to that of α-Fe2O3.

The γ-Fe2O3 enabled the colourimetric detection of glucose with a limit of detection (LOD) of 0.9 µM. However, the naked iron oxide possesses higher surface energy and thereby tend to aggregate, limiting their dispersion in solution and body fluids. In order avoid that, we have then reported on the synthesis of gold-loaded mesoporous iron oxide nanocube

(Au−NPFe2O3NC), which shows promising biocompatibility, well dispersibility in aqueous media, superparamagnetism and intrinsic peroxidase mimetic activity. Taking into account of these intrinsic properties, we have developed a molecular sensor with enhanced electrocatalytic and colourimetric (naked eye) detection of p53-autoantibodies (mutation in TP53 proteins is an early indicator of cancer and found to be present in almost 80% of all carcinogenesis). This detection platform is capable of giving a yes/no answer for the presence of target autoantibody, and the naked-eye screening can be exploited as a first-pass test for a large samples screening, followed by more precise quantification using electrochemical readout.

MicroRNAs are considered as a promising diagnostic and prognostic biomarkers for many diseases including cancer. Considering the superior superparamagnetism and electrocatalytic activity of the nanocube, a non-enzymatic, amplification-free electrochemical detection platform was developed for microRNA. The assay showed an excellent detection sensitivity down to 100 fM and specificity towards the analysis of miR-21 in cell lines and tissue samples derived from patients with oesophageal squamous cell carcinoma (ESCC). This detection platform simplifies the assay design by avoiding the complex chemistries underlying each step of the sensor fabrication by utilizing the superparamagnetic properties of nanocube (an external magnet is used to attach the nanocube on top of the electrode surface and miRNAs are allowed to adsorb on electrode surface directly). Nevertheless, one of the major sources of inefficiencies in miRNA biosensing is its low abundance in biological samples. To achieve ultra-sensitivity, we have integrated the 3-/4- MB/Fe(CN)6] redox cycle with the intrinsic electrocatalytic activity of the Au−NPFe2O3NC nanocube. The three-mode amplification system (e.g., electrocatalytic nanocube, MB and ferricyanide redox couples) facilitate an electrochemical detection platform for miRNA present in neurodegenerative disease (e.g. ALS) samples with a LOD of 100 aM.

The purpose of final sensor platform was to reflect on the extent to which the microRNA detection can move towards more personalised diagnostics of disease and enable a

290

user-friendly and portable approach that is suitable for point-of-care settings even in resource- limited settings. In this regard, we have fabricated a mesoporous gold electrode using self- assembled polymeric micelles as a pore directing agent. The mesoporous gold electrode fascinates a greater extent of nucleic acid adsorption owing to abundantly open pores and its good affinity to the Au surface, and single-step electrochemical detection. This method attains a wide dynamic linear range from 100 aM to 1 nM with an ultra-low limit detection of 100 aM. This method simplifies the assay building by circumventing multiple measures tangled in conventional biosensing approaches through recognition and transduction layers. All of the detection platforms developed in this PhD were demonstrated for their applicability in complex biological samples (a cohort of cancer and MND cell lines and patient samples), to ensure their translational potential. Importantly, the analytical performance of all microRNA assays was validated with the standard RT-qPCR approach.

8.2. Future recommendations

This PhD thesis has significantly contributed to the synthesis of mesoporous nanostructures, signal transduction system and their integration to colourimetric and electrochemical biosensor to achieve rapid, specific, ultra-sensitive detection of disease-specific biomarkers in a portable point-of-care approach. Consequently, these studies which were reported within this thesis will serve as a basis for future research. From the synthetic viewpoint, many issues need to be considered and addressed to make (superpara-) magnetic nanostructure suitable for a point-of-care platform, such as; (i) the stability of SMNPs in the aqueous system, (ii) biocompatibility, (iii) biofunctionalization of magnetic nanostructure as the half-life of biomolecule-nanostructure complexes is still low; (iv) magnetic susceptibility as the functional molecules cover the core magnetic molecules and (v) the combination of two or more nanostructures are still required to achieve functional magnetic nanostructures. Single magnetic nanostructure with in situ probe or capture biomolecule (antibody) functionalization can be considered for future magnetic nanostructure design for diagnostics device development. Considering the analytical aspect, microRNAs are unstable at room temperature and more susceptible to RNase degradation. RNA samples. Thus, extensive care required during storage, extraction and purification. False-positive response from the cross-talk amongst targets and closely related non-target microRNA sequences should be considered during the microRNA assay design. We have minimised this by utilizing magnetic-bead based purification and using innovative transduction surface, nonetheless, the

291

opportunity for circumventing the false-positive response considering other state-of-the-art approaches is still “wide open”. We have exploited the colourimetry and electrochemistry as a signal read-out, however, another promising direction could be using sophisticated surface- enhanced Raman scattering readout. This readout could read distinctive properties or surface of nanostructures (Raman enhancement) which aid in generating distinctive signals for target molecules. Moreover, to translate the proof-of-concept bioassays to clinics, each of the assays should be enabled to detect multiple biomarkers simultaneously. This is because, most of the cases, for instance, cancer is a multifaceted complex disease and there is no particular biomarker that can deliver comprehensive information about the exact disease. Therefore, a combined analysis of a group of novel biomarkers is suggested. With the recent tremendous advances in the fabrication technologies for microfluidic systems and microelectromechanical, a fully automated device that can role with minimal human involvement will become a reality in the future. In addition, extensive clinical evaluation (with a large cohort of patient samples, blinded studies, cross-institutional and cross-technological validation) of developed detection platform is essential to ensure adequate performance for clinical use.

292