PhD

3.º CICLO FCUP FCT/UNL 2014

synthesisand characterization encapsulating organic and inorganic fluorophores: Development of fluorescent silica nanoparticles

Development of

fluorescent silica

nanoparticles encapsulating organic and

inorganic D fluorophores: D Cristina Sofia Cristina Neves Santos dos synthesis and characterization

Cristina Sofia dos Santos Neves Tese de Doutoramento apresentada à

Faculdade de Ciências da Universidade do Porto, Faculdade

de Ciências e Tecnologia da Universidade Nova de Lisboa Química Sustentável 2014

Developmentof Eulália Pereira,Eulália Professora FaculdadeCiênciasdeAuxiliar, Coorientador Peter Eaton, FaculdadeInvestigadorCiênciasdeAuxiliar, Orientador 2014 Departamentodee Química Bioquímica Doutoramento em Química Sustentável CristinaSantos Sofia dos Neves characterizationand fluorophores:synthesis and inorganic encapsulatingorganic nanoparticles fluorescentsilica

D

“As ordens que levava não cumpri E assim contando tudo o que vi Não sei se tudo errei ou descobri”

Shophia de Mello Breyner (excerto do poema Deriva - VIII,1982)

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Acknowledgments

“No man is an island, Entire of itself, Every man is a piece of the continent, A part of the main.” (John Donne)

During the development of this work many challenges have emerged, and with them several people who helped me in one way or another, to overcome them. This thesis wouldn’t be the same without their support and for that I would like to take a few lines to thank those who believed in me and in this work.

First of all I would like to thank my supervisors Dr. Peter Eaton and Dr. Eulália Pereira for the opportunity to perform this work. Thank you for accepting me in the lab and for providing all means for the development of this project. Also for the guidance and words of encouragement that were needed in some curves of this way. Thank you also for the opportunities you gave me.

Part of the work present in this thesis wouldn’t be possible without the help of Dr. Salete Balula. Her guidance, support and friendship were of great importance to achieve my goals. Thanks for introducing me to the amazing world of polyoxometalates and giving me the chance to learn and grow up in this field.

To Dr. Carlos Granadeiro and Dr. Luis Cunha Silva a special thanks for the help with the synthesis and characterization of some nanoparticles in particular the help with X-ray crystallography and FT-RAMAN spectroscopy.

To Dr. Sandra Gago from the Chemistry Department and to Dr. Gabriel Feio from the Department of Materials Science (CENIMAT), Faculty of Sciences and Technology - University Nova de Lisboa, for the solid-state nuclear magnetic resonance experiments and all the patience and help with interpretation of those results.

To Dr. Duarte Ananias from Centre for Research in Ceramics and Composite Materials (CICECO) associated laboratory, University of Aveiro for the photoluminescence studies and his help with results interpretation. I also thank CICECO associated laboratory where part of the characterization techniques were carried out. 2 FCUP Acknowledgments

To Dr. Patricia Carvalho for all availability in the matters concerned with transmission electron microscopy and for all the suggestions and contributions related to this work.

To Dr. César Laia and Dr. João Lima from the group of photochemistry of the Chemistry Department, Faculty of Sciences and Technology - University Nova de Lisboa, for full availability on anisotropy and fluorescence lifetime measurements and for all the patience, help and support.

To Dr. Sónia Fraga from the laboratory of toxicology of the Faculty of Pharmacy from University of Porto I would like to thank all the help with the cytotoxic measurements. I also thank all the enthusiasm and the patience during this final stage of my work.

To Fundação para a Ciência e a Tecnologia (FCT) I thank the financial support through the PhD grant SFRH/BD/61137/2009.

Along these last five years in Porto I had the chance to work and live with several persons. Each one of them on their particular way made me feel at home. To my lab colleagues Pedro Quaresma and Leonor Soares I have to thank the way they integrated me in their world, how they helped me during my first steps in nanochemistry and nanotechnology fields. To them and also to others that were coming and going I thank the excellent environment, the fellowship and friendship, and those times that we laugh to tears. Also to Pedro Quaresma I thank the numerous times we discuss strategies and plans even when they were just guesses. To Ana Claudia for her optimism, and trying to make me see the bright side of everything, I know it was a great effort. I would also like to thank Catarina Loureiro, for her friendship and care.

I create many bonds over the years and I couldn’t go without mentioning some of them that were very important to me. First I’d like to thank Sónia Patricio, she was the first person I met when I arrived, she was the face that welcomed me and it was a pleasure to meet her and becoming her friend. To Carla Queirós for her friendship and caring even in my worst moments and a big thank you for helping me whenever I needed. For all the support, help, caring and words of encouragement during these rough times a sincere thanks to Daniela Leite, André Barbosa, Susana Ribeiro and Ana Margarida Silva. FCUP 3 Acknowledgments

I thank in a special way Silvia Lopes (or Lopez like in science people like to call her) the friendship over these years. I thank all the hugs that you gave me, the serious conversations and the ones not so serious, your joy with my victories and your support on my downs. I’ll always remember your laugh and your place in my heart is guaranteed. You are of great importance for me and I hope our paths never separate.

To Vitor Teixeira I thank all the support, help and caring along this journey. Thanks for make me laugh in those tricky moments, for listening and most important just for being there every time I needed.

To Manuela Moreira and Filipe Teixeira two great friends who played an important role during this journey I leave a big thank you. You were the biggest surprises that PDQS offered me. I’ll always remember our adventures in Lisbon, the nights without sleep, the nerves and stress before each presentation, the conversations after classes and all the stupid things we said or done just to make each other laugh. Thank you for all the care and support in good and bad times. I couldn't have better companions by my side during this challenge. We are fighters and together we reached a biggest trophy, our friendship.

To Ana Soares my sister by heart, there is no words to thank you. I know you will always be there for me no matter what, as I will be there for you to.

Roberto Vasconcelos Junior, even with an ocean between us, I know that the size of your friendship will keep us together. Hope to see you soon to celebrate our achievements. Thanks for being the friend that you are. Miss you…

To my mother a special thanks. Thank you for not let me giving up, I wouldn´t be the person I am today if it wasn’t for you and your effort. My accomplishments in life are due to you, to your example of courage, hardworking and dedication. I’m sure I wouldn´t come this far if it wasn´t for the great woman you are.

To Milene my sister I could write dozens of pages, after all we share a life together, but everything I could write wouldn’t be enough to express my gratitude. We overtake many things together and we are here today stronger than never. I love you and you will always be an inspiration to me.

For my love Luis that would turn the world upside down just to see a smile on my face I just don’t have enough words to say thank you. I know I wasn´t the sweetest 4 FCUP Acknowledgments

person along this journey, but nonetheless you’ll never gave up on me. You have been a supporting rock in all the bad moments and in the good ones you have been the most cheerful person. Thank you for believing in me when I doubted, for being a friend, a listener, a supporter and most important my companion for life.

To all of those who contributed somehow to this thesis and prevented me to go insane and aren't mentioned here, I appreciate your effort and caring. I leave you a sincere thanks and the following quote:

“Those who pass us by do not go alone, do not leave us alone. Leave a bit of themself, take a little of us.” (Antoine de Saint-Exupéry)

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AUTHOR PUBLICATIONS CONTAINING WORK RELATED WITH THIS THESIS:

Articles and book chapters in peer-reviewed journals

Published/submitted papers:

Cristina S. Neves, Carlos M. Granadeiro, Luis Cunha-Silva, Duarte Ananias, Sandra Gago, Gabriel Feio, Patricia A. Carvalho, Peter Eaton, Salete S. Balula, Eulália Pereira, Europium-polyoxometalates encapsulated into sílica nanoparticles: characterization and photoluminescence studies, European Journal of Inorganic Chemistry 16, 2877-2886 (2013).

Book chapter:

Pedro V. Baptista, Gonçalo Doria, Pedro Quaresma, Miguel Cavadas, Cristina S. Neves, Inês Gomes, Peter Eaton, Eulália Pereira, Ricardo Franco, Nanoparticles in Molecular Diagnosis, Progress in Molecular Biology and Translational Science, Vol. 104, 427-488, (Antonio Villaverde, Ed.) Elsevier (2011).

Awards:

Young Scientist Award on European Materials Research Society (E-MRS) 2013 Spring Meeting, Strasbourg, France, 26th – 31st May 2013.

Best Paper Award on XI International Conference on Nanostructured Materials, Rhodes, Greece, 23rd – 31st August 2012.

Articles not included in this dissertation, but published in the course of the Ph.D. work:

Silvia C. Lopes, Cristina Neves, Peter Eaton, Paula Gameiro, Improved model systems for bacterial membranes from differing species: the importance of varying composition in PE/PG/cardiolipin ternary mixtures, Molecular Membrane Biology, 29 (2012), 207-217.

Maria J. Medeiros, Cristina S. S. Neves, A. R. Pereira, Elizabeth Duñach, Electroreductive intramolecular cyclisation of bromoalkoxylated derivatives catalized by nickel (I) tetrametylcyclam in “green” media, Electrochimica Acta, 56 (2011), 4498- 4503. 6 FCUP Author publications

Silvia Lopes, Cristina S. Neves, Peter Eaton, Paula Gameiro, Cardiolipin, a key component to mimic the E. Coli bacterial membrane in model systems revealed by dynamic light scattering and steady-state fluorescence anisotropy, Analytical & Bioanalytical Chemistry, 398 (2010) 1357-1366.

Cristina S. Neves, Pedro Quaresma, Pedro V. Baptista, Patricia A. Carvalho, João P. Araújo, Eulália Pereira, Peter Eaton, New insights into the use of magnetic force microscopy to discriminate between magnetic and nonmagnetic nanoparticles, Nanotechnology, 21 (2010) 30576.

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Abstract

Production of nanosized probing (labelling) agents is anticipated to lead to advancements in understanding biological processes at the molecular level, and in the development of diagnostic tools and innovative therapies. Imaging agents such as luminescent silica nanoparticles that can incorporate organic or inorganic fluorophores have overcome many limitations of conventional contrast agents (organic dyes) such as poor photostability, low quantum yield, insufficient in vitro and in vivo stability, etc. For fluorophore-doped silica nanoparticles, the fluorescence source is the encapsulated fluorophore within the matrix of the nanoparticles. Silica encapsulation provides a protective layer around the fluorophore molecules, reducing diffusion of oxygen molecules which are responsible for photodegradation. Furthermore the high stability, chemical inertness and optical transparency of silica make it the ideal candidate for encapsulation while preserving the properties of the encapsulated material, in particularly the optical ones. The surface of silica nanoparticles can be easily functionalized for applications in the preparation of biosensors and cell labelling moreover several luminescent probes can be encapsulated inside the silica matrix.

The research work presented in this thesis aimed the preparation and characterization of luminescent core/shell silica nanoparticles using organic (rhodamine b isothiocyanate - RBITC) and inorganic (lanthanide-based polyoxometalates - LnPOMs) molecules as the source of luminescence.

Luminescent silica nanoparticles were synthesized by hydrolysis and polymerization of tetraethylortoslicate (TEOS) with aqueous ammonia in a water-in-oil reverse microemulsion. In the case of silica nanoparticles containing rhodamine b molecules, the dye was firstly coupled to a silane coupling agent (3-aminopropyl triethoxysilane – APTES), and the reaction product was incorporated into silica spheres using the reverse microemulsion technique for the alkaline hydrolysis of TEOS. The obtained nanoparticles had a mean diameter of 64 nm and were characterized by fluorescence spectroscopy, transmission electron microscopy (TEM) and dynamic light scattering (DLS). Lifetime measurements and steady-state anisotropy studies of the nanoparticles and the free dye were also performed to evaluate the effect of the encapsulation on the fluorescence emission properties of RBITC. Particle’s surface was also modified, so that the particles could bind to biologically active molecules such as oligonucleotides. 8 FCUP Abstract

Luminescent core/shell nanoparticles having LnPOMs as a luminescent source were also prepared. POMs acting as inorganic ligands can coordinate to lanthanide (Ln) ions to form new inorganic compounds with unique properties, such as excellent luminescent characteristics. The LnPOMs used were the mono-substituted 4- 11- [PW11O39Eu(H2O)3] and the sandwich-type [Eu(PW11O39)2] Keggin derivatives. The nanoparticles were characterized using several techniques (FT-IR, FT-Raman, 31P MAS NMR, TEM-EDS, ICP analysis) where the stability of the material and the integrity of the incorporated europium compound was examined. Furthermore, the photo- luminescence properties of the nanomaterials were evaluated and compared with the free LnPOMs. The nanocomposites exhibit a well-defined core/shell structure composed by a LnPOM core surrounded by a silica shell with mean diameters of approximately 16 nm and 51 nm for the mono-substituted and sandwich-type LnPOMs nanocomposites, respectively. Moreover, the silica surface of the most promising nanoparticles was successfully functionalized with appropriate organosilanes to enable the covalent binding to oligonucleotides.

The potential cytotoxicity of the fluorescent silica nanoparticles synthesized was evaluated in three different human cell lines (intestinal epithelial Caco-2 cells, neuroblastoma SH-SY5Y cells and hepatoma HepaRG cells). Nanoparticles cytotoxicity was evaluated by assessing cell viability using the Calcein-AM assay. Phase contrast microscopy was used to evaluate cell morphology and integrity after exposure to the nanoparticles. For the fluorescent silica nanoparticles encapsulating the organic dye RBITC, a cellular uptake experiment was also performed. The obtained results showed that, at the concentrations tested, the nanoparticles presented a non- toxic behavior.

Keywords

Fluorescent silica nanoparticles; rhodamine b isothiocyanate; europium- polyoxometalates; water-in-oil microemulsion; fluorescence lifetime; photoluminescence, cytotoxicity FCUP 9

Resumo

Prevê-se que a produção de sondas nanométricas (marcadores) pode levar a avanços na compreensão de processos biológicos a nível molecular, e ao desenvolvimento de ferramentas de diagnóstico e terapias inovadoras. Agentes de contraste de imagem, tais como nanopartículas de sílica luminescentes, que podem incorporar fluoróforos orgânicos ou inorgânicos, têm superado muitas limitações dos agentes de contraste convencionais (corantes orgânicos), tais como fraca fotoestabilidade, baixo rendimento quântico, insuficiente estabilidade in vitro e in vivo, etc. No caso das nanopartículas de sílica dopadas com fluoróforos, a fonte de fluorescência é fluoróforo encapsulado no interior da matriz das nanopartículas. A encapsulação com uma matriz de sílica proporciona uma camada de protecção em torno das moléculas do fluoróforo, reduzindo a difusão de moléculas de oxigénio, que são responsáveis pela sua fotodegradação. Além disso, a elevada estabilidade da sílica e o facto de ser quimicamente inerte, transparente e preservar as propriedades do material encapsulado, em particular as propriedades ópticas, tornam este material o candidato ideal para a encapsulação. A superfície das nanopartículas de sílica pode ser facilmente funcionalizada para aplicação na preparação de biossensores e marcação celular, além disso várias sondas luminescentes podem ser encapsuladas no interior da matriz de sílica.

O trabalho de pesquisa apresentado nesta tese consistiu na preparação e caracterização de nanopartículas de sílica utilizando como fonte de luminescência moléculas de fluoróforos orgânicos (rodamina b isotiocianato - RBITC) e inorgânicos (polioxometalatos à base de lantanídeos - LnPOMs).

As nanopartículas de sílica luminescentes foram sintetizadas através de uma microemulsão inversa de água-em-óleo, por meio de hidrólise e polimerização do tetraetilortoslicato (TEOS) na presença de amoníaco. No caso das nanopartículas de sílica contendo moléculas de rodamina b, o corante foi primeiramente acoplado a um agente de acoplamento de silano (3-aminopropil-trietoxisilano - APTES), e o produto da reacção foi incorporado em esferas de sílica, utilizando a técnica de microemulsão inversa para a hidrólise alcalina do TEOS. As nanopartículas obtidas possuem um diâmetro médio de 64 nm e foram caracterizadas por espectroscopia de fluorescência, microscopia electrónica de transmissão (TEM) e dispersão dinâmica de luz (DLS). Foram também realizadas medições de tempo de vida e estudos de anisotropia em 10 FCUP Resumo

estado estacionário das nanopartículas e do corante livre, para avaliar o efeito da encapsulação sobre as propriedades de emissão de fluorescência da RBITC. A superfície das partículas foi também modificada de modo a que se possam ligar a moléculas biologicamente activas, tais como, oligonucleótidos.

Também foram preparadas nanopartículas luminescentes do tipo “core-shell” usando como fonte luminescente os LnPOMs. Os POMs atuam como ligandos inorgânicos e podem coordenar com iões lantanídeos (Ln) para formar novos compostos inorgânicos com propriedades únicas, tais como excelente luminescência. 4- Foram utilizados LnPOMs do tipo Keggin, um monosubstituído [PW11O39Eu(H2O)3] e 11- outro do tipo sanduíche [Eu(PW11O39)2] . As nanopartículas foram caracterizadas utilizando várias técnicas (FT - IR, FT - Raman, 31P RMN MAS, TEM- EDS, análise de ICP), em que a estabilidade do material e a integridade do composto de európio incorporado foi examinada. Além disso, as propriedades de fotoluminescência dos nanomateriais foram avaliadas e comparadas com as dos LnPOMs não encapsulados. Os nanocompósitos apresentam uma estrutura “core-shell” bem definida, composto por um núcleo de LnPOM rodeado por uma capa de sílica, com diâmetros médios de cerca de 16 nm e 51 nm para os nanocompósitos encapsulando o LnPOMs monosubstituído e do tipo sandwich respectivamente. Além disso, a superfície das nanopartículas de sílica mais promissoras foi funcionalizada com organosilanos para permitir a ligação covalente com oligonucleótidos.

A potencial citotoxicidade das nanopartículas de sílica fluorescentes foi avaliada em três diferentes linhas celulares humanas (células epiteliais intestinais Caco-2, células de neuroblastoma SH-SY5Y e células hepáticas HepaRG). A citotoxicidade das nanopartículas foi avaliada através de ensaios de viabilidade celular utilizando o ensaio de Calceína-AM. A microscopia de contraste de fase foi utilizada para avaliar a morfologia e a integridade das células, após exposição às nanopartículas. No Caso das nanopartículas de sílica fluorescentes encapsulando o corante orgânico RBITC foi também realizada, uma experiência de captação celular. Os resultados obtidos mostraram que, nas concentrações testadas, as nanopartículas apresentam um comportamento não tóxico.

Palavras-chave Nanopartículas fluorescentes de sílica; rodamina b isotiocianato; európio- polioxometalatos; microemulsão de água em óleo; fotoluminescência; decaimento de fluorescência; citotoxicidade FCUP 11

General Index

Acknowledgments ...... 1

Abstract ...... 7

Resumo ...... 9

List of Tables ...... 17

List of Figures ...... 19

Abbreviations and Symbols...... 27

I - Introduction

1. An overview of nanoscale fluorescent materials ...... 33

1.1. Organic dye molecules ...... 34

1.1.1. Fluoresceins and rhodamines ...... 34

1.1.2. Cyanine dyes ...... 35

1.1.3. Alexa dyes ...... 36

1.2. Fluorescent Proteins ...... 37

1.3. Semiconductor quantum dots ...... 38

1.4. Dyed polymer nanoparticles ...... 40

1.5. Fluorophore doped silica nanoparticles ...... 40

1.6. References ...... 42

2. Fluorescent silica nanoparticles doped with organic dyes...... 45

2.1. Nanoparticle formation ...... 46

2.1.1. Stöber method ...... 46

2.1.2. Microemulsion method ...... 47

2.1.3. Incorporation of organic fluorophores ...... 48

2.2. Nucleation and growth of silica NPs ...... 49

2.2.1. Nucleation mechanisms ...... 49

2.2.2. Growth mechanisms ...... 50

2.2.3. Particle growth in the reverse micellar system ...... 50 12 FCUP General Index

2.3. Surface functionalization of dye-doped silica NPs ...... 51

2.4. Nanoparticle characterization ...... 54

2.4.1. Measure particle size ...... 54

2.4.2. Surface charge ...... 55

2.5. Biological applications of dye-doped silica NPs ...... 55

2.5.1. Cell targeting using dye-doped silica NPs ...... 56

2.5.2. Dye-doped silica NPs as intracellular nanosensors ...... 56

2.5.3. Dye-doped silica NPs for multiplexed bioanalysis ...... 57

2.5.4. Dye-doped silica NPs for nucleic acid analysis ...... 58

2.6. References ...... 60

3. Lanthanopolyoxometalates encapsulated into silica nanoparticles ...... 63

3.1. Polyoxometalates ...... 63

3.1.1. Definition ...... 63

3.1.2. Historical context ...... 65

3.1.3. Preparation ...... 66

3.2. Keggin anion ...... 67

3.3. Polyoxometalates containing lanthanide ions ...... 70

n- 3.3.1. Keggin-type lanthanide polyoxometalates ([Ln(XM11O39)y] ) ...... 70

3.3.2. Luminescence of lanthanide ions ...... 72

3.4. Silica encapsulation of LnPOMs ...... 75

3.5. Applications of Keggin-type LnPOMs ...... 76

3.6. References ...... 79

II – Research work

Scope of the thesis ...... 89

Experimental Background ...... 91

References ...... 97

4. Dye doped fluorescent silica nanoparticles ...... 99

4.1. Material and Methods ...... 99 FCUP 13 General Index

4.1.1 Chemicals ...... 99

4.1.2 Instrumentation and methodologies ...... 100

4.1.1.1. Elemental Analysis ...... 100

4.1.1.2. UV-visible spectroscopy ...... 100

4.1.1.3. Fluorescence spectroscopy, quantum yield and lifetime ...... 100

4.1.1.4. Steady-state anisotropy ...... 101

4.1.1.5. Transmission electron microscopy ...... 102

4.1.1.6. Dynamic light scattering and zeta potential ...... 102

4.1.3 Preparation of core-shell nanoparticles with rhodamine B isothiocyanate

(RBITC-APTES@SiO2) ...... 102

4.1.4 Surface functionalization of silica nanoparticles ...... 103

4.1.5 DNA grafting ...... 103

4.2. Results and Discussion ...... 104

4.2.1 Characterization of RBITC@SiO2 nanoparticles ...... 104

4.2.1.1. Characterization by electron microscopy ...... 104

4.2.1.1. Dynamic light scattering ...... 106

4.2.1.2. Characterization by UV-vis spectroscopy ...... 107

4.2.1.3. Fluorescence excitation and fluorescence emission spectra ...... 110

4.2.1.4. Fluorescence quantum yield ...... 112

4.2.1.5. Lifetime measurements ...... 115

4.2.1.6. Fluorescence anisotropy ...... 119

4.2.2 Characterization of RBITC-APTES@SiO2 NPs grafted with DNA ...... 122

4.2.3 Conclusions ...... 124

4.3. References ...... 127

5. Europium polyoxometalates encapsulated into silica nanoparticles ...... 131

5.1. Materials and Methods ...... 131

5.1.1. Chemicals ...... 131

5.1.2. Instrumentation and methodologies ...... 132 14 FCUP General Index

5.1.2.1. Elemental analysis ...... 132

5.1.2.2. Vibrational Spectroscopy ...... 132

5.1.2.3. Solid state NMR ...... 132

5.1.2.4. Transmission electron microscopy ...... 133

5.1.2.5. Scanning electron microscopy ...... 133

5.1.2.6. Dynamic light scattering ...... 133

5.1.2.7. Atomic force microscopy ...... 133

5.1.2.8. X-ray crystallography ...... 134

5.1.2.9. Photoluminescence and lifetime measurements ...... 134

5.1.2.10. Quantum efficiency ...... 135

5.1.3. Synthesis of europium polyoxometalates Eu(PW11)x (x = 1 and 2) ...... 136

5.1.4. Encapsulation of Eu(PW11)x (x = 1 and 2) into silica nanoparticles ...... 136

5.1.5. Functionalization of Eu(PW11)2@SiO2 ...... 137

5.2. Results and Discussion ...... 137

5.2.1. Characterization of Eu(PW11)x compounds ...... 138

5.2.1.1. X-ray crystallography ...... 138

5.2.1.2. Thermogravimetry ...... 140

5.2.1.3. 31P NMR spectroscopy ...... 141

5.2.2. Characterization of Eu(PW11)x@SiO2 nanoparticles ...... 142

5.2.2.1. Transmission Electron Microscopy ...... 143

5.2.2.1. Scanning Electron Microscopy ...... 146

5.2.2.2. Characterization by vibrational spectroscopy ...... 148

5.2.2.3. Photoluminescence properties ...... 151

5.2.3. Conclusions ...... 155

5.3. References ...... 156

6. Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles .... 161

6.1. Cytotoxicity assays ...... 162

6.2. Materials and Methods ...... 163 FCUP 15 General Index

6.2.1. Chemicals ...... 163

6.2.2. Cellular culture ...... 164

6.2.3. Nanoparticle uptake ...... 164

6.2.4. Cell viability by Calcein-AM assay ...... 164

6.2.5. Phase contrast microscopy ...... 165

6.2.6. Fluorescence spectroscopy ...... 165

6.2.7. Transmission electron microscopy ...... 165

6.2.8. Statistical analysis ...... 165

6.3. Results and Discussion ...... 165

6.3.1. Cellular uptake of silica nanoparticles ...... 166

6.3.2. Cell esterase activity (Calcein-AM assay) ...... 168

6.3.2.1. Effect of RBITC@SiO2 NPs on Caco-2, SH-SY5Y and Hepa RG cells viability ...... 169

6.3.2.2. Effect of Eu(PW11O39)2@SiO2 NPs on Caco-2, SH-SY5Y and Hepa RG cells viability ...... 172

6.3.3. Morphological analysis by phase contrast microscopy ...... 174

6.3.3.1. RBITC-APTES FSNPS ...... 174

6.3.3.2. Eu(PW11O39)2@ SiO2 NPs ...... 177

6.4. Conclusions ...... 179

6.5. References ...... 181

III – Concluding Remarks and Perspectives

Concluding Remarks and Perspectives...... 187 FCUP 16

FCUP 17

List of Tables

I - Introduction

Table 2.1 - Chemical binding for bioconjugation of silica NPs. (adapted from Yao et al. [2]) ...... 53

Table 3.1 - Commonly observed emission bands of the lanthanide ions Eu3+, Tb3+, Nd3+, Er3+ and Yb3+ in solution. (Adapted from Werts[50]) ...... 74

II – Research work

Table EB 1– Comparison between the three methods followed to prepare fluorescent silica NPs using the microemulsion technique...... 95

Table 4.1 - Average hydrodynamic diameter of RBITC FSNPs measured by DLS (by percentage of number of particles, measurements were repeated 5 times for each sample)...... 106

Table 4.2 – Amount of RBITC dye molecules per fluorescent silica nanoparticle ...... 110

Table 4.3 - Fluorescence quantum yields of RBITC, RBITC-APTES conjugate and FSNPs ...... 113

Table 4.4 - Lifetime data of RBITC and fluorescent silica nanoparticles (FSNPs) in absolute ethanol ...... 115

Table 4.5 - Anisotropy (r) and rotational diffusion coefficient (Dr) values of RBITC and fluorescent silica nanoparticles (FSNPs) adsorbed and covalently bound to silica NPs in absolute ethanol...... 120

Table 4.6 – Zeta potential ζ of FSNPs-GPTES and FSNPs-GPTES-DNA ...... 123

Table 5.1 - Crystal and structure refinement data for Eu(PW11)2 ...... 140

5 Table 5.2 - Experimental D0 lifetime, τ, radiative, kr, and non-radiative, knr, transition 5 rates and D0 quantum efficiency, q, for compounds Eu(PW11)2 and Eu(PW11)2@SiO2. The data have been obtained at room temperature (296 K)...... 154

III – Concluding Remarks and Perspectives

18 FCUP

FCUP 19

List of Figures

I - Introduction

Figure 1.1 - Several common fluorescent nanoscale materials including (a) organic dye molecules (tetramethylrhodamine); (b) green fluorescent protein; (c) polymer-coated, water soluble semiconductor quantum dots and (d) fluorophore-doped silica particles. (Adapted from Burns et al.[3]) ...... 33

Figure 1.2 - Plain and ball-and-stick structures of fluorescein isothiocyanate (FTIC). .. 34

Figure 1.3 – Plain and ball-and-stick structures of rhodamine 6G (top) and rhodamine b (bottom)...... 35

Figure 1.4 - Basic structure of cyanine dyes...... 36

Figure 1.5 – Plain and ball-and-stick structures of Alexa Fluor 350...... 36

Figure 1.6 – Plain and ball-and-stick structures of Alexa Fluor 430...... 37

Figure 1.7 - Structure of the Aequorea victoria green fluorescent protein. (Source: Ormö et al.[15]) ...... 38

Figure 1.8 - Photograph and spectra of CdSe quantum dots. The samples represent different sizes of QDs, which produce different colours upon UV light. An increase in particle size produces a red shift in the emission spectra. (Source: Nauman et al. [17]) ...... 39

Figure 1.9 - Schematic illustration of the surface functionalization of silica NPs with, for example, peptides, antibodies, aptamers, enzymes, DNA-fragments and different functional moieties. (Source: Schulz et al.[21]) ...... 41

Figure 2.1 - TEM images: (A) silica-based nanoparticles prepared by the Stöber method; and (B) silica nanoparticles prepared by the microemulsion process...... 47

Figure 2.2 - Typical structure of a reverse micelle (source: Malik et al.[11]) ...... 48

Figure 2.3 – Silica nanoparticles growth mechanism in a reverse micellar system composed. (Source: Osseo-Asare et al.[21]) ...... 51

Figure 2.4 - Schematic illustration of the surface functionalization of silica NPs for biological applications. (Source: Smith et al.[4]) ...... 52

Figure 2.5 - Representative bioconjugation schemes for attaching biomolecules to dye- doped silica NPs for bioanalysis. (source: Wang et al. [3]) ...... 54 20 FCUP List of Figures

Figure 2.6 - Confocal fluorescence microscopy images (overlaid and bright field)of pH sensors in rat basophilic leukemia mast cells showing a) reference dye (RBITC) channel, b) sensor dye (FTIC) channel, c) overlaid images and d) false-colour ratiometric imaging of pH in various intracellular compartments (Source: Burns et al.[30]) ...... 57

Figure 2.7 - Schematic representation of a sandwich assay based on dye-doped silica NPs. (Source: Zhao et al.[33]) ...... 59

Figure 3.1 - Ball-and-stick (left) and polyhedral (right) representations of the [16] fundamental unit MO6. (Source: Fernandez ) ...... 64

Figure 3.2 - Representation of the three possible unions between two MO6 octahedral units: A) corner-sharing, B) edge-sharing and C) face-sharing. Each corner represents an oxygen position. (Source: Fernandez[16]) ...... 64

Figure 3.3 - Polyhedral representation of common polyoxoanions: A) Lindqvist n- n- n- ([M6O19) ) isopolyanion; B) Anderson-Evans ([XM6O24] ); C) Keggin ([XM12O40] ); D) n- n- Wells-Dawson ([X2M18O62] ) and E) Preyssler ([XP5W30O110] ) heteropolyanions. (Source: Lopez et al. [22]) ...... 65

Figure 3.4 - Polyhedral representation of the Keggin structure showing the four groups

M3O13 in four different colors and the central tetrahedron XO4 in yellow. (Source: Al- Kadamany[35]) ...... 67

Figure 3.5 - Polyhedral representation of the five rotational isomers of the Keggin [22] anion. The rotated M3O13 groups are highlighted (dark blue). (source: Lopez et al. ) 68

Figure 3.6 - Ball and stick (left) and polyhedral representation (right) for the α- n- [XM12O40] Keggin anion showing the different classification of the oxygen atoms...... 68

(n+4)- Figure 3.7 - Formation scheme of the monolacunar anion [XM11O39] ...... 69

n- Figure 3.8 - Representation of the complexes of the type 1:1 [XM11M’(L)O39] (left) and n- 1:2 [M’(XM11O39)2] (right)...... 69

Figure 3.9 - Formation scheme of the monolacunar (A) and sandwich type (B) n- lanthanide-substituted Keggin anion [Ln(XM11O39)x] ...... 71

Figure 3.10 – Photoluminescence emission spectra of the Eu3+ ion in water. The 5 [50] radiative transitions take place from the D0 level. (Adapted from Werts ) ...... 73

Figure 3.11 - Interactions leading to the different electronic energy levels for Eu3+ configuration ([Xe] 4f65d0 – six electrons in the 4f orbitals). (Source: Werts[50]) ...... 75 FCUP 21 List of Figures

II – Research work

Figure EB 1 - UV-vis spectrum of fluorescent silica nanoparticles synthesized by Stober’s method through the adapted procedure described by Bringley[1]...... 91

Figure EB 2- TEM images of TRICT fluorescent silica nanoparticles prepared by Stöber’s method following a similar procedure to that described by Larson[3] et al...... 93

Figure EB 3 - TEM images of fluorescent silica nanoparticles prepared by the reverse microemulsion system following the procedures of Gao[4] (A); Zhang[5] (B) and Shi[6] (C)...... 96

Figure EB 4 - Fluorescence emission spectra of fluorescent silica nanoparticles prepared by the reverse microemulsion system following the procedures of Gao[4] (A); Zhang[5] (B) and Shi[6] (C)...... 96

Figure 4.1 - TEM images of RBITC-APTES FSNPS nanoparticles and corresponding size distribution histogram...... 105

Figure 4.2 - SEM images of RBITC-APTES FSNPs showing the spherical morphology of the NPs...... 105

Figure 4.3 - DLS hydrodynamic diameter distribution statistics graph (by percentage of number of particles) for RBITC FSNPS. Error bars show standard deviation of five different measurements...... 107

Figure 4.4 - UV-vis spectra of RBITC and RBITC fluorescent silica NPs (FSNPs) in ethanol at 25 ºC. UV-vis spectrum of FSNPs was fitted using a 2nd order exponential decay to remove silica scattering. Both samples were dissolved to a final concentration with almost the same absorbance (0.09)...... 108

Figure 4.5 - UV-vis spectra of RBITC and RBITC-APTES conjugate in ethanol at 25 ºC...... 109

Figure 4.6 - Fluorescence excitation spectra of RBITC, RBITC-APTES conjugate and FSNPs recorded at 25 ºC in absolute ethanol...... 111

Figure 4.7 - Fluorescence emission spectra of RBITC, RBITC-APTES conjugate and FSNPs recorded at 25 ºC in absolute ethanol...... 112

Figure 4.8 - (A) RBITC doped fluorescent silica NPs prepared by hydrolysis and polymerization of TEOS in a microemulsion method; (B) bare silica NPs with RBITC dye molecules adsorbed onto the nanoparticle’s surface; (C) fluorescent core-shell NPs with a silicon core and a shell of RBITC dye molecules and TEOS...... 113 22 FCUP List of Figures

Figure 4.9 - Fluorescence lifetime decay curves of RBITC (A), RBITC-APTES conjugate (B), RBITC-APTES FSNPs with dye covalently bound to silica matrix

(RBITC-APTES@SiO2) (C),and RBITC-APTES FSNPs with dye adsorbed to silica

surface (Ads:RBITC-APTES@SiO2) (D), all at ambient temperature (298 K) .The excitation was fixed at 370 nm and the emission was monitored at 550 nm...... 116

Figure 4.10 – Structures of rhodamine b isothiocyanate (RBITC) isomers. Left: rhodamine b 5-isothiocyanate and right: rhodamine b 6-isothiocyanate...... 117

Figure 4.11 – Structures of RBITC-APTES conjugate for RBITC 5-isomer (top) and RBITC 6-isomer (bottom) ...... 118

Figure 4.12 - Steady state emission fluorescence anisotropy of RBITC and RBITC

FSNPs with dye adsorbed (Ads:RBITC-APTES@SiO2) and dye covalently bound

(RBITC-APTES@SiO2) to silica matrix. The excitation wavelength was 530 nm...... 120

Figure 4.13 - Representation of the wobbling-in-cone model, where θc is the angle between the probe (dye) axis (direction of the optical transition moment) and the symmetry axis of the wobbling motion (cone axis)...... 121

Figure 4.14 - Strategy for immobilisation of thiolated oligonucleotides onto dye loaded silica nanoparticle surfaces...... 122

Figure 4.15 - UV-vis spectra of DNA and functionalized FSNPS before (FSNPs- GPTES) and after (FSNPs-GPTES-DNA) DNA immobilization in potassium phosphate buffer (10 mM, pH = 8). Inset: zoom in the UV-vis spectra of FSNPS before and after DNA immobilization...... 123

Figure 5.1 - (a) The structures of the sandwich type europium-phosphotungstate anion, 11− [Eu(PW11O39)2] ; (b) its {EuO8} coordination center displaying a square-antiprismatic geometry and (c) the mono-substituted europium-phosphotungstate anion, 4- [PW11Eu(H2O)3O39] drawn in polyhedral and ball-and-stick mixed model...... 138

Figure 5.2 - Thermogravimetric curves of EuPW11 (in blue) and Eu(PW11)2 (in red). 141

31 Figure 5.3 - P NMR spectra of monovacant precursor PW11 and Eu(PW11)x in D2O solution...... 142

Figure 5.4 - TEM images of (a,b) EuPW11@SiO2 and (d,e) Eu(PW11)2@SiO2 nanoparticles showing the core/shell structure (both materials prepared using 50 mg of corresponding europium compounds); (c,f) Size distribution histograms of

EuPW11@SiO2 and Eu(PW11)2@SiO2 nanoparticles respectively...... 143 FCUP 23 List of Figures

Figure 5.5 - EDS spectra of silica nanoparticles of mono-substituted compound

EuPW11@SiO2 and the sandwich-type Eu(PW11)2@SiO2 (both materials prepared using 50 mg of corresponding europium compounds). The copper peak comes from the support grid...... 144

Figure 5.6 - AFM topography and amplitude images respectively of EuPW11 (a,b) and

Eu(PW11)2 (c,d). Topography images show the presence of features with dimensions larger than expected for single POMs (i.e. larger than 2 nm)...... 145

Figure 5.7 - TEM images of (a,b) Eu(PW11)2@SiO2 NPs prepared using 95 mg (16 µmol) of corresponding europium polyoxometalate.; (c) Size distribution histogram of the mentioned EuPW11@SiO2 NPs. For direct comparison size distribution the histogram of EuPW11@SiO2 (d) NPs prepared using 8 µmol of the same europium polyoxometalate is also presented...... 146

Figure 5.8 - (a) STEM image of EuPW11@SiO2 NPs; (b) overlapping of EDS mapping for Si (red) and W (green), (c, d) separated EDX mapping for Si and W respectively 147

Figure 5.9 - (a) STEM image of Eu(PW11)2@SiO2 NPs; (b) EDS spectra of

Eu(PW11)2@SiO2, (c, d) separated EDS mapping for Si and W respectively. The Copper (Cu), aluminium (Al) and tin (Sn) peaks come from the support grid...... 147

Figure 5.10 - FT-IR spectra for EuPW11 (left) and for Eu(PW11)2 (right) and its corresponding core/shell nanoparticles with and without functionalization prepared using equal weight of europium-polyoxometalate...... 148

Figure 5.11 - FT-Raman spectra for EuPW11 (A) and for Eu(PW11)2 (B) and the same particles in silica-coated core:shell form, with and without functionalization (both materials prepared using 50 mg of corresponding europium compound)...... 149

31 Figure 5.12 - Solid state P MAS NMR spectra of the monovacant precursor PW11, potassium salt EuPW11 and its corresponding silica-coated core/shell nanoparticles (A), and of potassium salt Eu(PW11)2 and their corresponding silica-coated core/shell nanoparticles with and without functionalization (B). All nanoparticles prepared using 50 mg of corresponding europium compound...... 150

Figure 5.13 - Excitation spectra of EuPW11 (A) and Eu(PW11)2 (B) and their corresponding core:shell nanoparticles at ambient temperature (298 K, black lines) and 14 K (red lines) while monitoring the emission at 614 nm...... 151

Figure 5.14 - Ambient temperature (298 K) emission spectra of EuPW11 (A) and

Eu(PW11)2 (B) and their corresponding core:shell nanoparticles at ambient conditions 24 FCUP List of Figures

(black lines, pressure of 1 bar) and with a high vacuum (red lines, pressure of ca. 5×10- 6 mbar). The excitation was fixed at 394 nm...... 152

3+ 5 Figure 5.15 - Eu D0 decay curves of EuPW11 (A) and Eu(PW11)2 (B) and its corresponding core/shell nanoparticles at ambient temperature (298 K) and pressure (1 bar). The excitation was fixed at 394 nm and the emission was monitored at ca. 614 nm...... 154

Figure 6.1 - TEM images of RBITC-APTES FSNPs dried from high glucose DMEM at a concentration of 3.2 µg/ml...... 167

Figure 6.2 - (A) Fluorescence excitation and emission spectra of stock solution of RBITC-APTES FSNPs (1.6mg/ml) in ethanol and RBITC-APTES FSNPs solution (3.2 µg/ml) in high glucose, phenol red-free DMEM ; (B) zoom of the fluorescence excitation and emission spectra of RBITC-APTES FSNPs solution (3.2 µg/ml) in high glucose, phenol red-free DMEM...... 168

Figure 6.3 - Effect of RBITC-APTES@SiO2 NPs, RBITC and SiO2 NPs on esterase activity of human intestinal epithelial Caco-2 cells, as assessed by the calcein-AM assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage of control (vehicle-treated cells) and data are presented as mean ± SEM (n=4-28 per group)...... 169

Figure 6.4 - Effect of RBITC-APTES@SiO2 NPs, RBITC and SiO2 NPs on esterase activity of human neuroblastoma SH-SY5Y cells, as assessed by the calcein-AM assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage of control (vehicle-treated cells) and data are presented as mean ± SEM (n=4-28 per group)...... 169

Figure 6.5 - Effect of RBITC-APTES@SiO2 NPs, RBITC and SiO2 NPs on esterase activity of human hepatoma Hepa RG cells, as assessed by the calcein-AM assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage of control (vehicle-treated cells) and data are presented as mean ± SEM (n=8-28 per group). . 170

3+ Figure 6.6 - Effect of Eu(POMs)@SiO2 NPs, Eu , POMs and and SiO2 NPs on esterase activity of human intestinal epithelial Caco-2 cells, as assessed by the calcein-AM assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage of control (vehicle-treated cells) and data are presented as mean ± SEM (n=8-24 per group)...... 172

3+ Figure 6.7 - Effect of Eu(POMs)@SiO2 NPs, Eu , POMs and SiO2 NPs on esterase activity of human neuroblastoma SH-SY5Y cells, as assessed by the calcein-AM FCUP 25 List of Figures

assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage of control (vehicle-treated cells) and data are presented as mean ± SEM (n=8-24 per group)...... 173

3+ Figure 6.8 - Effect of Eu(POMs)@SiO2 NPs, Eu , POMs and and SiO2 NPs on esterase activity of human hepatoma Hepa RG cells, as assessed by the calcein-AM assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage of control (vehicle-treated cells) and data are presented as mean ± SEM (n=8-28 per group)...... 173

Figure 6.9 - Representative phase contrast microscopy images of Caco-2 cells at 24 hours after incubation with SiO2 nanoparticles, RBITC dye and RBITC-APTES@SiO2 nanoparticles (100x magnification)...... 175

Figure 6.10 - Representative phase contrast microscopy images of SH-SY5Y cells at

24 hours after incubation with SiO2 nanoparticles, RBITC dye and RBITC-

APTES@SiO2 nanoparticles (100x magnification)...... 176

Figure 6.11 - Representative phase contrast microscopy images of Hepa RG cells at

24 hours after incubation with SiO2 nanoparticles, RBITC dye and RBITC-

APTES@SiO2 nanoparticles (100x magnification)...... 177

Figure 6.12 - Representative phase contrast microscopy images of Caco-2 cells at 24 hours after incubation with SiO2 nanoparticles, Eu salt, Eu(PW11O39)2 and

Eu(PW11O39)2@SiO2 nanoparticles (100x magnification)...... 178

Figure 6.13 - Representative phase contrast microscopy images of SH-SY5Ycells at 24 hours after incubation with SiO2 nanoparticles, Eu salt, Eu(PW11O39)2 and

Eu(PW11O39)2@SiO2 nanoparticles (100x magnification)...... 178

Figure 6.14 - Representative phase contrast microscopy images of Hepa RGcells at 24 hours after incubation with SiO2 nanoparticles, Eu salt, Eu(PW11O39)2 and

Eu(PW11O39)2@SiO2 nanoparticles (100x magnification)...... 179

III – Concluding Remarks and Perspectives 26 FCUP

FCUP 27

Abbreviations and Symbols

AFM - atomic force microscopy

APTES - 3-(aminopropyl)triethoxysilane

ATP - Adenosine TriPhosphate

BSA – bovine serum albumin

Caco-2 - Human epithelial colorectal adenocarcinoma cells

CCK-8 – cell counting kit 8

CPTES – (3-chloropropyl)-trimethoxysilane

CMC – carboxymethyl chitosan

CTES – carboxyethylsilanetriol

Cy3 and Cy5 – cyanine dyes 3 and 5 respectively

DLS - dynamic light scattering

DMEM - Dulbecco’s modified eagle’s medium

DNA - deoxyribonucleic acid

EDS - X-ray spectroscopy

FRET - fluorescence resonance energy transfer

FSNPs - fluorescent silica nanoparticles

FTIC - fluorescein isothiocyanate

FT-IR – Fourier transform infrared spectroscopy

FT-Raman - Fourier transform Raman spectroscopy

Gd-DTPA – gadolinium-diethylenetriaminepntaacetic acid complex 28 FCUP Abbreviations and Symbols

GFP - green fluorescent protein

GPTMS - 3-glycidoxypropyltrimethoxy silane

HBSS – Hank’s balanced salt solution

HC – high component lifetime

HEPA RG - Human hepatoma cells

HepG2 – Human hepatocellular liver carcinoma cells

ICP-MS – Inductively coupled plasma mass spectroscopy

LC – low component lifetime

LMCT- ligand-to-metal charge transfer

Ln - lanthanide

LnPOMs - lanthanide-substituted polyoxometalates

MAS – Magic-angle spinning

MB - methylene blue

MLCT - metal-to-ligand charge transfer

MOF - metal-organic-framework

MPTS - 3-mercaptopropyltrimethoxysilane

MRI – magnetic resonance imaging

MTT - 3-(4, 5-dimethyl-2-thiazolyl)-2, 5-diphenyl-2H-tetrazolium bromide

NMR – nuclear magnetic resonance

NP - nanoparticle

NPs – nanoparticles

ODS – oxidative desulfurization FCUP 29 Abbreviations and Symbols

PEG - poly(ethylene glycol)

PEBBLEs – probes encapsulated by biologically localized embedding

POMs – polyoxometalates

PVA - polyvinyl alcohol

QDs - Quantum Dots

R6G - rhodamine 6G

RBITC - rhodamine b isothiocyanate

RBITC 5-isomer - rhodamine b-5-isothiocyanate

RBITC 6-isomer - rhodamine b-6-isothiocyanate

ROS – reactive oxygen species

ROX - 6-carboxyl-X-rhodamine

Rubpy - tris(bipyridine)ruthenium(II) dichloride

SEM - scanning electron microscopy

SEPs – surfactant encapsulated polyoxometalates

SH-SY5Y - Human neuroblastoma cells

Si – silicon

SMM - single molecular magnets

TEM - transmission electron microscopy

TEOS - tetraethylortosilicate

TG – thermogravimetry

TMR – tetramethylrhodamine

TMR-Dex – tetramethylrhodamine dextran 30 FCUP Abbreviations and Symbols

TRITC - tetramethylrhodamine isothiocyanate

UV-vis – ultraviolet-visible spectroscopy

W/O - water-in-oil microemulsion

W0 - water-to-surfactant molar ratio

ε – absorption coefficient

ζ - zeta potential

I Introduction

FCUP 33

1. An overview of nanoscale fluorescent materials

Nanoscale materials are a broadly defined set of substances that have at least one critical dimension less than 100 nm and possess unique optical, magnetic, electrical or other properties.[1] Thus, when particle size is be nanoscale, properties such as melting point, fluorescence, electrical conductivity, magnetic permeability, and chemical reactivity can change as a function of the size of the particle.

Nanoscale fluorescent materials gained particular interest in the fields of chemistry, biology, medical science and biotechnology.[2] Because nanoscale fluorescent particles may not be visible to the naked eye, it is possible to use them as hidden fluorescent substances which only become fluorescent once they have been excited by light of a specific wavelength. This property makes them of particular interest for biological applications where in past decades many progresses have been made.

Due to their high signal-to-noise ratio, excellent spatial resolution, and ease of implementation, fluorescent materials are ideal to investigate biology down to nanoscale.[3, 4] Every application has its own particular restrictions, but the most important properties for any fluorescent material are the same: brightness and stability. There are several classes of materials currently employed as fluorescent emitters/probes, which includes organic dye molecules, fluorescent proteins, semiconductor quantum dots, polymer/dye-based nanoparticles and silica/fluorophores hybrid particles (Figure 1.1), and each of them have their own advantages and disadvantages.

Figure 1.1 - Several common fluorescent nanoscale materials including (a) organic dye molecules (tetramethylrhodamine); (b) green fluorescent protein; (c) polymer-coated, water soluble semiconductor quantum dots and (d) fluorophore-doped silica particles. (Adapted from Burns et al.[3]) 34 FCUP An overview of nanoscale fluorescent materials

1.1. Organic dye molecules

Organic dye molecules are the smallest fluorescent emitters used today. These fluorophores are commercially available with emissions from UV to the near infrared region of the spectrum (~300-900nm). These dye molecules have a small size (~1nm) which makes them an excellent choice for many applications especially in biology where they are the most commonly used fluorophores. Although they present some limitations like short stokes shifts (difference between maximum wavelengths of absorption and emission bands), rapid photobleaching, poor photochemical stability and decomposition under repeated excitation, the organic dyes are still widely used due to their low cost, availability and ease of usage. Examples of commonly used organic dyes include fluorescein, rhodamine, cyanine and Alexa dyes.

1.1.1. Fluoresceins and rhodamines

Fluoresceins are amine-reactive organic fluorophores widely used in biolabelling.[5, 6] They belong to the xanthene class of dyes[7] with absorption and fluorescence

maxima in the visible region (e.g. fluorescein λabs = 490 nm and λem = 512 nm in water). Fluoresceins have high extinction coefficient and quantum yield and also high solubility in water. However they present some major drawbacks such as photobleaching, pH sensitivity, relatively broad emission spectra and tendency for self-quenching after bioconjugation.[5, 8, 9] As a result the use of fluorescein dyes in ultra-sensitive biological studies is limited.

Figure 1.2 - Plain and ball-and-stick structures of fluorescein isothiocyanate (FTIC). FCUP 35 An overview of nanoscale fluorescent materials

Like fluoresceins, rhodamine dyes also belong to the xanthene class of dyes. Rhodamine dyes have strong absorption and emission spectra in the visible region and many derivatives are strongly fluorescent. When compared with fluorescein dyes rhodamine dyes are more photostable and less sensible to pH.[5] The rhodamine dyes such as rhodamine 6G, Texas red or rhodamine B are well known as fluorescent markers in biology. However, poor water solubility of rhodamine dyes limits their application in biolabelling.

Figure 1.3 – Plain and ball-and-stick structures of rhodamine 6G (top) and rhodamine b (bottom).

1.1.2. Cyanine dyes

Cyanine dyes belong to a family of long wavelength fluorophores extensively used in fields like photography, lasers and more recently in biolabelling and cell imaging.[5, 10] As represented in Figure 1.4 the basic structure of cyanine dyes includes two aromatic or heterocyclic rings linked by a polymethine chain with conjugated carbon-carbon double band.[11] These dyes present emission spectra in the range of 600-900 nm and a high extinction coefficient (>10000 M-1.cm-1). 36 FCUP An overview of nanoscale fluorescent materials

X Y

N n N

R R

- X, Y = O, S, C(CH3)2 or C = CH2; n = 0-4; R = (CH2)xSO3

Figure 1.4 - Basic structure of cyanine dyes.

However, there are some drawbacks in the use of cyanine dyes that include low availability of these dyes as labeling probes, short fluorescence lifetimes and low fluorescence quantum yield. Apart from that cyanine dyes tend to aggregate in aqueous solution leading to low fluorescence intensities.[11]

1.1.3. Alexa dyes

Alexa dyes are a group of new fluorescent molecules resulting from the sulfonation of aminocoumarin, rhodamine or cyanine dyes. The excitation and emission wavelengths range of Alexa dyes cover the entire spectrum from ultraviolet to red and match the principal output wavelengths of common excitation sources. These dyes are generally more stable, exhibit high photostability and are less sensitive to pH changes. Alexa dyes that absorb above 480 nm have the highest extinction coefficient comparable to those of fluorescein and rhodamine.[5]

However, sulfonation makes Alexa dyes hydrophilic and negatively charged which may lead to nonspecific electrostatic interactions with positively charged structures. In addition these dyes are also more expensive than the conventional ones.

Figure 1.5 – Plain and ball-and-stick structures of Alexa Fluor 350. FCUP 37 An overview of nanoscale fluorescent materials

Figure 1.6 – Plain and ball-and-stick structures of Alexa Fluor 430.

1.2. Fluorescent Proteins

Fluorescent proteins such as green fluorescent protein (GFP) are an endogenous agent that use an enzyme mediated process inside the body to generate visible light when a substrate is degraded.[12] GFP (see Figure 1.7) was originally isolated from jellyfish Aequora Victoria[13] and is composed of 238 amino acids residues. It exhibits bright green fluorescence under light excitation from blue to ultraviolet.[5] This protein is widely used in biochemistry and cell biology and has become an established marker for gene expression and protein targeting.[14] GFP’s excitation wavelength is 490 nm and its emission wavelength is 510 nm, which is a drawback because it overlaps with the autofluorescence of many tissues.[12] Furthermore there is also a problem of potential aggregation of the fluorescent proteins that can lead to quenching. Moreover and similar to organic dyes, fluorescent proteins suffer from photobleaching. Fluorescent proteins also have short time blinking meaning that they can’t undergo repeated cycles of fluorescent emission.[3, 5] 38 FCUP An overview of nanoscale fluorescent materials

Figure 1.7 - Structure of the Aequorea victoria green fluorescent protein. (Source: Ormö et al.[15])

1.3. Semiconductor quantum dots

Semiconductor nanocrystals, also known as Quantum Dots (QDs), are a novel inorganic fluorophores class which have become popular in the past two decades due to their exceptional photophysical properties. These semiconductors nanocrystals have sizes in the range of 1-10 nm and their fluorescence is due to quantum confinement effects. Also due to quantum confinement the absorption and emission wavelengths of QDs is size dependent, meaning that they can be synthesized in a wide range of sizes with the wavelength of light emitted related to the size of the QDs (Figure 1.8). QDs are composed of combined elements from periodic groups II–VI (e.g. ZnS, CdSe, CdTe), III–V (e.g. InP, GaAs, InAs), or IV–VI (e.g. PbS, PbSe, PbTe) and they provide a new class of biomarkers that can overcome the usual limitations of organic dyes.[16] QDs exhibit high photostability, broad absorption, narrow and symmetric emission spectra, slow excited decay rates and large absorption cross-sections. Their broad absorption and narrow emission spectra allows a single laser to excite QDs of a wide size-range, with each dot emitting its own specific colour, contrasting to organic-based fluorophores, which are characterized by narrow Stokes shifts.[17] Also when compared with traditional organic fluorophores they present unique fluorescence properties.[18] Unlike organic fluorophores which bleach after only a few minutes on exposure to light, QDs are extremely stable and can undergo repeated cycles of excitation and fluorescence for hours with a level of brightness and photobleaching threshold.[12, 19] QDs are used in biological applications such as cellular labelling, cell tracking and tissue imaging.[20] However, QDs themselves are hydrophobic and non-biocompatible, requiring layers of polymeric or inorganic material to make them compatible for biological applications.[3] Additionally some QDs contain toxic components, such as FCUP 39 An overview of nanoscale fluorescent materials

cadmium or lead (from cadmium and lead-based QDs respectively) which can result in the release of Cd2+ and Pb2+, toxic ions, and therefore in cell death in biological media. To overcome this issue surface modification of QDs can be employed.

Figure 1.8 - Photograph and spectra of CdSe quantum dots. The samples represent different sizes of QDs, which produce different colours upon UV light. An increase in particle size produces a red shift in the emission spectra. (Source: Nauman et al. [17])

Among the methods for surface modification are the conjugation of QDs with mercaptoacetic acid and silica coating.[5, 12] However, QDs capped with small molecules as the case of mercaptoacetic acid can be easily degraded by hydrolysis or oxidation of the capping agent. On the other hand silica coating can improve QDs stability without interfering with their optical properties. Also capping QDs with silica makes it possible to encapsulate several QDs in one nanoparticle enhanced their optical signal. Although due to the hydrophobic nature of QDs it is difficult to encapsulate them directly in silica without making them hydrophilic in the first place.[5] Overall, the toxicity of the elements that compose the QDs together with their hydrophobic nature makes their use difficult for in vivo applications. 40 FCUP An overview of nanoscale fluorescent materials

1.4. Dyed polymer nanoparticles

Dyed polymer nanoparticles are an interesting group of fluorescent probes that present a large and diverse group of sensors with one or more analytical reagents incorporated into the polymeric matrix.[16] Typical polymer-based dye-doped nanoparticles are made of hydrophobic polymers such as styrene, olefin, vinylpyridine and vinylpyrrolidone polymers.[17] In these NPs the dye molecules are incorporated inside the polymer matrix through covalent attachment of the dye molecules to the polymer chain or by physical entrapment in a cross-linked particle.[3, 5] Contrary to single organic fluorophores each polymer-based dye-doped nanoparticle can incorporate several molecules of the fluorophore embedded in the polymer shell and protected from the outer environment. As a result each nanoparticle is brighter than the single fluorophores due to the large number of dye molecules per particle and since the fluorophores are protected from the external environment they are more stable to photobleaching.[5] Although these nanoparticles are reasonably photostable, their application in bioimaging is limited due to the hydrophobic nature of these probes. Phenomena like clustering and non-specific binding can occur under imaging applications in aqueous environments. To overcome these problems, the surface of these nanoparticles can be functionalized with hydrophilic coatings like polymers such as poly(ethylene glycol) (PEG).[17] Polymer NPs have found so far frequent applications in intracellular sensing and cell targeting. However, these NPs usually exhibit low incorporation rates and little protection to the dyes.[3, 21] Apart from that the post coating step can also bring other problems like larger particles size, swelling, particle aggregation and leaking of the fluorophores through surface defects.[5, 19]

1.5. Fluorophore doped silica nanoparticles

Fluorophore doped silica nanoparticles have been developed for ultrasensitive bioanalysis and diagnosis over the past several years.[2] These fluorescent NPs consist of a fluorophore dispersed within a silica matrix and contrary to dyed polymer NPs the hydrophilic nature of the dye-doped silica NPs reduces the problems with non-specific binding and clustering.[17] Typically traditional organic dyes are encapsulated but even inorganic fluorophores such as lanthanides can be used. By simply changing the fluorophore a wide range of fluorescence wavelengths, either visible or near infrared, can be obtained.[22, 23] Encapsulation of the fluorescent molecules within the silica framework can overcome some of the functional limitations of the bare molecules. FCUP 41 An overview of nanoscale fluorescent materials

When the fluorophores are encapsulated into a silica matrix the resulting NPs are brighter than single molecules, since they can incorporate more than one molecule per particle. As a matrix material for fluorescent probes, silica provides a chemically and mechanically stable shell, which can protect and stabilize the encapsulated fluorophores and enhance their photophysical properties such as fluorescence and brightness.[3]

The application of silica NPs in biological samples presents a large number of advantages. Silica NPs are robust, mechanically stable and transparent, are easy to prepare and exhibit good monodispersity. Additionally, their surfaces can be easily functionalized for further

conjugation with antibodies, peptides, Figure 1.9 - Schematic illustration of the surface DNA, etc. (Figure 1.9). Moreover pH functionalization of silica NPs with, for example, peptides, antibodies, aptamers, enzymes, DNA- changes do not lead to swelling and fragments and different functional moieties. (Source: porosity changes, and silica particles Schulz et al.[21]) are not prone to microbial attack.[17]

These fluorescent silica based NPs can be synthesized in a wide size range from 10 nm to several hundreds of nanometers meaning that the size can be adjusted for a specific application. Furthermore in most of biological studies these fluorescent NPs were found to be biocompatible and haven´t show significant toxic effects.[21]

In chapters 2 and 3 the preparation, chemical composition and uses of this class of fluorescent NPs will be discussed in more detail.

42 FCUP An overview of nanoscale fluorescent materials

1.6. References

1. Buzea, C., Pacheco, I.I. and Robbie, K., Nanomaterials and nanoparticles: Sources and toxicity. Biointerphases, 2007. 2(4): p. Mr17-Mr71. 2. Yao, G., Wang, L., Wu, Y., Smith, J., Xu, J., Zhao, W., Lee, E. and Tan, W., FloDots: luminescent nanoparticles. Analytical and Bioanalytical Chemistry, 2006. 385(3): p. 518-24. 3. Burns, A., Ow, H. and Wiesner, U., Fluorescent core-shell silica nanoparticles: towards "Lab on a Particle" architectures for nanobiotechnology. Chemical Society Reviews, 2006. 35(11): p. 1028-1042. 4. Lakowicz, J.R., Principles of Fluorescence Spectroscopy. 3rd ed. 1999, New York: Kluwer Academic. 5. Wang, F., Tan, W.B., Zhang, Y., Fan, X.P. and Wang, M.Q., Luminescent nanomaterials for biological labelling. Nanotechnology, 2006. 17(1): p. R1-R13. 6. Holmes, K.L. and Lantz, L.M., Protein labeling with fluorescent probes. Methods in Cell Biology, 2001. 63: p. 185-204. 7. Lide, D.R. and Milne, G.W.A., Handbook of Data on Organic Compounds: Compounds 1001-15600 1994, Boca Raton: CRC Press. 8. Egawa, Y., Hayashida, R., Seki, T. and Anzai, J., Fluorometric determination of heparin based on self-quenching of fluorescein-labeled protamine. Talanta, 2008. 76(4): p. 736-41. 9. Lakowicz, J.R., Malicka, J., D'Auria, S. and Gryczynski, I., Release of the self- quenching of fluorescence near silver metallic surfaces. Analytical Biochemistry, 2003. 320(1): p. 13-20. 10. Escobedo, J.O., Rusin, O., Lim, S. and Strongin, R.M., NIR dyes for bioimaging applications. Current Opinion in Chemical Biology, 2010. 14(1): p. 64-70. 11. Sameiro, M. and Goncalves, T., Fluorescent Labeling of Biomolecules with Organic Probes. Chemical Reviews, 2009. 109(1): p. 190-212. 12. Sharma, P., Brown, S., Walter, G., Santra, S. and Moudgil, B., Nanoparticles for bioimaging. Advances in Colloid and Interface Science, 2006. 123-126: p. 471- 85. 13. Shimomura, O., Johnson, F.H. and Saiga, Y., Extraction, purification and properties of aequorin, a bioluminescent protein from the luminous hydromedusan, Aequorea. Journal of Cellular and Comparative Physiology, 1962. 59: p. 223-39. FCUP 43 An overview of nanoscale fluorescent materials

14. Tsien, R.Y., The green fluorescent protein. Annual Review of Biochemistry, 1998. 67: p. 509-544. 15. Ormo, M., Cubitt, A.B., Kallio, K., Gross, L.A., Tsien, R.Y. and Remington, S.J., Crystal structure of the Aequorea victoria green fluorescent protein. Science, 1996. 273(5280): p. 1392-5. 16. Ruedas-Rama, M.J., Walters, J.D., Orte, A. and Hall, E.A.H., Fluorescent nanoparticles for intracellular sensing: A review. Analytica Chimica Acta, 2012. 751: p. 1-23. 17. Naumann, M.J.M.a.C.A., ed. Biofunctionalization of Nanomaterials. Nanotechnologies for the Life Sciences ed. Kumar, C.S.S.R. Vol. 1. 2005, WILEY-VCH Verlag GmbH & Co. KGaA: Weinheim. p.1-39. 18. Michalet, X., Pinaud, F., Lacoste, T.D., Dahan, M., Bruchez, M.P., Alivisatos, A.P. and Weiss, S., Properties of fluorescent semiconductor nanocrystals and their application to biological labeling. Single Molecules, 2001. 2(4): p. 261-276. 19. Santra, S., Zhang, P., Wang, K., Tapec, R. and Tan, W., Conjugation of biomolecules with luminophore-doped silica nanoparticles for photostable biomarkers. Analytical Chemistry, 2001. 73(20): p. 4988-93. 20. Medintz, I.L., Uyeda, H.T., Goldman, E.R. and Mattoussi, H., Quantum dot bioconjugates for imaging, labelling and sensing. Nature Materials, 2005. 4(6): p. 435-46. 21. Schulz, A. and McDonagh, C., Intracellular sensing and cell diagnostics using fluorescent silica nanoparticles. Soft Matter, 2012. 8(9): p. 2579-2585. 22. Herz, E., Burns, A., Bonner, D. and Wiesner, U., Large stokes-shift fluorescent silica nanoparticles with enhanced emission over free dye for single excitation multiplexing. Macromolecular Rapid Communications, 2009. 30(22): p. 1907- 10. 23. Xu, J., Liang, J., Li, J. and Yang, W., Multicolor dye-doped silica nanoparticles independent of FRET. Langmuir, 2010. 26(20): p. 15722-5.

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FCUP 45

2. Fluorescent silica nanoparticles doped with organic dyes

Fluorescent dye-doped silica NPs have been extensively used for a wide range of applications in biological detection and diagnosis in the past several years. Dye doped- silica NPs are extremely bright and photostable because a large number of fluorescent dye molecules are encapsulated in the silica matrix that serves to protect the dye from photodegradation. This also enables NPs to exhibit strong emission signal when properly excited which can dramatically lower the analyte detection limit in biological samples.

Using appropriate synthetic conditions, a large variety of either organic or inorganic dye molecules can be incorporated inside a single silica particle. Despite the fact that incorporating a large amount of dye molecules into a single NP would be expected to lead to some fluorescence quenching phenomena due to the particle small volume, the goal of obtaining a particle with brighter luminescence is largely successful.[1]

Photostability is an important criterion in observation of fluorescence signal, especially under intense excitation and one of the major concerns when using bare dyes in bioanalysis. However, when dye molecules are encapsulated into a silica matrix which provides an effective barrier from the surrounding environment, the doped dye molecules are protected from oxygen and both photobleaching and photodegradation phenomena that often affect conventional dyes can be minimized. Moreover, the encapsulation enables the fluorescence to be constant providing an accurate measurement making these NPs suitable for applications where high intensity or intense excitations are needed.[1, 2] In addition, when dye doped silica NPs are used in real biological media the dye molecules are protect against degradation by the complex biological environment due to high resistance to chemical and metabolic degradation of the silica matrix.[3]

Silica is a good matrix material due to its flexible chemistry which allows surface modification with different types of functional groups. This property makes silica a versatile and biocompatible substrate for biomolecule immobilization. Biochemically modified dye-doped silica NPs can be used to express the activity of a desired process, such as enzyme immobilization, or can be used as an affinity ligand to capture or 46 FCUP Fluorescent silica nanoparticles doped with organic dyes

modify target molecules or cells.[3] In addition the silica surface makes these NPs chemically inert and physically stable.[4] Furthermore, studies on the cytotoxicity of these NPs have showed the benign nature of silica based NPs, exhibiting low or no cytotoxicity.[3]

Owing to these properties the use of dye-doped silica NPs as labelling reagents for bioanalysis and bioimaging have been widely studied and used.[1, 3] For example these NPs have been used to create assays for oligonucleotides, proteins and antibodies[5], cell targeting[3] and intracellular sensors[6].

2.1. Nanoparticle formation

Commonly, silica NPs are formed as a result of the base-catalysed hydrolysis of an organic precursor like tetraethylortosilicate (TEOS) and subsequent condensation to a silica network as shown in the equations (1) and (2).[6]

(1)

(2)

Silica NPs are generally synthesized by one of two chemical routes. These are sol- gel synthesis also known as the Stöber method[7] and the reverse microemulsion approach[8]. In the Stöber method, TEOS is hydrolysed in a dilute ethanolic solution and in the microemulsion method, TEOS is hydrolysed inside the water droplets of a water-in-oil microemulsion.[6] Both methods offer the possibility to incorporate fluorophore molecules inside the silica matrix. In the case of dye-doped silica NPs, the dye encapsulation is achieved by either covalent attachment of the dye with silica precursors (e.g. 3-(aminopropyl)triethoxysilane, APTES), before the hydrolysis in Stöber’s method, or by first solubilizing the dye in the microemulsion reaction media and then carrying out the polymerization.[9]

2.1.1. Stöber method

The Stöber method is a physical chemistry process for the generation of silica particles that was discovered in 1968 by Werner Stöber and Arthur Fink.[7] This method has been used to prepare spherical silica nanoparticles with different sizes. Silica NPs are formed by the addition of TEOS to an excess of water containing ammonia and a low molar-mass alcohol such as ethanol. The resulting silica particles have diameters FCUP 47 Fluorescent silica nanoparticles doped with organic dyes

between 50 and 2000 nm depending on type of organic precursor and alcohol used and on water/ethanol volume ratios.[10] In general a lower concentration of water leads to particles of smaller size. The Stöber method is a relatively simple procedure to make silica NPs and can be carried out in only few hours. Although Stöber’s method presents the advantage of having a reaction that can be scaled up easily to yield large amounts of nanoparticles, it can also lead to particles with non-uniform sizes (see Figure 2.1).[9]

Figure 2.1 - TEM images: (A) silica-based nanoparticles prepared by the Stöber method; and (B) silica nanoparticles prepared by the microemulsion process.

2.1.2. Microemulsion method

The reverse micelle system, also known as the water-in-oil (W/O) microemulsion system is an alternative method to form silica NPs. A reverse microemulsion is an isotropic and thermodynamically stable single-phase system consisting of water, oil and surfactant.[2] Figure 2.2 shows a typical structure of a reverse micelle where the nanodroplets of water are surrounded by surfactant and dispersed in the continuous bulk oil phase. The water nanodroplets serve as nanoreactors for the synthesis of the nanoparticles whose size is dependent on the size of those nanodroplets, which is [2] controlled by the water-to-surfactant molar ratio (W0). 48 FCUP Fluorescent silica nanoparticles doped with organic dyes

Figure 2.2 - Typical structure of a reverse micelle (source: Malik et al.[11])

In addition, during the later stages of growth, steric stabilization provided by the surfactant layer prevents the nanoparticles from aggregating.[11] Similar to Stöber’s method the particles synthesized by microemulsion technique are formed by the hydrolysis and polymerization of silane precursors in presence of ammonia. The reverse microemulsion method produces fairly uniform and monodisperse nanoparticles (Figure 2.1 B), but takes 24 to 48 hours to complete. The advantage of the reverse microemulsion method apart from the fact that it produces highly spherical and monodisperse nanoparticles of various sizes is the ability to encapsulate a wide variety of organic and inorganic fluorophores as well other materials such as luminescent quantum dots. In case of the last ones previously to encapsulation QDs have to undergo a ligand exchange process in order to make them hydrophilic.

2.1.3. Incorporation of organic fluorophores

The Stöber and reverse microemulsion methods offer the possibility to incorporate fluorophore molecules inside the silica matrix either by physical entrapment or by covalent binding. The physical entrapment of dye molecules is usually obtained by adding the fluorophore to the reaction media however this incorporation method leads very often to dye leakage from the NPs. Covalent binding of the dye to the silica matrix is obtained by reaction of the dye to a silane agent such as APTES before the hydrolysis and condensation of TEOS. This approach reduces dye leakage from the silica matrix and also enables the incorporation of a variety of organic dye molecules into the silica NPs.[3]

The first report on the covalent incorporation of organic fluorophores into colloidal silica NPs was made in 1992 by Van Blaaderen and co-workers.[12] They report the FCUP 49 Fluorescent silica nanoparticles doped with organic dyes

successful covalent linkage of fluorescein isothiocyanate (FTIC) to APTES and subsequent incorporation of the dye-silane agent into the silica matrix. Since then different kinds of fluorescent organic dyes such as methylene blue (MB)[13], rhodamine 6G (R6G)[14], tetramethylrhodamine isothiocyanate (TRITC)[15] or rhodamine b isothiocyanate (RBITC)[16] have been incorporated into the matrix of silica NPs by covalent attachment.

Besides single-dye doping, multiple-dye incorporation into the silica matrix is also [17] possible. Recently Wang et al. reported the simultaneously incorporation of three organic dyes, FTIC, R6G and 6-carboxyl-X-rhodamine (ROX) into the same silica matrix for multiplexed signalling in bioanalysis. These NPs uses fluorescence resonance energy transfer (FRET) as the emission scheme. By controlling the doping ratios of the dyes the FRET-mediated emission signals can be tuned and the NPs exhibit different colours under the same single wavelength excitation. This allows simultaneous detection of multiple FRET targets.

2.2. Nucleation and growth of silica NPs

Formation of silica nanoparticles occurs in three stages: silica polymerization and nucleation of silica nanospheres, followed by particle growth and/or ripening and particle aggregation.[18] In the initial stage, silica monomers polymerize by condensation of dimers and trimers to colloids which then form the nanoparticles. During the second stage these particles grows by further addition of silica monomers, trimers or larger colloids and/or by Ostwald ripening. The final stage often occurs when particles stick to each other, and spontaneously form irregular particle clusters or aggregates.

2.2.1. Nucleation mechanisms

Nucleation of a new phase can occur when the overall free energy of the system is at its lowest. Nucleation can be heterogeneous (when initiated at nucleation sites such as phase boundaries, surfaces or impurities like dust) or homogeneous (when occurs randomly and spontaneously without the use of surfaces).

Homogeneous nucleation is generally more difficult to occur since the creation of a nucleus implies the formation of an interface at the boundaries of a new phase. For homogeneous nucleation to occur the solution needs to be supersaturated with respect to the new forming phase.[19] 50 FCUP Fluorescent silica nanoparticles doped with organic dyes

On the other hand heterogeneous nucleation occurs more often than homogeneous nucleation. Since heterogeneous nucleation takes place at preferential sites, it requires less energy than homogeneous nucleation because the effective surface energy at those sites is lower, thus diminishing the free energy barrier and this facilitates the nucleation. In this type of nucleation, some energy is released by the partial destruction of the previous surface allowing the new phase to form without the need for supersaturation.[19]

2.2.2. Growth mechanisms

In classical growth theory it is assumed that particle growth occurs by molecule-by- molecule attachment to a pre-existing surface.[18] Based on this theory, the molecules diffuse onto the particle surface where it will attach itself to a suitable growth site. Alternatively to the classical growth is the growth model based on Ostwald ripening[20], which consists of a mass transfer process where smaller particles in solution dissolve and deposit on larger particles in order to reach a more thermodynamically stable state. Thus small particles decrease in size until they disappear and large particles grow even larger. This shrinking and growing of particles will result in an increase in mean particle size. Ostwald ripening is often found in water-in-oil emulsions where oil molecules will diffuse through the aqueous phase and join larger oil droplets.

2.2.3. Particle growth in the reverse micellar system

Particle growth in the reverse microemulsion system is illustrated in Figure 2.3. This mechanism can be viewed as a fluid phase consisting of reverse micelles that are filled with silica particles or empty (free of particles but containing hydrolysed TEOS molecules). Additionally there is a fraction of TEOS molecules that remain in the oil phase during the reaction.[21] Particle growth can then result from the transfer of the hydrolysed TEOS in the reverse micelles to the micelles filled with silica nanoparticles. Direct interaction of TEOS that remain in the oil phase (non-hydrolysed TEOS) with the particle filled micelles can also occur. Prior to growth, hydrolysis of TEOS has to proceed in the water-shell (or hydration layer) that surrounds the particles.[21] FCUP 51 Fluorescent silica nanoparticles doped with organic dyes

Figure 2.3 – Silica nanoparticles growth mechanism in a reverse micellar system composed. (Source: Osseo-Asare et al.[21])

2.3. Surface functionalization of dye-doped silica NPs

Controlling the surface chemical composition of the nanoparticles can confer them with stability, biocompatibility and enables their use in a wide range of bioapplications.[22] Due to the versatility of silica chemistry it is possible to modify the particle’s surface with various functional groups for biological applications (Figure 2.4). A variety of methods are available for particle surface modification, among them are the physical absorption and the chemical binding.

Physical absorption relies on the formation of noncovalent interactions and is commonly employed to modify the silica NPs surface with avidin. Avidin is a glycoprotein with an overall positive charge, which can attach to the negatively charged silica surface through electrostatic interactions (see Figure 2.5 bottom).[1, 14]

52 FCUP Fluorescent silica nanoparticles doped with organic dyes

Figure 2.4 - Schematic illustration of the surface functionalization of silica NPs for biological applications. (Source: Smith et al.[4])

The chemical binding takes advantage of the condensation of alkoxy groups of organosilanes with silanol groups on the particle surface. The particle surface is first

modified with functional groups such as, thiol (-SH), amine (-NH2) and carboxyl (COOH) groups through an additional silica coating (post-coating) that contains the functional groups of interest. Afterwards biomolecules such as proteins, antibodies, oligonucleotides, etc. can be conjugated to silica nanoparticles through interaction with those functional groups following standard conjugation methods. These binding methods are listed in Table 2.1 and schematically represented in Figure 2.5. For example, thiol functionalized NPs can conjugate with dissulfide modified oligonucleotides by a dissulfide coupling chemistry while amine modified NPs can be coupled to a wide variety of haptens (small molecules that react with a specific antibody) via succinimidyl esters and isothiocyanates.[3] The carboxyl modified NPs are suitable for covalent coupling with proteins or other amine containing biomolecules trough carbodiimide chemistry.[3] In case of Stöber nanoparticles, the surface modification is usually achieved after nanoparticle synthesis to avoid potential secondary nucleation. Surface modification of microemulsion NPs in the other hand can be done in the same manner or via direct hydrolysis and co-condensation of TEOS and other organosilanes in the microemulsion reaction media.[3, 23] After the bioconjugation step, the nanoparticles can be separated from unbound biomolecules by centrifugation, dialysis, filtration, or other laboratory techniques.

53 FCUP Fluorescent silica nanoparticles doped with organic dyes

Table 2.1 - Chemical binding for bioconjugation of silica NPs. (adapted from Yao et al. [2])

Organosilanes used on surface Functional Target Structure Bioconjugation method modification group on NPs biomolecules

OCH3 Dissulfide modified HS 3-mercaptopropyltrimethoxysilane oligonucleotides Si OCH3 -SH Thiol-dissulfide exchange (MPTS) OCH3 (-S-S-)

H3C O Antibodies (3-aminopropyl)-triethoxysilane O Si NH2 -NH2 Amine-thiocyanate coupling (APTES) O CH3 (-NCS)

CH3

Proteins or other O OH amine containing Carboxyethylsilanetriol OH + -COOH biomolecules Carbodiimide chemistry (CTES) NaO Si

OH (-NH2) 54 FCUP Fluorescent silica nanoparticles doped with organic dyes

Figure 2.5 - Representative bioconjugation schemes for attaching biomolecules to dye-doped silica NPs for bioanalysis. (source: Wang et al. [3])

2.4. Nanoparticle characterization

Characterization of NPs is important to elucidate the structure, characteristics and mechanism of nanoparticle formation. Evaluation of the nanoparticles in relation to their photostability, surface properties, size and morphology provides information that can be used to improve and enhance the synthesis protocol. Typical particle characterization methods include particle size and shape measurements, determination of surface charge and functionality, and determination of the optical and spectral characteristics. Chemical characterization is also important to quantify the amounts of doped dye molecules and surface-immobilized biomolecules.[3]

2.4.1. Measure particle size

Several techniques are currently available for measuring particle size including transmission electron microscopy (TEM), scanning electron microscopy (SEM), atomic force microscopy (AFM) and dynamic light scattering (DLS). TEM and SEM are commonly used for size characterization of nanoparticles in vacuum, while AFM is used for both dry and wet samples at normal atmospheric pressure. DLS allows particle size measurements in aqueous media giving information on nanoparticle size distribution and relative dispersion.[3] DLS determines the hydrodynamic diameter of the NPs meaning that it measures the Brownian movement of the various particles FCUP 55 Fluorescent silica nanoparticles doped with organic dyes

present in a given solution or dispersion. Therefore the diameter depends not only on the particle size by itself but also in the solvation sphere around the NPs.

Particle size is affected by the amount of TEOS and ammonia both in microemulsion and Stöber methods. In the reverse microemulsion method, the particle size is also affected by the nature of the surfactant (anionic, cationic or nonionic according to the load of its hydrophilic part) and by water to surfactant molar ratio.

NPs prepared by the reverse microemulsion method are spherical with smooth surfaces and low polydispersity while NPs prepared by the Stöber method are less monodisperse, less spherical and less smooth.[3]

2.4.2. Surface charge

The zeta potential ζ of a particle is a measure of the overall charge carried by the particle in a particular medium. The value of the measured ζ is indicative of the repulsive forces that are present and can be used to predict the long-term colloidal stability of the particles. If all the particles in suspension have a high negative or positive ζ, they repel each other, and have no tendency to agglomerate.[3, 4] Based on this, in the case of silica NPs, due to the strong negative charge on the particles surface, provided by silanol groups (Si–OH → Si–O-), suspensions should tend to be well dispersed and without aggregation. However in biological media due to pH and high salt concentrations it is possible that particles aggregate. Based on ζ measurements it is also possible to determine whether bioconjugation reactions occurred if the reacting species are charged (e.g. proteins and DNA).[3, 4]

2.5. Biological applications of dye-doped silica NPs

Dye-doped silica NPs have been extensively used for a wide range of applications in biological detection and diagnosis due to their important properties such as high optical intensity and photostability and easy bioconjugation. The first biological applications of these particles were used for the targeting of cancer cells [24] however the encapsulation of dyes into silica NPs as also been carried out for development of intracellular nanosensors[6], multiplexed analysis[1] and DNA analysis[1, 3].

56 FCUP Fluorescent silica nanoparticles doped with organic dyes

2.5.1. Cell targeting using dye-doped silica NPs

Different kinds of fluorescent dyes have been incorporated into silica NPs for targeting cells purpose. After NPs surface modification with the functional groups of interest, the fluorescence intensity of the NPs is used to determine localization of NPs in cellular media through fluorescence microscopy, flow cytometry or confocal microscopy. In 2008 Santra and coworkers[25] reported the incorporation of a metallorganic dye (Rubpy) inside silica matrix for leukemia cell recognition. Later on the same dye was incorporated into galactose-conjugated silica NPs to be used as an immunofluorescence assay for cell labelling and identification of liver cancer cells in blood.[26] The organic rhodamine dye and its derivatives have been also incorporated into silica NPs for cell targeting purposes. Ow and coworkers reported the synthesis of silica NPs doped with TRITC and their use as markers for biological imaging by labelling a cell surface receptor (FcϵRI) of rat basophilic leukemia mast cells.[27] In another work based on rhodamine b isothiocyanate (RBITC) doped silica NPs modified with Annexin V[28], was reported the successful application of these NPs as biomarkers for cell apoptosis. These NPs could specifically recognise early-stage apoptotic cells through the binding between Annexin V and a phospholipid component (phosphatidylserine) on the outer membrane of apoptotic cells.

2.5.2. Dye-doped silica NPs as intracellular nanosensors

Some of the first silica based nanosensors employed for the detection of small molecules in live cells measured intracellular dissolved oxygen.[24] However, several applications of dye-doped silica NPs as intracellular nanosensors for different analytes have been reported. The basis of detection is the change in NPs emission intensity in the presence of the analyte of interest. The earliest example on the use of these NPs for intracellular sensing was reported by Kopelman in 2001.[29] Here they reported the incorporation of two fluorophores: the oxygen-sensitive ruthenium complex (Ru(II)- 2+ tris(4,7-diphenyl-1,10-phenantroline) ([Ru(dpp)3] ) and the analyte-insensitive reference dye Oregon Green 488 dextran into silica NPs called PEBBLEs (probes 2+ encapsulated by biologically localized embedding). The ([Ru(dpp)3] complex exhibit good photostability and high quantum yield and its fluorescence is quenched in the presence of oxygen.

A similar ratiometric nanosensor was developed by Burns and coworkers[30] for pH sensing. These pH-sensitive silica NPs were developed by incorporation of a pH FCUP 57 Fluorescent silica nanoparticles doped with organic dyes

sensitive indicator fluorophore (FTIC that exists in several protonation states depending on pH) and a reference dye (TRITC) to the silica matrix during synthesis. NPs were used to determine pH changes in intracellular components of rat basophilic leukemia mast cells using confocal fluorescence microscopy (see Figure 2.6). Another example of a pH nanosensor based on dye-doped silica NPs is the work of Peng et al.[31] where they report the use of a nanoparticle based pH sensor for noninvasive monitoring of intracellular pH changes in living cells by drug stimulation. The pH sensor is also a two- fluorophore-doped nanoparticle sensor that contains FTIC as pH-sensitive indicator and Rubpy as a reference dye. The NPs measured pH changes inside murine macrophages during drug stimulation and in HeLa cancer cells during apoptosis. These studies demonstrate how dye-doped silica NPs can be used to monitor pH in certain cell compartments and to investigate cellular processes.

Figure 2.6 - Confocal fluorescence microscopy images (overlaid and bright field)of pH sensors in rat basophilic leukemia mast cells showing a) reference dye (RBITC) channel, b) sensor dye (FTIC) channel, c) overlaid images and d) false- colour ratiometric imaging of pH in various intracellular compartments (Source: Burns et al.[30])

2.5.3. Dye-doped silica NPs for multiplexed bioanalysis

The need to observe and target many biological events simultaneously has led to the development of multiplexed fluorescent tags. Compared to single-target detection methods, multiplexed assays reduce the time and cost per analysis, allow for 58 FCUP Fluorescent silica nanoparticles doped with organic dyes

simpler assay protocols, decrease the sample volume required, make comparison of samples feasible and measurements reproducible and reliable.[32] Wang and coworkers have adopted a two-dye encapsulated NPs system for multiplexed bacteria detection with flow cytometry system.[3, 32] In this system three different antibodies were conjugated to the NPs doped with the two dyes. Each labeled nanoparticle specifically recognizes and binds to the corresponding antigen-presenting bacteria. When the bacteria-nanoparticle mixture passes through a flow cytometer each kind of bacterium- nanoparticle complex exhibit the unique fluorescence signature of the attached NPs.[1] This scheme enables very rapid, highly selective, and sensitive bioassays.

2.5.4. Dye-doped silica NPs for nucleic acid analysis

The intense fluorescence signal of one fluorescent silica nanoparticle can be effectively used in DNA hybridization analysis. In this application, the NPs are generally used to replace standard fluorescent dyes, in order to reveal the presence of bound DNA molecules on the microarray capture dots.[5] Zhao et al.[33] have developed an ultrasensitive DNA assay to detect gene products using a highly fluorescent and photostable bioconjugated dye-doped silica nanoparticle using TMR as the source of fluorescence. This test is based on a sandwich assay setup where three different DNA species are present: captured DNA which is immobilized on a glass surface; a probe sequence that is attached to the dye-doped silica NPs; and an unlabeled target sequence, which is complementary to both capture and probe sequences through different parts of the sequence. The capture DNA is first immobilized on the glass substrate and hybridizes with unlabeled target DNA, then probe DNA attached to the TMR-doped silica NPs is added for hybridization (Figure 2.7). The detection of target DNA is done by monitoring the fluorescence signals of NPs-probe DNA conjugates left on the glass substrates after subsequent washing steps. This sandwich assay proved to be successful in DNA analysis on sub femtomolar concentration limits.[33] This scheme has also been introduced into a protein microarray platform by Wang et al.[9] in order to obtain a system with an improved signal.

FCUP 59 Fluorescent silica nanoparticles doped with organic dyes

Figure 2.7 - Schematic representation of a sandwich assay based on dye-doped silica NPs. (Source: Zhao et al.[33])

Another work based on dye-doped silica NPs for microarray analysis is the one of Zhou[34] and coworkers where they report the use of silica core-shell NPs encapsulating cyanine dyes as labeling in DNA microarray based bioanalysis. These NPs were prepared by attaching dye-alkanethiol (dT)20 oligomers chemisorbed to the surface of colloidal gold (Au) particles. Then Au NPs were coated with a silica layer through thiol functional groups. Both Cy3-and Cy5-doped Au/silica core-shell particles were prepared and applied to two-colour microarray detection in a sandwich assay format. This system exhibited sensitivity ten times higher than the bare cyanine dyes with a detection limit of 1 pM for target DNA in a sandwich hybridization.[34]

Dye-doped silica NPs can be developed by either Stöber or reverse microemulsion methodologies, however the microemulsion systems yields more uniform and monodisperse NPs. The silica surface can be easily modified with various functional groups enabling its conjugation with a variety of biomolecules. Due to the flexible conjugation, excellent photostability and ultrasensitivity of dye-doped silica NPs these materials can be a powerful tool in bioanalysis.

60 FCUP Fluorescent silica nanoparticles doped with organic dyes

2.6. References

1. Yan, J.L., Estevez, M.C., Smith, J.E., Wang, K.M., He, X.X., Wang, L. and Tan, W.H., Dye-doped nanoparticles for bioanalysis. Nano Today, 2007. 2(3): p. 44- 50. 2. Yao, G., Wang, L., Wu, Y., Smith, J., Xu, J., Zhao, W., Lee, E. and Tan, W., FloDots: luminescent nanoparticles. Analytical and Bioanalytical Chemistry, 2006. 385(3): p. 518-24. 3. Wang, L., Wang, K.M., Santra, S., Zhao, X.J., Hilliard, L.R., Smith, J.E., Wu, J.R. and Tan, W.H., Watching silica nanoparticles glow in the biological world. Analytical Chemistry, 2006. 78(3): p. 646-654. 4. Smith, J.E., Wang, L. and Tan, W.T., Bioconjugated silica-coated nanoparticles for bioseparation and bioanalysis. Trac-Trends in Analytical Chemistry, 2006. 25(9): p. 848-855. 5. Baptista, P.V., Doria, G., Quaresma, P., Cavadas, M., Neves, C.S., Gomes, I., Eaton, P., Pereira, E. and Franco, R., Nanoparticles in Molecular Diagnostics, in Progress in Molecular Biology and Translational Science; Nanoparticles in Translational Science and Medicine, Villaverde, A., Editor. 2011, Elsivier: London. p. 427-488. 6. Schulz, A. and McDonagh, C., Intracellular sensing and cell diagnostics using fluorescent silica nanoparticles. Soft Matter, 2012. 8(9): p. 2579-2585. 7. Stober, W., Fink, A. and Bohn, E., Controlled Growth of Monodisperse Silica Spheres in Micron Size Range. Journal of Colloid and Interface Science, 1968. 26(1): p. 62-&. 8. Arriagada, F.J. and Osseoasare, K., Synthesis of Nanosize Silica in Aerosol Ot Reverse Microemulsions. Journal of Colloid and Interface Science, 1995. 170(1): p. 8-17. 9. Sharma, P., Brown, S., Walter, G., Santra, S. and Moudgil, B., Nanoparticles for bioimaging. Advances in Colloid and Interface Science, 2006. 123-126: p. 471- 85. 10. Bogush, G.H., Tracy, M.A. and Zukoski, C.F., Preparation of Monodisperse Silica Particles - Control of Size and Mass Fraction. Journal of Non-Crystalline Solids, 1988. 104(1): p. 95-106. 11. Malik, M.A., Wani, M.Y. and Hashim, M.A., Microemulsion method: A novel route to synthesize organic and inorganic nanomaterials. Arabian Journal of Chemistry, 2012. 5(4): p. 397-417. FCUP 61 Fluorescent silica nanoparticles doped with organic dyes

12. Vanblaaderen, A. and Vrij, A., Synthesis and Characterization of Colloidal Dispersions of Fluorescent, Monodisperse Silica Spheres. Langmuir, 1992. 8(12): p. 2921-2931. 13. Deng, T., Li, J.S., Jiang, J.H., Shen, G.L. and Yu, R.Q., Preparation of near-IR fluorescent nanoparticles for fluorescence-anisotropy-based immunoagglutination assay in whole blood. Advanced Functional Materials, 2006. 16(16): p. 2147-2155. 14. Tapec, R., Zhao, X.J.J. and Tan, W.H., Development of organic dye-doped silica nanoparticles for bioanalysis and biosensors. Journal of Nanoscience and Nanotechnology, 2002. 2(3-4): p. 405-409. 15. Larson, D.R., Ow, H., Vishwasrao, H.D., Heikal, A.A., Wiesner, U. and Webb, W.W., Silica nanoparticle architecture determines radiative properties of encapsulated fluorophores. Chemistry of Materials, 2008. 20(8): p. 2677-2684. 16. Verhaegh, N.A.M. and Vanblaaderen, A., Dispersions of Rhodamine-Labeled Silica Spheres - Synthesis, Characterization, and Fluorescence Confocal Scanning Laser Microscopy. Langmuir, 1994. 10(5): p. 1427-1438. 17. Wang, L. and Tan, W.H., Multicolor FRET silica nanoparticles by single wavelength excitation. Nano Letters, 2006. 6(1): p. 84-88. 18. Tobler, D.J., Shaw, S. and Benning, L.G., Quantification of initial steps of nucleation and growth of silica nanoparticles: An in-situ SAXS and DLS study. Geochimica et Cosmochimica Acta, 2009. 73: p. 5377-5393. 19. Tobler, D.J., Molecular pathways of silica nanoparticle formation and biosilicification, in School of Earth and Environment. 2008, University of Leeds: Leeds. p. 1-258. 20. Ostwald, W., Analytische Chemie, 3rd edition. 1901, Englemann. 21. Osseo-Asare, K. and Arriagada, F.J., Growth kinetics of nanosize silica in a nonionic water-in-oil microemulsion: A reverse micellar pseudophase reaction model. Journal of Colloid and Interface Science, 1999. 218(1): p. 68-76. 22. Jiang, S., Win, K.Y., Liu, S.H., Teng, C.P., Zheng, Y.G. and Han, M.Y., Surface- functionalized nanoparticles for biosensing and imaging-guided therapeutics. Nanoscale, 2013. 5(8): p. 3127-3148. 23. Deng, G., Markowitz, M.A., Kust, P.R. and Gaber, B.P., Control of surface expression of functional groups on silica particles. Materials Science & Engineering C-Biomimetic and Supramolecular Systems, 2000. 11(2): p. 165- 172. 62 FCUP Fluorescent silica nanoparticles doped with organic dyes

24. Ruedas-Rama, M.J., Walters, J.D., Orte, A. and Hall, E.A.H., Fluorescent nanoparticles for intracellular sensing: A review. Analytica Chimica Acta, 2012. 751: p. 1-23. 25. Santra, S., Zhang, P., Wang, K.M., Tapec, R. and Tan, W.H., Conjugation of biomolecules with luminophore-doped silica nanoparticles for photostable biomarkers. Analytical Chemistry, 2001. 73(20): p. 4988-4993. 26. Peng, J., Wang, K., Tan, W., He, X., He, C., Wu, P. and Liu, F., Identification of live liver cancer cells in a mixed cell system using galactose-conjugated fluorescent nanoparticles. Talanta, 2007. 71(2): p. 833-40. 27. Ow, H., Larson, D.R., Srivastava, M., Baird, B.A., Webb, W.W. and Wiesner, U., Bright and stable core-shell fluorescent silica nanoparticles. Nano Letters, 2005. 5(1): p. 113-7. 28. Shi, H., He, X.X., Wang, K.M., Yuan, Y., Deng, K., Chen, J.Y. and Tan, W.H., Rhodamine B isothiocyanate doped silica-coated fluorescent nanoparticles (RBITC-DSFNPs)-based bioprobes conjugated to Annexin V for apoptosis detection and imaging. Nanomedicine-Nanotechnology Biology and Medicine, 2007. 3(4): p. 266-272. 29. Xu, H., Aylott, J.W., Kopelman, R., Miller, T.J. and Philbert, M.A., A real-time ratiometric method for the determination of molecular oxygen inside living cells using sol-gel-based spherical optical nanosensors with applications to rat C6 glioma. Analytical Chemistry, 2001. 73(17): p. 4124-4133. 30. Burns, A., Sengupta, P., Zedayko, T., Baird, B. and Wiesner, U., Core/Shell fluorescent silica nanoparticles for chemical sensing: towards single-particle laboratories. Small, 2006. 2(6): p. 723-6. 31. Peng, J.F., He, X.X., Wang, K.M., Tan, W.H., Wang, Y. and Liu, Y., Noninvasive monitoring of intracellular pH change induced by drug stimulation using silica nanoparticle sensors. Analytical and Bioanalytical Chemistry, 2007. 388(3): p. 645-654. 32. Wang, L., Yang, C. and Tan, W., Dual-luminophore-doped silica nanoparticles for multiplexed signaling. Nano Letters, 2005. 5(1): p. 37-43. 33. Zhao, X.J., Tapec-Dytioco, R. and Tan, W.H., Ultrasensitive DNA detection using highly fluorescent bioconjugated nanoparticles. Journal of the American Chemical Society, 2003. 125(38): p. 11474-11475. 34. Zhou, X.C. and Zhou, J.Z., Improving the signal sensitivity and photostability of DNA hybridizations on microarrays by using dye-doped core-shell silica nanoparticles. Analytical Chemistry, 2004. 76(18): p. 5302-5312. FCUP 63

3. Lanthanopolyoxometalates encapsulated into silica nanoparticles

Polyoxometalates (POMs) are a highly versatile and easily modified class of inorganic compounds. The diversity of structures as well as the high number of elements that can make up the structure of a polyoxometalate (POM) allows for a wide range of applications in fields such as catalysis, medicine, biology and more recently in nanotechnology.[1, 2] Lanthanide-containing polyoxometalates (LnPOMs) in particular, are readily obtained through the coordination of lanthanide ions to lacunary POMs, and exhibit interesting luminescent properties and other specific characteristics that result from the synergy between the properties of lanthanide ions and POM units.[3-5] LnPOMs have been applied in different areas such as catalysis,[6, 7] magnetism,[8, 9] luminescence[3, 10] and medicine.[11, 12]

However, the use of LnPOMs in biological applications is hindered by the possible toxicity of the lanthanides and by their interaction with biological media, which may lead to coordination of biological ligands, hydrolysis, and other reactions that may adversely affect the properties of LnPOMs. One strategy to avoid possible adverse interactions with biological molecules is the encapsulation of LnPOMs into an inert nanomaterial. The high stability, chemical inertness and optical transparency of silica makes it the ideal candidate for encapsulation while preserving the properties of the encapsulated material, in particularly the optical properties.[13] Moreover, the surface of silica nanoparticles can be easily functionalized enabling their application in the preparation of biosensors and cell labeling.[14, 15]

3.1. Polyoxometalates

3.1.1. Definition

Polyoxometalates (POMs) are anionic species consisted by polyhedral units of transition metal polyoxoanions (MOx, generally MO6 octahedrons – Figure 3.1), linked together by shared oxygen atoms to form a large and closed 3-dimensional framework. 64 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles

Figure 3.1 - Ball-and-stick (left) and polyhedral (right) representations of the fundamental unit MO6. (Source: Fernandez[16])

The linkage of MO6 units can be done by edge and corner-sharing of MO6 octahedrons. Less often this linkage can also be accomplished by face-sharing of the

MO6 octahedrons (Figure 3.2).

Figure 3.2 - Representation of the three possible unions between two MO6 octahedral units: A) corner-sharing, B) edge- sharing and C) face-sharing. Each corner represents an oxygen position. (Source: Fernandez[16])

p- POMs can be classified in two main classes, the isopolyanions ([MmOy] ) and the q- heteropolyanions ([XxMmOy] ). The isopolyanions are anions composed of a metal- oxide framework while the heteropolyanions apart from this framework also have an internal heteroatom X. The metal atoms that make up the framework, also called addenda atoms, are typically V, Nb, Ta, Mo and W in the V or VI oxidation states (electronic configuration d0 or d1).[17, 18] When more than one addenda atom is present in the framework the cluster is called a mixed addenda cluster. In general, any element can participate as X in a POM cluster since there are no strict physical requirements for this position, X can be a non-metal (e.g. P), a semi-metal (such as B and Si), a transition metal (e.g. Co and Fe) or a p-block metal (as Al).[19, 20] The heteroatom X,

when present, is the central atom and forms a central tetrahedron XO4. The heteroatoms can be classified as primary or secondary (also called peripherals). The primary heteroatoms are indispensable for the heteropolyanions basic structure since they cannot be removed without destroying the anion. In the other hand and since they FCUP 65 Lanthanopolyoxometalates encapsulated into silica nanoparticles

are not essential for the maintenance of the POM structure, the secondary heteroatoms can be removed from the heteropolyanion giving rise to other anionic stable species.[21]

In Figure 3.3 are represented some examples of the two different classes of POMs. Among them, the most studied and well-known structures are the Keggin-type and Wells-Dawson. In this work, were used the Keggin-type heteropolyanions and for this reason special attention will be given in section 3.2 to this type of structure.

n- Figure 3.3 - Polyhedral representation of common polyoxoanions: A) Lindqvist ([M6O19) ) isopolyanion; B) Anderson- n- n- n- n- Evans ([XM6O24] ); C) Keggin ([XM12O40] ); D) Wells-Dawson ([X2M18O62] ) and E) Preyssler ([XP5W30O110] ) heteropolyanions. (Source: Lopez et al. [22])

3.1.2. Historical context

In 1826 Berzelius[23] reported the discovery of the first POM, the phosphomolybdate, 3- of formula [PMo12O40] . Years later, in 1862, and after the discovery of the silicotungstic acid and its salts by Marignac[24] the analytical composition of these compounds began to be analysed. In 1929, Pauling[25] gave the first steps trying to 3- understand the structure of POMs, proposing that the anion [PMo12O40] discovered by

Berzelius had structure formed by a central tetrahedron XO4 surrounded by twelve MO6 octahedra sharing corners. However, in 1933 Keggin[26] solved, by X-ray diffraction, the structure of the phosphotungstic acid (H3PW12O40·5H2O) demonstrating that the anion n- [XM12O40] was composed by octahedral units, but contrary to that suggested by Pauling these units share between them not only corners but also edges.

Later on, in 1937 Anderson[27] suggested that the structure of the heteropolyanions n- 6- [XM6O24] and the isopolyanions [Mo7O24] was planar and formed exclusively by MO6 octahedra sharing edges between them. In the case of the heteropolyanions, this hypothesis was confirmed in 1948 when Evans[28] determined the structure of 6- [TeMo6O24] and this anion was then designated by Anderson-Evans. However, regarding the structure of the heteropolyanions the hypothesis suggested by Pauling was rejected in 1950 when Lindqvist[29] presented the correct structure for the 66 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles

6- heptamolybdate anion ([Mo7O24] ) proving that the geometry of the heteropolyanions was non-planar. Three years later, Dawson[30] reported the structure of another anion 6- the heteropolyanion of formula [P2W18O62] that confirmed the structure proposed by Wells[31] few years earlier. This anion known as Wells-Dawson consisted of a

diamagnetic anion with eighteen MO6 octahedra sharing edges and corners, being the two tetrahedral positions occupied by the heteroatoms. Since then, countless structures have been synthesised and characterised.

The turning point came when spectroscopic techniques (such as infrared, Raman and nuclear magnetic resonance – NMR), were used for the characterisation. Later on the use of single crystal X-Ray diffraction and the demand of new synthetic routes also allowed the growth of the POMs chemistry.[20] In the last decades, a lot of experimental information has been collected and today POMs constitute an immense class of polynuclear metal-oxygen clusters.[20]

3.1.3. Preparation

Heteropolyanions and isopolyanions are usually prepared and isolated from both aqueous and non-aqueous solutions. The most common method of synthesis involves m− dissolving [MOn] anions which, after acidification, assemble to yield a packed

molecular array of MO6 units, as indicated in the following equations (1) and (2):

(1)

(2)

Generally, pH conditions must be taken into account so that the reaction can be controlled. The sequence in which the reagents are added to the reaction media is sometimes also important. One of the latest steps in synthetic procedures, and maybe the most important if POMs are to be completely characterised, is the isolation of crystals so that their features can be studied in greater depth. Clusters are precipitated by adding countercations (alkali metals, organic cations like TBA, ammonia, etc.) and subsequent separation.[16]

POMs solubility depends on the cations that surround its structure. Generally POMs have a low solvation energy network and their solubility is determined by cations solvation energy. Thus, the POMs acids are very soluble in polar solvents such as water and esters. The potassium, sodium and ammonium salts are water soluble while FCUP 67 Lanthanopolyoxometalates encapsulated into silica nanoparticles

the organic molecule salts such as tetrabutylammonium[32, 33] or tetrabutylphosphonium derivatives[34] are generally soluble in nonaqueous solvents.

3.2. Keggin anion

As mentioned before among the most studied and known POMs structures are the Keggin anions. This anion that got its name from the author who made its structural [26] n- 6+ 6+ 5+ characterization in 1934 has a general formula [XM12O40] (M = Mo , W ; X = P , As5+, Ge6+, Si6+, B3+, Fe3+, Co2+, etc.) and presents a tetrahedral symmetry.[21] The Keggin structure is constituted by a central atom X, tetrahedral bonded to four oxygen atoms forming a XO4 group. The XO4 tetrahedron is surrounded by twelve octahedrons

MO6 that can be arranged into four groups of three octahedral units, M3O13, by edge- sharing of the MO6 units (Figure 3.4). These octahedral units bind each other through [5] corner-shared oxygen atoms and through the central XO4 tetrahedron.

Figure 3.4 - Polyhedral representation of the Keggin structure showing the four groups M3O13 in four different colors and [35] the central tetrahedron XO4 in yellow. (Source: Al-Kadamany )

The Keggin anion has several geometrical isomers (rotational isomers or isomers [36] Baker-Figgis) , resulting from a 60º rotation of a M3O13 group relatively to the isomer α (Keggin anion). From the isomer α, isomers β, γ, δ and ε can be obtained by a 60º rotation of one, two, three or four groups M3O13 respectively (Figure 3.5). These rotational orientations of the M3O13 units lower the symmetry of the overall structure. Among these isomers, the α-isomer is the most studied in which the metal centers are all equivalent.[5, 21] 68 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles

Figure 3.5 - Polyhedral representation of the five rotational isomers of the Keggin anion. The rotated M3O13 groups are highlighted (dark blue). (source: Lopez et al.[22])

The metal-oxygen bonds present in a POM structure are arranged in such a framework that can be divided according to position of the oxygen atoms in the structure. Thus an oxygen atom linked to the central atom X (in case of the

heteropolyanions) is designated by Oa, atoms that share a corner or an edge are [37-39] designated as Ob and Oc respectively and Od represents a terminal oxygen. In Figure 3.6 is shown a schematic representation of the relative positions of the different oxygen atoms present in a POM structure using as example the Keggin anion α- n- [XM12O40] .

n- Figure 3.6 - Ball and stick (left) and polyhedral representation (right) for the α-[XM12O40] Keggin anion showing the different classification of the oxygen atoms. FCUP 69 Lanthanopolyoxometalates encapsulated into silica nanoparticles

From the Keggin anion it is possible to obtain several lacunar structures by [5, 21] (n+4)- removing one or more MOx octahedrons. The monolacunar anion [XM11O39] derives from the removal of a MO4+ unit (a metal with its terminal oxygen) by alkaline hydrolysis, giving origin to a gap with five oxygen atoms potentially coordinating (Figure 3.7).

(n+4)- Figure 3.7 - Formation scheme of the monolacunar anion [XM11O39]

The monolacunar anion [XM11O39](n+4)- can then coordinate with metallic cations and originate complexes of the type 1:1 [XM11M’(L)O39]n- or 1:2 [M’(XM11O39)2]n- (Figure 3.8).

n- n- Figure 3.8 - Representation of the complexes of the type 1:1 [XM11M’(L)O39] (left) and 1:2 [M’(XM11O39)2] (right).

The complexes of the type 1:1 are formed when the metallic cation (M’) is a transitional metal (such as V3+, Mn2+ or Co2+) or an element from the p group (e.g. Al3+, Ga3+ or Ge4+). In these complexes, to maintain the octahedral coordination of the ion M’ a monodentate ligand (L) is used. In the case of lanthanide ions, since they are larger ions, bind preferentially to two lacunar units forming 1:2 complexes in which the metal coordinates through eight bonds (four with each lacunar unit).

70 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles

3.3. Polyoxometalates containing lanthanide ions

Lanthanide (Ln) ions, when combined with POMs, confer additional properties, such as excellent luminescent characteristics. This property makes the lanthanide- substituted POMs useful in biological applications where the use of a luminescent probe is needed.

Most of the designed and prepared LnPOMs are modifications of the classic POM anions (Keggin, Well-Dawson, Preyssler and others).[5, 10] The synthesis of these compounds is normally made in two steps. First, the POM structure is transformed into a vacant species (lacunary polyanion) by the removal of at least one MOx group from its structure. Afterwards, the lacunary POM acts as an inorganic ligand that can coordinate with Ln3+ cations through the free oxygen atoms in the lacunary region of the POM, giving rise to the LnPOM.[5]

n- 3.3.1. Keggin-type lanthanide polyoxometalates ([Ln(XM11O39)y] )

Keggin-type LnPOMs are generally generated from the lacunary forms of the Keggin anion and can be obtained through the removal of one or more MOx groups by hydrolysis in alkaline conditions.[5]

7- Lacunar anion [PW11O39] is formed when one of the W(VI) metals and its 4- terminally bound oxo group are missing. Lanthanide substituted [PW11Ln(H2O)3O39] is obtained when the lacunar POM acts as an inorganic ligand coordinating lanthanide cations (such as Eu3+, Tb3+, Sm3+ and others). When Keggin structure loses one of the transition metal oxyanions the lacunar POM is formed. Then the lacunar POM coordinates with lanthanide cations and the lanthanide substituted is obtained. If we have two lacunar POMs coordinated to a lanthanide cation we obtain a sandwich type lanthanide substituted (Figure 3.9). FCUP 71 Lanthanopolyoxometalates encapsulated into silica nanoparticles

Figure 3.9 - Formation scheme of the monolacunar (A) and sandwich type (B) lanthanide-substituted Keggin anion n- [Ln(XM11O39)x] .

The Keggin-type LnPOMs was firstly described in 1971 by Peacock and Weakley.[40] The authors describe the preparation of 1:1 and 1:2 complexes of the n- 3+ 4+ 3+ 3+ [Ln(XW11O39)2] type with X = P and Ln = Ce , Ce , Pr and Nd (for 1:1 complex) and X = Si and Ln = Ce3+, Ce4+, Sm3+, Eu3+ and Ho3+ (for 1:2 complex). POM units can be considered as ligands, which bind to lanthanide ions through the four oxygen atoms present in the gap.[5] Currently, almost all the trivalent lanthanide ions as well as the tetravalent Ce4+ and Tb4+ have been incorporated in complexes of the type m- [Ln(XM11O39)2] with M = W and X = P, As, Si, Ge, B, Ga , Zr, Cu or M = Mo and X = P, As, Si and Ge.[5, 41] As an example is the work of Gaunt et al. where the authors 11– describe the synthesis and crystal structures of the 1:2 [Ln(PMo11O39)2] complexes with the entire lanthanide series.[42, 43]

Comparatively to the large number of dimeric (1:2) complexes, the reports regarding the crystal structures of monomeric complexes (1:1) based on the α-isomer of the Keggin anion are much limited. Sadakene[44] and Mialane[45] have reported the 72 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles

5- 3+ 3+ 3+ crystal structures of the silicotungstates [Ln(SiW11O39)(H2O)n] with Ln = La , Ce , Eu3+, Yb3+. Zhang[46] and co-workers have also reported the crystalline structure of the 4- phosphotungstate [Ce(PW11O39)(H2O)2] . All of these complexes form a one- dimensional polymeric chain. Another report of a crystal structure of a monomeric complex (1:1) is the work of Niu et al.[47] These authors isolated and characterized the crystal structures of two series of rare 2:2 lanthanophosphotungstate anions: 6- 3+ 3+ 3+ [{Ln(H2O)3(α-PW11O39)}2] (Ln = Nd and Gd ) and [{Ln(H2O)(α- 10- 3+ 3+ 3+ 3+ 3+ 3+ 3+ PW11O39)(CH3COO}2] (Ln = Sm , Eu , Gd , Tb , Ho and Er ).

3.3.2. Luminescence of lanthanide ions

The lanthanide series comprises a group of fifteen metallic chemical elements with atomic number increasing from 57 (lanthanum - La) to 71 (lutetium - Lu). These fifteen lanthanide elements, along with the chemically similar elements scandium (Sc) and yttrium (Y), are often collectively known as the rare earth elements.[48] The informal chemical symbol Ln is used in general discussions of lanthanide chemistry to refer to any lanthanide. They are termed lanthanide because the lighter elements in the series are chemically similar to lanthanum.

The electronic structure of the lanthanide elements, with minor exceptions, is 4fn 6s2. The exceptions (La, Ce, Gd and Eu) have an electronic configuration 4fn-15d16s2 with n = 1, 2, 8 e 15, respectively. The lanthanide series is characterized by the gradual filling of the 4f electron orbitals, from 4f0 (La) to 4f14 (Lu) even though for this element the filling is not regular. The chemistry of the lanthanides differs from main group elements and transition metals because of the nature of the 4f orbitals. These orbitals are shielded from the atom's environment by the 5s2 and 5p6 layers, which make the 4f electrons practically undisturbed by the ligands present in the first and second coordination spheres.

Lanthanide elements when in form of coordination complexes generally exhibit their +3 oxidation state (Ln3+) , although particularly stable 4f configurations can also give +4 (Ce, Tb) or +2 (Eu, Yb) ions. This configuration is obtained by removal of all electrons from the 6s and 5d electronic orbitals and frequently one electron from the 4f electronic orbital. Emission of lanthanide ions is due to transitions inside the 4f shell (intra configurational f-f transitions). Electrons from the 4f electronic orbital have a high probability of being found close to the nucleus and thus are strongly affected as the FCUP 73 Lanthanopolyoxometalates encapsulated into silica nanoparticles

nuclear charge increases across the series (from lanthanum to lutenium), this results in a corresponding decrease in ionic radius referred to as the lanthanide contraction.[48]

One of the most interesting properties of these ions is their photoluminescence. The emission of lanthanide ions covers a large part of the electromagnetic spectrum. Several lanthanide ions show luminescence in the visible or near-infrared spectral regions while Gd3+ emits in the ultraviolet region. Thus, the colour of the emitted light is dependent on the lanthanide ion. For example, Eu3+ emits red light, Tb3+ green light, Sm3+ orange light, and Tm3+ blue light. In the case of Yb3+, Nd3+, and Er3+ these ions are known for their near-infrared luminescence, while the La3+ and Lu3+ ions do not present emission properties.[49] This is due to the fact that these ions have the 4f electronic orbital empty and completely filled, respectively.

The electrons in the 4f orbitals have limited interaction with the chemical environment of the lanthanide ion, since the 4f orbitals are well shielded by the electrons in 5s2 and 5p6 shells. For this reason lanthanides have long excited lifetime states and an emission spectrum characterized by narrow and well defined peaks (see example for lanthanide ion Eu3+ in Figure 3.10).[50]

Figure 3.10 – Photoluminescence emission spectra of the Eu3+ ion in water. The radiative transitions take place from the 5 [50] D0 level. (Adapted from Werts )

In Table 3.1 are listed some of the commonly observed emission bands of the lanthanide ions Eu3+, Tb3+, Nd3+, Er3+ and Yb3+ in solution. There are many ways to distribute the electrons over the seven 4f orbitals (see Figure 3.11), but some electron distributions are energetically more favourable than others. Not all transitions are allowed since they have to obey selection rules. One of these is Laporte’s rule in which 74 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles

the sum of the angular momenta of the electrons in the initial and final states must change by an odd integer.[51] That is the case of electronic transitions between f orbitals in lanthanides ions. As a result lanthanide ions suffer from weak light absorption and because of the low molar absorption coefficients ε (smaller than 10 L mol-1 cm-1) only a very limited amount of radiation is absorbed by direct excitation in the 4f levels.[49]

Table 3.1 - Commonly observed emission bands of the lanthanide ions Eu3+, Tb3+, Nd3+, Er3+ and Yb3+ in solution. (Adapted from Werts[50])

Ion Transition λemission (nm) Ion Transition λemission (nm)

5 7 4 4 D0 → F0 580 F3/2 → F9/2 880

5 7 3+ 4 4 D0 → F1 590 Nd F3/2 → F11/2 1060

5 7 4 4 D0 → F2 613 F3/2 → F13/2 1330 Eu3+ 5 7 D0 → F3 650

5 7 D0 → F4 690

5 7 3+ 4 4 D0 → F5 710 Er I13/2 → I15/2 1150

5 7 D4 → F6 490

5 7 D4 → F5 545

3+ 5 7 3+ 2 2 Tb D4 → F4 590 Yb F5/2 → F7/2 980

5 7 D4 → F3 620

5 7 D4 → F2 650

However, the problem related with weak light absorption can be improved through the use of an organic chromophore-containing ligand coordinated to the lanthanide ion the so called indirect excitation, sensitization or antenna effect. In this energy transfer process the antenna chromophore, with a high molar excitation coefficient, absorbs the incoming radiation and transfers the energy to the ion, leading to indirect excitation of the lanthanide. The sensitisation process is often extremely efficient, and high emission quantum yields for lanthanide ions, especially for Eu3+ and Tb3+ ions, can be achieved.[52] FCUP 75 Lanthanopolyoxometalates encapsulated into silica nanoparticles

Another way to enhance the luminescence of lanthanide (III) complexes is by charge transfer states.[53, 54] Charge transfer states often occur in inorganic ligand chemistry involving metals and depending on the direction of charge transfer they are classified as either ligand-to-metal (LMCT) or metal-to-ligand (MLCT) charge transfer. In the case of LnPOMs the charge transfer states are generally of the type LMCT, associated to the O→Ln and O→M transitions.[3, 55, 56] LMCT transitions between the Ln3+ orbitals and the POM orbitals results in intense bands, usually with high intensity in the absorption spectra. In these processes of charge transfer, the radiation is absorbed by LMCT state of high intensity, which subsequently transfers the energy to the lanthanide ion.

Figure 3.11 - Interactions leading to the different electronic energy levels for Eu3+ configuration ([Xe] 4f65d0 – six electrons in the 4f orbitals). (Source: Werts[50])

3.4. Silica encapsulation of LnPOMs

The direct use of POMs for in vivo applications is hindered by their possible toxicity. One possible way to overcome this issue is by coating the composites with amorphous silica.

The first work reporting the encapsulation of a luminescent polyoxometalate 13- ([Eu(SiMoW10O39)2] ) into silica nanoparticles was performed by Green et al. in 76 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles

2005.[57] Thereafter, other studied were developed and some examples of that are the works of Balula[58] et al. in 2006 and Granadeiro[59] et al. in 2010. In the first example the authors reported the incorporation of new lanthanotungstocobaltates 7- 3+ 3+ 3+ 3+ 3+ 3+ ([Ln(CoW11O39)(H2O)3] with Ln = Ce , La , Eu , Sm and Tb ) into spherical silica particles in an attempt to prepare new hybrid materials with combined properties such as, luminescence and magnetism.[58] In the second work the authors have described 9- 3+ 3+ 3+ 3+ the preparation of core/shell nanoparticles using [Ln(W5O18)2] (Ln = Eu , Tb , Gd ) Lindqvist-type derivatives.[59] These multi-wavelength systems allow tuning the excitation wavelength by changing the lanthanide center or through the coordination with an organic ligand to the POM. The same method has also been used by Sousa and co-workers to prepare heterogeneous catalysts based on POM-supported silica nanoparticles.[60] The authors used iron(III)-substituted Keggin derivatives with the corresponding nanocomposites exhibiting higher selectivity for the oxidation of geraniol than the isolated POMs.

Another approach for the synthesis of LnPOM-containing silica nanoparticles is the method described by Zhao and co-workers.[61, 62] In this method, the silica nanoparticles are prepared using surfactant-encapsulated polyoxometalates (SEPs) which are obtained through the replacement of the counterions of the POM structure with cationic surfactants. Silica nanoparticles were obtained using the Preyssler- and Lindqvist-type europium-tungstates corresponding SEPs via a sol-gel reaction with TEOS. The former composites exhibited an interesting photochromic behaviour allowing for the in situ encapsulation of metallic nanoparticles, while the latter have been shown to have potential applications in cell labelling.

3.5. Applications of Keggin-type LnPOMs

The large diversity of structures and the high number of elements that may constitute POMs, leads to the application of these compounds in a variety of fields such as catalysis, medicine, biology, analytical chemistry and more recently in nanotechnology.[2] Apart from that the ability of POMs to behave as inorganic ligands capable of coordinating to lanthanide ions has also received extraordinary scientific interest, leading to the preparation and characterization of a wide diversity of LnPOMs. The combination of lanthanide cations and POMs originates novel compounds with notable properties, remarkable structural features and potential applications in various FCUP 77 Lanthanopolyoxometalates encapsulated into silica nanoparticles

technological areas, such as catalysis, optical/magnetic sensors and medical imaging.[5] LnPOMs can be incorporated in several composites, namely nanostructured thin films, silica NPs, layered double hydroxides and metal-organic frameworks (MOFs).

In the particular case of the Keggin type LnPOMs, this type of composite has been applied as a catalyst in the oxidation of alcohols, alkenes and aldehydes, and also as electrocatalysts and photocatalysts.[5] Griffith and coworkers[63] were the first to report 11- 3+ 3+ 3+ the use of lanthanophosphopolyoxotungstates ([Ln(PW11O39)2)] with Ln = La , Pr , 3+ 3+ Sm and Tb ) as oxidation catalysts in the presence of hydrogen peroxide (H2O2).

These LnPOMs with H2O2 as co-oxidant catalyse the oxidation of primary and secondary alcohols to aldehydes and ketones and the epoxidation of alkenes. More recently Kholdeeva et al.[64] reported the oxidation of formaldehyde mediated by cerium 4- (Ce) containing POM. The [CeSiW11O39] complex was shown to be a selective and effective catalyst for the aerobic oxidation of formaldehyde to formic acid under mild conditions, including the ambient ones. A novel application of LnPOMs as catalyst in oxidative desulfurization (ODS) process was report by Ribeiro et al.[6] In this work the authors describe the use of a LnPOM incorporated into a metal-organic-framework (MOF) as catalyst to complete desulfurization of sulphur refractory compounds from model oil. The LnPOMs-MOFs composite revealed to be an effective catalyst for ODS of oils containing refractory sulphur compounds. Furthermore these composites are recyclable which allows their use in several cycles.[6] In electrocatalysis field Cheng and coworkers[65], for example, reported the use of mixed addenda molybdotungstates coordinated with neodymium (Nd) as catalysts. In this report the authors studied the 13- catalytic property of [Nd(SiMo7W4)] and this complex proved to be an efficient catalyst in the reduction of bromate to bromide in aqueous solution. POMs can also be applied as photocatalysts in degradation of organic pollutants. In the last two decades they gathered attention exhibiting potential application to degrade and mineralize organic pollutants in wastewater. An example of this is the work of Feng et al.[66] where is reported the study on photodegradation of the organic pollutant Azo dye by polyoxometalates/polyvinyl alcohol complexes. Azo dyes contain some aromatic hydrocarbons such as methyl orange, Congo red or Panceau 2R, hazardous chemicals for the water environment. The photocatalysts were prepared by using LnPOMs as the active sites and polyvinyl alcohol (PVA) as a support. The LnPOMs were first immobilized into PVA support in order to decrease water solubility of POMs and enable their recovery from the reaction system and subsequent recycling. A series of 78 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles

3+ 3+ 3+ 3+ 3+ 3+ photocatalysts [Ln(PW11O39)2]/PVA with Ln = La , Ce , Pr , Nd and Sm ) were prepared and used to degrade the three kinds or aromatic hydrocarbons presented in Azo dyes mentioned above. The photocatalysts exhibited efficient catalytic activity to degrade Azo dyes with high degradation conversions. Furthermore

[Ce(PW11O39)2]/PVA showed the best catalytic activity exhibiting potential for practical applications.

In the case of Keggin type LnPOMs containing gadolinium these have found applications as MRI contrast agents. Feng and coworkers[67] for example reported the 9- 11- use of two gadolinium (Gd) containing POMs ([GdW10O36] and [Gd(PW11O39)2] ) for in vitro and in vivo tissue-specific MRI contrast agents. Both LnPOMs presented favourable tissue-specificity to liver and kidney with relaxivities slightly higher than the 9- commercial and widely used MRI contrast agent Gd-DTPA. Furthermore [GdW10O36] was found to be helpful in the diagnosis of stomach pathological states. Sun et al.[68] reported a similar study of two gadolinium-sandwich complexes with tungstosilicates 13- 11- [Gd(SiW11O39)2] and [Gd3O3(SiW9O34)2] . Again both complexes were used as tissue-specific contrast agents and were evaluated by in vivo relaxation measurements. Similar to the previous work both gadolinium complexes exhibit higher relaxivity than the widely used Gd-DTPA contrast agent. MRI experiments showed signal enhancement in liver and kidney. However, toxicity test on these complexes have shown that these complexes were too toxic and need to be modified for further clinic use.

Recently, Coronado et al.[8, 9] have published studies on the behavior of POMs as 9- single molecular magnets (SMM), particularly for complexes of the type [Ln(W5O18)2] 3+ 3+ 3+ 13- 3+ 3+ 3+ 3+ 3+ with Ln = Ho , Er and [Ln(SiW11O39)2] with Ln = Dy , Ho , Er , Yb . These compounds exhibit a slower relaxation of magnetization, which is a characteristic behavior of the SMM. Thus, the application of POMs can be extended to new fields such as, quantum computers and high-density magnetic memories.[69]

Despite the remarkable potentialities revealed by these compounds industrial and technological applications of LnPOMs are still limited. The development of more effective methods of incorporation, encapsulation and/or immobilization of the LnPOMs in support systems and the design and development of new materials could be the route to overcome this limited applicability.

FCUP 79 Lanthanopolyoxometalates encapsulated into silica nanoparticles

3.6. References

1. Liu, S.Q. and Tang, Z.Y., Polyoxometalate-based functional nanostructured films: Current progress and future prospects. Nano Today, 2010. 5(4): p. 267- 281. 2. Long, D.L. and Cronin, L., Towards polyoxometalate-integrated nanosystems. Chemistry - A European Journal, 2006. 12(14): p. 3698-706. 3. Yamase, T., Photo- and electrochromism of polyoxometalates and related materials. Chemical Reviews, 1998. 98(1): p. 307-325. 4. Pope, M.T., in Handbook on the Physics and Chemistry of Rare Earths, Gschneider, J.C.B.K.A. and Pecharsky, V.K., Editors. 2007, Elsiver. p. 337-382. 5. Granadeiro, C.M., de Castro, B., Balula, S.S. and Cunha-Silva, L., Lanthanopolyoxometalates: From the structure of polyanions to the design of functional materials. Polyhedron, 2013. 52: p. 10-24. 6. Ribeiro, S., Granadeiro, C.M., Silva, P., Paz, F.A.A., de Biani, F.F., Cunha- Silva, L. and Balula, S.S., An efficient oxidative desulfurization process using terbium-polyoxometalate@MIL-101(Cr). Catalysis Science & Technology, 2013. 3(9): p. 2404-2414. 7. Siozaki, R., Inagaki, A., Kominami, H., Yamaguchi, S., Ichihara, J. and Kera, Y., Catalytic properties of holmiumdecatungstate modified with cetylpyridinium cation [{C5H5N(CH2)(15)CH3}(7)H2Ho(III)W10O36; Cetyl-HoW10] for H2O2- oxidation of alcohols and olefins and its working states under an organic- solvent-free condition. Journal of Molecular Catalysis A: Chemical, 1997. 124(1): p. 29-37. 8. AlDamen, M.A., Clemente-Juan, J.M., Coronado, E., Marti-Gastaldo, C. and Gaita-Arino, A., Mononuclear lanthanide single-molecule magnets based on polyoxometalates. Journal of the American Chemical Society, 2008. 130(28): p. 8874-5. 9. AlDamen, M.A., Cardona-Serra, S., Clemente-Juan, J.M., Coronado, E., Gaita- Arino, A., Marti-Gastaldo, C., Luis, F. and Montero, O., Mononuclear Lanthanide Single Molecule Magnets Based on the Polyoxometalates [Ln(W5O18)(2)](9-) and [Ln(beta(2)-SiW11O39)(2)](13-) (Ln(III) = Tb, Dy, Ho, Er, Tm, and Yb). Inorganic Chemistry, 2009. 48(8): p. 3467-3479. 10. Granadeiro, C.M., Ferreira, R.A.S., Soares-Santos, P.C.R., Carlos, L.D. and Nogueira, H.I.S., Lanthanopolyoxometalates as Building Blocks for 80 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles

Multiwavelength Photoluminescent Organic-Inorganic Hybrid Materials. European Journal of Inorganic Chemistry, 2009(34): p. 5088-5095. 11. Yamase, T., Anti-tumor, -viral, and -bacterial activities of polyoxometalates for realizing an inorganic drug. Journal of Materials Chemistry, 2005. 15(45): p. 4773-4782. 12. Hasenknopf, B., Polyoxometalates: Introduction to a class of inorganic compounds and their biomedical applications. Frontiers in Bioscience- Landmark, 2005. 10: p. 275-287. 13. Guerrero-Martinez, A., Perez-Juste, J. and Liz-Marzan, L.M., Recent progress on silica coating of nanoparticles and related nanomaterials. Advanced Materials, 2010. 22(11): p. 1182-95. 14. Slowing, I.I., Trewyn, B.G., Giri, S. and Lin, V.S.Y., Mesoporous silica nanoparticles for drug delivery and biosensing applications. Advanced Functional Materials, 2007. 17(8): p. 1225-1236. 15. Wang, J., Liu, G.D. and Lin, Y.H., Electroactive silica nanopartictes for biological Labeling. Small, 2006. 2(10): p. 1134-1138. 16. Fernández, X.L., Theoretical Study of the Basicity and the Redox Properties of Heteropolyanions, in Departament de Química Física i Inorgànica. 2003, Universitat Rovira i Virgili: Tarragona. 17. Borrás-Almenar, J.J., Coronado, E., Müller, A. and Pope, M., eds. Polyoxometalate Molecular Science. 2003, Kluwer Academic Publishers: Dordrecht. 18. Pope, M.T. and Müller, A., eds. Polyoxometalates: from platonic solids to anti- retroviral activity. 1994, Kluwer Academic Publishers: Dordrecht. 19. Cavaleiro, A.M.V., Pedrosa de Jesus, J.D. and Nogueiva, H.I.S., Complexes of Keggin-Type Monolacunary Heteropolytungstates: Synthesis and Characterisation, in Metal Clusters in Chemistry, P. Braunstein, L. A. Or0 and Raithby, P.R., Editors. 1999, Wiley-VCH: Weinheim, Germany. p. 444-458. 20. Pope, M.T., Heteropoly and Isopoly Oxometalates. 1983, Berlin: Springer- Verlag. 21. Gamelas, J., Cavaleiro, A., Santos, I. and Balula, M.S., Os Polioxometalatos. Do Anião de Keggin às Nanocápsulas. Boletim da Sociedade Portuguesa de Química, 2003. 90: p. 45-51. 22. Lopez, X., Carbo, J.J., Bo, C. and Poblet, J.M., Structure, properties and reactivity of polyoxometalates: a theoretical perspective. Chemical Society Reviews, 2012. 41(22): p. 7537-71. FCUP 81 Lanthanopolyoxometalates encapsulated into silica nanoparticles

23. Berzelius, J.J., Beitrag zur näheren kenntniss des molybdäns. Annalen der Physik, 1826. 82(4): p. 369-392. 24. Marignac, C., Comptes rendus de l'Académie des Sciences, 1862. 55: p. 888- 892. 25. Pauling, L., The molecular structure of the tungstosilicates and related compounds. Journal of the American Chemical Society, 1929. 51: p. 2868- 2880. 26. Keggin, J.F., The structure and formula of 12-phosphotungstic acid. Proceedings of the Royal Society of London Series a-Containing Papers of a Mathematical and Physical Character, 1934. 144(A851): p. 0075-0100. 27. Anderson, J.S., Constitution of the poly-acids. Nature, 1937. 140: p. 850-850. 28. Evans, H.T., The Crystal Structures of Ammonium and Potassium Molybdotellurates. Journal of the American Chemical Society, 1948. 70(3): p. 1291-1292. 29. Lindqvist, I., A Crystal Structure Investigation of the Paramolybdate Ion. Arkiv for Kemi, 1950. 2(4): p. 325-341. 30. Dawson, B., The Structure of the 9(18)-Heteropoly Anion in Potassium 9(18)- Tungstophosphate, K6(P2w18o62).14h2o. Acta Crystallographica, 1953. 6(2): p. 113-126. 31. Wells, A.F., Structural Inorganic Chemistry Structural Inorganic Chemistry. Vol. 1945. 1945, Oxford: Oxford University Press. 344-345. 32. Bartis, J., Dankova, M., Lessmann, J.J., Luo, Q.H., Horrocks, W.D. and Francesconi, L.C., Lanthanide complexes of the alpha-1 isomer of the [P2W17O61](10-) heteropolytungstate: Preparation, stoichiometry, and structural characterization by W-183 and P-31 NMR spectroscopy and europium(III) luminescence spectroscopy. Inorganic Chemistry, 1999. 38(6): p. 1042-1053. 33. Lis, S., But, S. and Meinrath, G., Synthesis and spectroscopic characterisation of chosen heteropolyanions and their Ln(III) complexes containing tetrabutylammonium counter ion. Journal of Alloys and Compounds, 2004. 374(1-2): p. 366-370. 34. Lis, S., But, S. and Meinrath, G., Spectroscopic characterization of chosen Ln(III) polyoxometalate complexes with organic counter cations in solid and in non-aqueous solutions. Journal of Alloys and Compounds, 2006. 408: p. 958- 961. 82 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles

35. Al-Kadamany, G., Synthesis, Structure and Catalytic Activity of Titanium, Zirconium and Hafnium-Containing Polyoxometalates, in School of Engineering and Science. 2010, Jacobs University: Bremen. p. 147. 36. Baker, L.C.W. and Figgis, J.S., A New Fundamental Type of Inorganic Complex - Hybrid between Heteropoly and Conventional Coordination Complexes - Possibilities for Geometrical Isomerisms in 11-Heteropoly, 12-Heteropoly, 17- Heteropoly, and 18-Heteropoly Derivatives. Journal of the American Chemical Society, 1970. 92(12): p. 3794-&. 37. Poblet, J.M., Lopez, X. and Bo, C., Ab initio and DFT modelling of complex materials: towards the understanding of electronic and magnetic properties of polyoxometalates. Chemical Society Reviews, 2003. 32(5): p. 297-308. 38. Rocchicciolideltcheff, C., Fournier, M., Franck, R. and Thouvenot, R., Vibrational Investigations of Polyoxometalates .2. Evidence for Anion Anion Interactions in Molybdenum(Vi) and Tungsten(Vi) Compounds Related to the Keggin Structure. Inorganic Chemistry, 1983. 22(2): p. 207-216. 39. Rocchiccioli-Deltcheff, C., Thouvenot, R. and Franck, R., Spectres i.r. et Raman d'hétéropolyanions α - XM12O40n− de structure de type Keggin (X = BIII, SiIV, GeIV, PV, AsV et M = WVI et MoVI). Spectrochimica Acta Part A: Molecular Spectroscopy, 1976. 32(3): p. 587-597. 40. Peacock, R.D. and Weakley, T.J.R., "Heteropolytungstate complexes of lanthanide elements. Part I. Preparation and reactions". Journal of the Chemical Society A, 1971. 11: p. 1836-1839. 41. Muller, A., Peters, F., Pope, M.T. and Gatteschi, D., Polyoxometalates: Very Large Clusters-Nanoscale Magnets. Chemical Reviews, 1998. 98(1): p. 239- 272. 42. Gaunt, A.J., May, I., Sarsfield, M.J., Collison, D., Helliwell, M. and Denniss, I.S., A rare structural characterisation of the phosphomolybdate lacunary anion, [PMo11O39](7-). Crystal structures of the Ln(III) complexes, (NH4)(11)[Ln(PMo11O39)(2)]center dot 16H(2)O (Ln = Ce-III, Sm-III, Dy-III or Lu-III). Dalton Transactions, 2003(13): p. 2767-2771. 43. Copping, R., Gaunt, A.J., May, I., Sarsfield, M.J., Collison, D., Helliwell, M., Denniss, I.S. and Apperley, D.C., Trivalent lanthanide lacunary phosphomolybdate complexes: a structural and spectroscopic study across the series [Ln(PMo11O39)2]11. Dalton Transactions, 2005(7): p. 1256-62. 44. Sadakane, M., Dickman, M.H. and Pope, M.T., Controlled Assembly of Polyoxometalate Chains from Lacunary Building Blocks and Lanthanide-Cation FCUP 83 Lanthanopolyoxometalates encapsulated into silica nanoparticles

Linkers Supported in part by the National Science Foundation (CHE9727417) and Georgetown University. Angewandte Chemie International Edition in English, 2000. 39(16): p. 2914-2916. 45. Mialane, P., Lisnard, L., Mallard, A., Marrot, J., Antic-Fidancev, E., Aschehoug, P., Vivien, D. and Secheresse, F., Solid-State and solution studies of [Ln(n)(SiW11O39)] polyoxoanions: an example of building block condensation dependent on the nature of the rare earth. Inorganic Chemistry, 2003. 42(6): p. 2102-8. 46. Zhang, C., Ma, P.T., Chen, H.N., Wang, J.P. and Niu, J.Y., Synthesis, structure, and properties of a 1-D cerium based on monovacant Keggin-type polyoxotungstate. Journal of Coordination Chemistry, 2011. 64(12): p. 2178- 2185. 47. Niu, J.Y., Wang, K.H., Chen, H.N., Zhao, J.W., Ma, P.T., Wang, J.P., Li, M.X., Bai, Y. and Dang, D.B., Assembly Chemistry between Lanthanide Cations and Monovacant Keggin Polyoxotungstates: Two Types of Lanthanide Substituted Phosphotungstates [{(alpha-PW11O39H)Ln(H2O)(3)}(2)](6-) and [{(alpha- PW11O39)Ln(H2O)(eta(2),mu-1,1)-CH3COO}(2)](10-). Crystal Growth & Design, 2009. 9(10): p. 4362-4372. 48. Gschneidner, K.A. and Eyring, L.R., eds. Handbook on the Physics and Chemistry of Rare Earths: Metals. Vol. 1. 1978, North-Holland: Nova Iorque, EUA,. 49. Binnemans, K., Lanthanide-Based Luminescent Hybrid Materials. Chemical Reviews, 2009. 109(9): p. 4283-4374. 50. Werts, M.H.V., Making sense of lanthanide luminescence. Science Progress, 2005. 88(2): p. 101-131. 51. Eliseeva, S.V. and Bunzli, J.C., Lanthanide luminescence for functional materials and bio-sciences. Chemical Society Reviews, 2010. 39(1): p. 189- 227. 52. Van der Tol, E.B., Ramesdonk, H.J.V., Verhoeven, J.W., Steemers, F.J., Kerver, E.G., Verboom, W. and Reinhoudt, D.N., Tetraazatriphenylenes as Extremely Efficient Antenna Chromophores for Luminescent Lanthanide Ions. Chemistry - A European Journal, 1998. 4(11): p. 2315-2323. 53. Faustino, W.M., Malta, O.L. and de Sa, G.F., Intramolecular energy transfer through charge transfer state in lanthanide compounds: A theoretical approach. Journal of Chemical Physics, 2005. 122(5). 84 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles

54. D'Aleo, A., Picot, A., Beeby, A., Williams, J.A.G., Le Guennic, B., Andraud, C. and Maury, O., Efficient Sensitization of Europlum, Ytterbium, and Neodymium Functionalized Tris-Dipicolinate Lanthanide Complexes through Tunable Charge-Transfer Excited States. Inorganic Chemistry, 2008. 47(22): p. 10258- 10268. 55. Ferreira, R.A.S., Nobre, S.S., Granadeiro, C.M., Nogueira, H.I.S., Carlos, L.D. and Malta, O.L., A theoretical interpretation of the abnormal D-5(0)-> F-7(4) intensity based on the Eu3+ local coordination in the Na-9[EuW10O36] center dot 14H(2)O polyoxometalate. Journal of Luminescence, 2006. 121(2): p. 561- 567. 56. Ballardini, R., Chiorboli, E. and Balzani, V., Photophysical properties of Eu(SiW11O39)2 13- and Eu(BW11O39)2 15-. Inorganica Chimica Acta, 1984. 95(6): p. 323-327. 57. Green, M., Harries, J., Wakefield, G. and Taylor, R., The synthesis of silica nanospheres doped with polyoxometalates. Journal of the American Chemical Society, 2005. 127(37): p. 12812-12813. 58. Balula, M.S.S., Nogueira, H.I.S. and Cavaleiro, A.M.V., New Polyoxotungstates with Ln(III) and Co(II) and their Immobilization in Silica Particles. Materials Science Forum, 2006. Vols. 514-516: p. 1206-1210. 59. Granadeiro, C.M., Ferreira, R.A.S., Soares-Santos, P.C.R., Carlos, L.D., Trindade, T. and Nogueira, H.I.S., Lanthanopolyoxotungstates in silica nanoparticles: multi-wavelength photoluminescent core/shell materials. Journal of Materials Chemistry, 2010. 20(16): p. 3313-3318. 60. Sousa, J.L.C., Santos, I.C.M.S., Simoes, M.M.Q., Cavaleiro, J.A.S., Nogueira, H.I.S. and Cavaleiro, A.M.V., Iron(III)-substituted polyoxotungstates immobilized on silica nanoparticles: Novel oxidative heterogeneous catalysts. Catalysis Communications, 2011. 12(6): p. 459-463. 61. Zhao, Y.Y., Li, Y., Li, W., Wu, Y.Q. and Wu, L.X., Preparation, Structure, and Imaging of Luminescent SiO2 Nanoparticles by Covalently Grafting Surfactant- Encapsulated Europium-Substituted Polyoxometalates. Langmuir, 2010. 26(23): p. 18430-18436. 62. Zhao, Y.Y., Qi, W., Li, W. and Wu, L.X., Covalent Dispersion of Surfactant- Encapsulated Polyoxometalates and In Situ Incorporation of Metal Nanoparticles in Silica Spheres. Langmuir, 2010. 26(6): p. 4437-4442. FCUP 85 Lanthanopolyoxometalates encapsulated into silica nanoparticles

63. Griffith, W.P., Moreea, R.G.H. and Nogueira, H.I.S., Lanthanide complexes as oxidation catalysts for alcohols and alkenes. Polyhedron, 1996. 15: p. 3493- 3500. 64. Kholdeeva, O.A., Timofeeva, M.N., Maksimov, G.M., Maksimovskaya, R.I., Neiwert, W.A. and Hill, C.L., Aerobic oxidation of formaldehyde mediated by a Ce-containing polyoxometalate under mild conditions. Inorganic Chemistry, 2005. 44(3): p. 666-672. 65. Cheng, L., Zhang, X.M., Xi, X.D., Liu, B.F. and Dong, S.J., Electrochemical behavior of the molybdotungstate heteropoly complex with neodymium, K10H3[Nd(SiMo7W4O39)(2)]center dot xH(2)O in aqueous solution. Journal of Electroanalytical Chemistry, 1996. 407(1-2): p. 97-103. 66. Feng, C., Zhuo, X. and Liu, X.J., Study on photodegradation of Azo dye by polyoxometalates/polyvinyl alcohol Journal of Rare Earths, 2009. 27: p. 717- 722. 67. Feng, J.H., Li, X.J., Pei, F.K., Sun, G.Y., Zhang, X. and Liu, M.L., An evaluation of gadolinium polyoxornetalates as possible MRI contrast agent. Magnetic Resonance Imaging, 2002. 20(5): p. 407-412. 68. Sun, G.Y., Feng, J.H., Wu, H.F., Pei, F.K., Fang, K. and Lei, H., Investigation of sandwiched gadolinium(III) complexes with tungstosilicates as potential MRI contrast agents. Magnetic Resonance Imaging, 2004. 22(3): p. 421-426. 69. Lehmann, J., Gaita-Arino, A., Coronado, E. and Loss, D., Spin qubits with electrically gated polyoxometalate molecules. Nature Nanotechnology, 2007. 2(5): p. 312-7. 86 FCUP

II Research work

FCUP 89

Scope of the thesis

The main purpose of this research was to develop fluorescent silica nanoparticles that could incorporate organic (rhodamine b isothiocyanate – RBITC) and inorganic (lanthanide-based polyoxometalates – LnPOMs) fluorophores. Due to the unique chemical and optical properties such as bright fluorescence, high photostability and biocompatibility fluorescent silica nanoparticles have received an increasing interest for biological applications in the past few years and are considered as an alternative to the classical fluorophores.

Fluorescent silica nanoparticles containing RBITC molecules as the fluorophore were synthesized using the reverse microemulsion technique for the alkaline hydrolysis of TEOS. In this particular case, the dye was firstly coupled to a silane coupling agent (3-aminopropyl triethoxysilane – APTES), and the reaction product was incorporated into silica spheres by hydrolysis and polymerisation of TEOS in alkaline media. The obtained nanoparticles were further functionalized, so that the particles could bind to biologically active molecules, such as oligonucleotides. Lifetime measurements and steady-state anisotropy studies of the nanoparticles and the free dye were also performed to evaluate the effect of the encapsulation on the fluorescence emission properties of RBITC. (Chapter 4)

Regarding the production of fluorescent silica nanoparticles encapsulating inorganic fluorophores, two europium-polyoxometalates with different europium coordination environments were encapsulated into a silica matrix through the reverse microemulsion technique mentioned previously. The prepared nanoparticles were characterized and the stability of the material and the integrity of the europium compounds incorporated were also examined. Furthermore the photo-luminescence properties of the synthesized nanoparticles were evaluated and compared with the free europium- polyoxometalates. (Chapter 5)

To evaluate the potential cytotoxicity of the synthesized NPs a cell viability test was performed in three different human cell models (intestinal epithelial Caco-2 cells, neuroblastoma SH-SY5Y cells and hepatoma RG cells), using a calcein-AM assay. After incubation of cells with the NPs, cell morphology was evaluated by phase contrast microscopy and in the particular case of NPs containing RBITC molecules a cellular uptake experiment was also performed. (Chapter 6) 90 FCUP

FCUP 91

Experimental Background

As mentioned previously in chapter 2 (section 2.1) there are two general routes for synthesizing fluorescent silica NPs, the Stöber method and the reverse microemulsion technique.

Since Stöber’s method is described as a relatively simple procedure to make silica NPs that can be carried out in only few hours, this method was the first choice to prepare the fluorescent silica NPs. In the first attempt to produce fluorescent silica NPs incorporating an organic dye, an adapted procedure described by Bringley[1] and coworkers, based on Stöber’s method, was used. In this procedure, a RBITC dye solution (49.6 µM or 66.7 µM) in ethanol was heated to 65ºC using a controlled temperature bath. To the previous solution was then added 7.62 mL of TEOS followed by 12.0 mL of distilled water and 6.40 mL of a 25% ammonia solution. The mixture was stirred at 65ºC for 3.0 h and afterwards was left to cool to 15 ºC. To the cooled mixture was added 200 mL of ethanol and then the solvent was removed by rotary evaporation. The obtained powder was resuspended in ethanol and washed by repeated cycles of centrifugation/resuspension in ethanol. However, after each washing cycle dye leaching was observed and the final obtained NPs presented low fluorescence intensity under a UV chamber. Furthermore, the corresponding UV-vis spectrum (Figure EB 1) of these nanoparticles does not show any band from the RBITC dye.

1.1

1.0

0.9

Abs 0.8

0.7

0.6

300 400 500 600 700 800 Wavelength (nm)

Figure EB 1 - UV-vis spectrum of fluorescent silica nanoparticles synthesized by Stober’s method through the adapted procedure described by Bringley[1]. 92 FCUP Experimental Background

Since this adapted procedure proved to be inefficient for encapsulation of the fluorophore within the silica matrix another attempt to prepare fluorescent silica nanoparticles was made following a method described in literature by Rossi[2] and coworkers. This procedure was also based on Stöber’s method and is briefly described as follows: to an ethanol solution (25 mL) containing ammonium hydroxide (25% aqueous solution) and RBITC (0.5 mg/mL in water) was added 1.3 mL of TEOS. The

mixture was stirred for 1 h at room temperature and further sonicated for 10 min. Afterwards nanoparticles were isolated by centrifugation and washed by repeated cycles of centrifugation/resuspension in ethanol. Like in the method described previously, dye leaching was observed during washing steps leading to nanoparticles with low or no fluorescence intensity. These results could be due to the fact that both methods describe that dye encapsulation within silica matrix is achieved by physical entrapment of the dye. The physical entrapment of dye molecules is usually obtained by adding the fluorophore to the reaction media but this incorporation method has the disadvantages of low entrapment probability and of dye leakage from the NPs.

To overcome the issue of dye leakage, the fluorescent silica nanoparticles were synthesized following a procedure described by Larson[3] and coworkers. The method reported was also based on Stöber’s method and is described as a two-part process, where dye molecules are first covalently conjugated to a silica precursor such as APTES and then TEOS and ammonium hydroxide are subsequently added to form a silica network around the dye-silica precursor. Dye-silica precursor was synthesized by addition reaction between TRITC and APTES in molar ratio of 1:50, in ethanol under argon atmosphere for 12 h. For the synthesis of the nanoparticles, the aforementioned dye-silica precursor (5 mL; 1.70x10-5 M) was condensed with an aliquot of TEOS (160 µL) and then added to a reaction vessel containing 500 µL of ammonia, 2 mL of water and 125 mL of ethanol, the mixture was left to react overnight. Afterwards TEOS (4.4 mL) was subsequently added in aliquots of 500 µL every 15 min to grow the silica shell. Nanoparticles were isolated by centrifugation and washed by repeated cycles of centrifugation/resuspension in ethanol. In comparison to the methods tested previously from Bringley[1] and Rossi[2], the nanoparticles obtained by this method present a high fluorescence signal. However TEM analysis of these nanoparticles shown that they are not uniform in size and appear to be an intricate net of silica aggregates (Figure EB 2). This could be an issue for further functionalization and bioapplications of these nanoparticles. FCUP 93 Experimental Background

Figure EB 2- TEM images of TRICT fluorescent silica nanoparticles prepared by Stöber’s method following a similar procedure to that described by Larson[3] et al.

Although Stöber’s method presents the advantage of having a reaction that can be scaled up easily to yield large amounts of nanoparticles, it can also lead to particles with non-uniform sizes. So in this scenario and despite the fact of taking 24 to 48 hours to complete the reaction the microemulsion technique appears to be a good alternative to produce fairly uniform and monodisperse nanoparticles.

To produce uniform fluorescent silica nanoparticles through the reverse microemulsion methodology three approaches were tested to determine the most suitable to achieve the purposed aim. These three approaches are all very similar and are listed in Table EB 1. The first one is based in the work of Gao[4] et al. where a water-in-oil microemulsion was prepared by mixing 1.77 mL of triton X-100 (surfactant), 7.5 mL of cyclohexane, 1.8 mL of n-hexanol, and 340 mL of water. An aqueous solution of RBITC (0.1 M; 140 µL) was then added and the mixture was left to homogenize for 30 minutes after which 100 mL of TEOS and 60 mL of ammonium hydroxide were added to the mixture. The reaction was allowed to continue for 24 h. After reaction was complete the nanoparticles were isolated from the reaction media by addition of 20 mL of pure acetone and washed by repeated cycles of centrifugation/resuspension in ethanol to remove any surfactant or unreacted molecules.

In the second approach based on the work of Zhang[5] and coworkers a conjugated reaction between RBITC and APTEs was firstly carried out to enable the covalent 94 FCUP Experimental Background

binding of RBITC to the silica matrix. For that purpose 0.1 mL of APTS was added to 2.5 mL of an anhydrous ethanol solution containing 27.5 mg of RBITC (20.5 mM) and left to react for 24 h in the dark. After that period the solution was centrifuged and the obtained powder was dried in a desiccator for further use. In a second step the reverse microemulsion was prepared by mixing 5.3 mL of Triton X-100, 5.4 mL of n-hexanol, 22.5 mL of cyclohexane, 3.0 mg of RBITC-APTES conjugate dissolved in 1.5 mL of deionized water, and 300 µL of ammonium hydroxide. The microemulsion was stirred for 30 min before 300 µL of TEOS was added. The solution was stirred for another 24 h after which 20 mL of pure acetone were added to precipitate the nanoparticles from the microemulsion. Nanoparticles were then washed by repeated cycles of centrifugation/resuspension in ethanol to remove any surfactant or unreacted molecules.

The third approach is similar to the previous method described and is based in the work of Shi[6] et al. An aqueous solution of RBITC (0.1 M) was prepared and then mixed with an equimolar quantity of APTES, and left to react overnight at room temperature. The RBITC-APTES conjugate was directly used to prepare the fluorescent silica nanoparticles trough the reverse microemulsion technique as follows: 1.77 mL of Triton X-100 was mixed with 7.5 mL of cyclohexane, 1.8 mL of n-hexanol, 400 µL of water and 100 µL of the as prepared RBITC-APTES conjugate. After stirring for 1 hour, 200 µL of TEOS and 100 µL of ammonium hydroxide were then added to the previous mixture and the reaction was allowed to continue for 24 hours at room temperature. When the reaction was completed the nanoparticles were isolated by addition of 20 mL of acetone followed by a washing step through repeated cycles of centrifugation/resuspension in ethanol.

FCUP 95 Experimental Background

Table EB 1– Comparison between the three methods followed to prepare fluorescent silica NPs using the microemulsion technique.

Method

Gao Zhang Shi

1.77 mL Triton X-100 1.80 mL n-hexanol 0.1 mL APTES RBITC aqueous solution 7.50 mL cyclohexane 27.5 mg RBITC (0.1 M), with equimolar 140 µL RBITC aqueous 2.5 mL anhydrous ethanol quantity of APTES in water solution (0.1 M) Homogenize for 30 min React 24h in dark React overnight 5.33 mL Triton X-100 1.77 mL Triton X-100 5.40 mL n-hexanol 1.80 mL n-hexanol Add: 22.50 mL cyclohexane 7.50 mL cyclohexane 100 µL TEOS 3 mg RBITC-APTES 400 µL water 60 µL ammonium hydroxide (dissolved in deionized water 100 µL RBITC-APTES - 1.5 mL) aqueous solution 300 µL ammonium hydroxide 24h reaction Homogenize for 30 min Homogenize for 60 min Add: Add: 200 µL TEOS Washing 300 µL TEOS 100 µL ammonium hydroxide 24h reaction 24h reaction

Washing Washing

All of these three approaches yield uniform and monodisperse nanoparticles (Figure EB 3) with a relatively high fluorescence signal (Figure EB 4). However, since the synthesis route based on method described by Gao[4] uses the a physical strategy to incorporate the dye within the silica matrix which can further lead to dye leaking problems, this method was not considered to be the ideal for the synthesis of the aimed fluorescent nanoparticles. Regarding the two synthesis strategies based on the methods from Zhang[5] and Shi[6] that uses the covalent binding of the dye to the silica matrix, by reaction of the dye to the silane agent APTES before the hydrolysis and condensation of TEOS, the one adapted from Shi appears to be the best synthesis route. Comparing to the Zhang method, Shi’s approach takes less reaction time and uses water instead of anhydrous ethanol, which makes the experimental work easier, since there is no need to work with an inert atmosphere. 96 FCUP Experimental Background

Figure EB 3 - TEM images of fluorescent silica nanoparticles prepared by the reverse microemulsion system following the procedures of Gao[4] (A); Zhang[5] (B) and Shi[6] (C).

1000

C B 800

600

400 A Intensity (a.u)

200

0 550 575 600 625 650 675 700 Wavelength (nm)

Figure EB 4 - Fluorescence emission spectra of fluorescent silica nanoparticles prepared by the reverse microemulsion system following the procedures of Gao[4] (A); Zhang[5] (B) and Shi[6] (C).

Based on the results shown above, the experimental work presented in the next section, regarding the development of fluorescent silica nanoparticles encapsulating the organic fluorophore RBITC, was done following the adapted procedure from Shi and coworkers. In the case of fluorescent silica nanoparticles encapsulating the inorganic fluorophores (lanthanopolyoxometalates), these were also synthesized through the reverse microemulsion technique but following a procedure described by Ye[7] et al. for similar fluorescent silica nanoparticles. FCUP 97 Experimental Background

References

1. Bringley, J.F., Penner, T.L., Wang, R., Harder, J.F., Harrison, W.J. and Buonemani, L., Silica nanoparticles encapsulating near-infrared emissive cyanine dyes. Journal of Colloid and Interface Science, 2008. 320(1): p. 132-9. 2. Rossi, L.M., Shi, L., Quina, F.H. and Rosenzweig, Z., Stober synthesis of monodispersed luminescent silica nanoparticles for bioanalytical assays. Langmuir, 2005. 21(10): p. 4277-80. 3. Larson, D.R., Ow, H., Vishwasrao, H.D., Heikal, A.A., Wiesner, U. and Webb, W.W., Silica nanoparticle architecture determines radiative properties of encapsulated fluorophores. Chemistry of Materials, 2008. 20(8): p. 2677-2684. 4. Gao, F., Wang, L., Tang, L. and Zhu, C., A Novel Nano-Sensor Based on Rhodamine-b-Isothiocyanate –Doped Silica Nanoparticle for pH Measurement. Microchimica Acta, 2005. 152: p. 131-135. 5. Zhang, R.R., Wu, C.L., Tong, L.L., Tang, B. and Xu, Q.H., Multifunctional Core- Shell Nanoparticles as Highly Efficient Imaging and Photosensitizing Agents. Langmuir, 2009. 25(17): p. 10153-10158. 6. Shi, H., He, X.X., Wang, K.M., Yuan, Y., Deng, K., Chen, J.Y. and Tan, W.H., Rhodamine B isothiocyanate doped silica-coated fluorescent nanoparticles (RBITC-DSFNPs)-based bioprobes conjugated to Annexin V for apoptosis detection and imaging. Nanomedicine-Nanotechnology Biology and Medicine, 2007. 3(4): p. 266-272. 7. Ye, Z.Q., Tan, M.Q., Wang, G.L. and Yuan, J.L., Novel fluorescent europium chelate-doped silica nanoparticles: preparation, characterization and time- resolved fluorometric application. Journal of Materials Chemistry, 2004. 14(5): p. 851-856.

98 FCUP

FCUP 99

4. Dye doped fluorescent silica nanoparticles

Incorporation of fluorescent organic molecules inside silica matrixes shields the molecules from several environmental factors that can interfere with their fluorescence emission, photostability or quantum yield. For these reasons fluorescent dye molecules are widely used nowadays doped into silica matrixes to produce fluorescent nanomaterials.

Dye doped fluorescent silica nanoparticles containing RBITC dye molecules were synthesized using the reverse microemulsion technique for the alkaline hydrolysis of TEOS. In this particularly case of silica nanoparticles containing RBITC dye molecules, the dye was firstly coupled to a silane coupling agent (3-aminopropyl triethoxysilane – APTES), and the reaction product was incorporated into silica spheres by hydrolysis and polymerisation of TEOS in alkaline media. The obtained nanoparticles were then characterized by fluorescence and ultraviolet-visible (UV-Vis) spectroscopy, transmission electron microscopy (TEM) and dynamic light scattering (DLS). To investigate the influence of silica encapsulation in the fluorescence emission properties of the dye, lifetime measurements and steady state anisotropy studies were perform for both nanoparticles and the free dye in solution.

The produced RBITC encapsulated silica nanoparticles had a mean diameter of around 64 nm. Fluorescence and UV-Vis spectra of the same nanoparticles show the typical fluorescence and absorption band of RBITC indicating that silica encapsulation does not interfere with the emission properties of the fluorophore. It was also noticed that silica encapsulation improved RBITC quantum yield and fluorescence lifetime when compared to free RBITC in solution. Particle’s surface have also been modified, so that the particles could bind to biologically active molecules, such as oligonucleotides.

4.1. Material and Methods

4.1.1 Chemicals

Rhodamine b isothiocyanate (mixed isomers Aldrich), rhodamine b (dye content ~95% Sigma), (3-aminopropyl)triethoxysilane (≥98% Sigma-Aldrich), water (molecular 100 FCUP Dye doped fluorescent silica nanoparticles

biology reagent Sigma), tetraethoxysilane (99% Aldrich), Triton X-100 (Aldrich), 1- hexanol (98% Merck), cyclohexane (99% Aldrich), ammonia (25% Merck), ethanol (99.5% Panreac), acetone (99.9% Fluka), acetonitrile (99.9% ROMIL), (3- glycidyloxypropyl)trimethoxysilane (99% Aldrich), potassium phosphate monobasic (≥99% Sigma-Aldrich), potassium phosphate dibasic (≥99% Sigma-Aldrich), were used as received. Oligonucleotide sequences (5’-gat cgc ctc cac gtc c-3’) were acquired from STAB vida (Lisbon, Portugal) and were purified through a NAP-5 column from GE Healthcare (UK) before use.

4.1.2 Instrumentation and methodologies

4.1.1.1. Elemental Analysis

Elemental analysis for carbon and hydrogen were performed on a Leco CHNS-932, the technique was carried out in the University of Santiago de Compostela.

4.1.1.2. UV-visible spectroscopy

Absorption spectra were observed on a Varian Cary bio50 spectrophotometer, using quartz cells with 1 cm path length. Absorption spectra of RBITC doped silica nanoparticles were fitted using a second order exponential decay in order to remove the background scattering from silica. Based on the absorbance, the amount of RBITC molecules was calculated with an assumption that the molar absorption coefficient (ε) of RBITC molecules are equal in a RBITC solution and a RBITC doped nanoparticle solution if they have the same absorbance. Using the known values including the density of silica matrix (2.2 g mL-1), the concentration and the size of the nanoparticles, the amount of RBITC molecules in each nanoparticle was calculated.[1, 2]

4.1.1.3. Fluorescence spectroscopy, quantum yield and lifetime

Fluorescence measurements were performed in a Varian Cary Eclipse spectrofluorometer, equipped with a constant-temperature cell holder (PeltierMulticell Holder).

Fluorescence quantum yield determination was done as described by Fery-Forgues and Lavabre.[3] Absorption spectra were recorded with a Varian Cary bio50 spectrophotometer, equipped with a Varian Cary single cell Peltier accessory, using FCUP 101 Dye doped fluorescent silica nanoparticles

quartz cells with 1 cm path length, thermostated at 25 °C. Rhodamine b was used as standard for quantum yield determination. Steady-state fluorescence measurements were carried out with a Varian spectrofluorometer, model Cary Eclipse, equipped with a constant-temperature cell holder (PeltierMulticell Holder) with 5 mm slit width for excitation and emission. All emission spectra were recorded at 25 °C using the maximum λexc and the appropriate λem range for rhodamine b isothiocyanate and considering the different solvents used.

For the calculation of relative quantum yield, from scattering corrected spectra, the following equation was used:

(Eq. 4.1)

The ratio of the rhodamine b reference standard absorption (Ast) to the absorption of the sample (As), was found first and multiplied by the quantum yield of the rhodamine b solution (Φst) as well as the square ratio of the sample refractive index (ns) and the reference standard refractive index (nst). This product was then multiplied by the ratio of the integrated area under the emission spectrum of the sample (Fs) to that of the standard (Fst) to find the relative quantum yield of the particles and the free dye.

For lifetime measurements the samples were excited at 370 nm using a nanoled (IBH).The electronic start pulses were shaped in a constant fraction discriminator (Canberra 2126) and directed to a time to amplitude converter (TAC, Canberra 2145). Emission wavelength was selected by a monochromator (Oriel 77250) imaged in a fast photomultiplier (9814B Electron Tubes Inc.), the PM signal was shaped as before and delayed before entering the TAC as stop pulses. The analogue TAC signals were digitized (ADC, ND582) and stored in multichannel analyser installed in a PC (1024 channels, 1.95 ns/ch). Fluorescence lifetime values were obtained by fitting the data to appropriate decay models. These measurements were carried out by Dr. César Laia and Dr. João Lima from the group of photochemistry of the chemistry department, Faculty of Sciences and Technologies – University Nova de Lisboa.

4.1.1.4. Steady-state anisotropy

Steady-state anisotropy fluorescence emission spectra were obtained on a Jobin Yvon Spex, Fluorolog FL3-22, using quartz cells with 1 cm path length. In the emission anisotropy measurements samples were excited at 530 nm. These measurements 102 FCUP Dye doped fluorescent silica nanoparticles

were carried out in the chemistry department of the Faculty of Sciences and Technologies from the University Nova de Lisboa.

4.1.1.5. Transmission electron microscopy

TEM images were obtained using a HITACHI H-8100 instrument operating at an acceleration voltage of 200 kV. Samples for TEM analysis were prepared by depositing ethanol suspensions of the nanoparticles on carbon coated copper grids and allowing them to completely dry. TEM images were analysed using image J software (version 1.44p), available through http://imagej.nih.gov/ij. TEM was carried out in the institute of materials and surfaces science and engineering (ICEMS) from Instituto Superior Técnico (IST).

4.1.1.6. Dynamic light scattering and zeta potential

DLS measurements were performed at 25ºC, using a Malvern ZetasizerNanoZS compact scattering spectrometer with a 4.0 mW He-Ne laser (633 nm wavelength) at a scattering angle of 173º. The average hydrodynamic diameter and the size distribution of the samples were determined using Malvern Dispersion Technology Software 5.10. For zeta potential measurements NPs were redispersed in potassium phosphate buffer (10 mM, pH=8) and analysed in disposable polystyrene zeta potential cuvettes with gold-coated electrodes (Malvern) at 25.0 ºC. All measurements were repeated five times to verify the reproducibility of the results.

4.1.3 Preparation of core-shell nanoparticles with rhodamine B

isothiocyanate (RBITC-APTES@SiO2)

Nanoparticles were synthesized using the reverse microemulsion technique for the alkaline hydrolysis of tetraethoxysilane (TEOS) following a procedure described in literature.[4] For the silica nanoparticles containing rhodamine b isothiocyanate (RBITC) molecules, the dye was firstly coupled to a silane coupling agent (3-aminopropyl triethoxysilane – APTES), and the reaction product was incorporated into silica spheres by hydrolysis and polymerization of TEOS in alkaline media.

RBITC dye (1.0x10-4 mol) was prepared in aqueous solution and then mixed with an equimolar quantity of APTES, reacting overnight at room temperature. The monomer precursor RBITC-APTES was directly used to prepare silica-coated fluorescent FCUP 103 Dye doped fluorescent silica nanoparticles

nanoparticles using the reverse microemulsion technique as reported elsewhere.[5-7] The reverse microemulsion was prepared by mixing 1.77 mL of Triton X-100, 7.50 mL of cyclohexane, 1.80 mL of n-hexanol, 400 μL of water and 100 μL of the RBITC- APTES solution mentioned above. After stirring for 1 hour, 200 μL of TEOS were then added as a precursor for silica formation, followed by the addition of 100 μL of ammonia to initiate the polymerization process. The reaction was allowed to continue for 48 hours at room temperature. When it was completed the nanoparticles were isolated from the reaction media by addition of 20 mL of pure acetone. NPs were then washed by repeated cycles of centrifugation/resuspension in ethanol to remove any surfactant or unreacted molecules. After washing steps NPs were dried in a desiccator and stored for further use. The nanoparticles prepared following the described methodology are mentioned in this work as RBITC-APTES FSNPs or RBITC-

APTES@SiO2.

4.1.4 Surface functionalization of silica nanoparticles

Modification of silica NPs surface is simple, typically achieved using organosilane linkers. With appropriate linkers it is possible to bind a variety of biologically active molecules including proteins and oligonucleotides.[5, 8-10] The surface of RBITC- [11] APTES@SiO2 was modified by a grafting methodology adapted from literature. The dried RBITC FSNPs (RBITC-APTES@SiO2) (68 mg) were dissolved in acetonitrile (7 mL) and then (3-glycidyl-oxypropyl)-trimethoxysilane (GPTEs) at a concentration of 2 mmol was added. The reaction with GPTEs was completed by heating the mixture to reflux under argon for 24 h. The resulting functionalized nanoparticles were then centrifuged, washed with acetonitrile several times, and dried under vacuum for further use. The amount of organosilane grafted onto the particle surface was determined by C and H elemental analysis. The resulting functionalized NPs (RBITC-

APTES@GPTEsSiO2) contain 0.20 mmol of GPTEs per 1 g of material.

4.1.5 DNA grafting

The functionalized NPs were grafted with a 16 pair single stranded DNA oligonucleotide (5’-gat cgc ctc cac gtc c-3’) by adjusting a procedure from literature.[12] Briefly, the as prepared and functionalized dye-doped fluorescent silica nanoparticles were redispersed in potassium phosphate buffer (10 mM, pH=8) to a concentration of 1.6 mg/ml in mass and then mixed with the thiolated oligonucleotide (ratio oligo/NP = 104 FCUP Dye doped fluorescent silica nanoparticles

100). The mixture was left for incubation at room temperature for 24h under stirring. Afterwards nanoparticles were centrifuged and washed with potassium phosphate buffer for three times redispersed in buffer and storage at - 20ºC for further use.

4.2. Results and Discussion

Fluorescent silica nanoparticles (FSNPs) containing RBITC molecules were synthesized using the reverse microemulsion technique for the alkaline hydrolysis of TEOS. The dye was firstly coupled to a silane coupling agent (3-aminopropyl triethoxysilane – APTES), and the reaction product was incorporated into silica spheres by hydrolysis and polymerisation of TEOS in alkaline media. The obtained

nanoparticles (RBITC-APTES@SiO2) have a mean diameter of approximately 64 nm with a spherical morphology and a narrow particle size distribution. The particle surface was also modified, so that the particles could bind to biologically active molecules, such as oligonucleotides. Lifetime measurements and steady-state anisotropy studies of the nanoparticles and the free dye were also performed to evaluate the effect of the encapsulation on the fluorescence emission properties of RBITC.

4.2.1 Characterization of RBITC@SiO2 nanoparticles

4.2.1.1. Characterization by electron microscopy

The size and shape of RBITC-APTES doped fluorescent silica nanoparticles were measured from TEM images. Based upon the size (diameter) measurement of more than 100 particles, the average nanoparticle diameter obtained was 64.4 nm (± 7.5 nm standard deviation). TEM images (Figure 4.1) show that the dye doped fluorescent silica nanoparticles have a spherical shape, are fairly monodisperse and size distribution histogram demonstrates a narrow particle size distribution (Figure 4.1 bottom right). FCUP 105 Dye doped fluorescent silica nanoparticles

Figure 4.1 - TEM images of RBITC-APTES FSNPS nanoparticles and corresponding size distribution histogram.

Figure 4.2 - SEM images of RBITC-APTES FSNPs showing the spherical morphology of the NPs. 106 FCUP Dye doped fluorescent silica nanoparticles

Representative SEM images of the RBITC-APTES FSNPs are shown in Figure 4.2, and corroborate the results obtained by TEM showing the spherical morphology of the NPs.

4.2.1.1. Dynamic light scattering

The average diameters of RBITC-APTES FSNPs were also determined by DLS. The results obtained are presented in Table 4.1 and in Figure 4.3. It is expected to obtain larger diameters values by DLS than the ones measured by TEM since DLS measures the effective diameter of a particle in a liquid environment - the so called hydrodynamic diameter, whereas TEM measures size of individual dehydrated particles. In practice, DLS measures not only the size of the NPs but also the additional layer corresponding to the solvent that moves together with the particles on their Brownian motion. The obtained size measured by DLS can also include any other molecules attached or adsorbed to the particle’s surface such as surfactant molecules, which by TEM cannot be measured due to poor contrast of those molecules. Furthermore, DLS gives an ensemble size average of dispersed particles, which may include aggregates. For these reasons the results presented in Table 4.2 are expected and in accordance with the ones obtained from TEM analysis.

Table 4.1 - Average hydrodynamic diameter of RBITC FSNPs measured by DLS (by percentage of number of particles, measurements were repeated 5 times for each sample).

Hydrodynamic Polydispersity Standard Sample Measurement diameter (nm) index deviation (±)

1 95.83 0.238 46.75

2 102.9 0.206 46.70 RBITC-APTES 3 96.65 0.211 44.39 FSNPs 4 87.99 0.234 42.56

5 88.79 0.188 38.49

Polydispersity index refers to the variability in particle size and is equal to the square of the division product of the standard deviation / mean diameter ( )

In a non-agglomerated suspension, the hydrodynamic diameter measured by DLS will be similar or slightly larger than the TEM size. If the particles are agglomerated, the FCUP 107 Dye doped fluorescent silica nanoparticles

DLS measurement is often much larger than the TEM size and can have a large variability in the particle size (polydispersity index >0.1).

25 Size: 78.82 nm

20

15

10

5 Number of particles (%)

0 0 20 40 60 80 100 120 140 160 180 200 Hydrodynamic diameter (nm)

Figure 4.3 - DLS hydrodynamic diameter distribution statistics graph (by percentage of number of particles) for RBITC FSNPS. Error bars show standard deviation of five different measurements.

Comparing the size dispersion obtained by both techniques (DLS and TEM) it can be observed that DLS dispersion is higher relative to TEM. This is not only related to what was previously mentioned about the hydrodynamic diameter of the particles, but also to the fact that DLS gives an ensemble size average of dispersed particles, which may include aggregates and therefore giving rise to a higher polydispersity. On the other hand, since TEM can measure size of individual dehydrated particles, each particle is sized individually and therefore aggregates can be excluded reflecting a lower polydispersity compared to that of DLS. In summary TEM provides the size distribution of dehydrated particles and DLS measurements yield an ensemble average of the particle size in solution.

4.2.1.2. Characterization by UV-vis spectroscopy

Free dye and FSNPs were dissolved in ethanol with almost the same UV absorbance by adjusting the concentration. The total amount of RBITC dye per unit volume can be derived through the absorption with the assumption that the molar absorption coefficient (ε) of free dye molecules in solution and in FSNPs suspension are almost the same when they have equivalent UV absorbance value.[2] 108 FCUP Dye doped fluorescent silica nanoparticles

As the TEM characterization shows (see section 4.2.1.1) RBITC doped FSNPs had a relatively low polydispersity with an average size of 64.4 ± 7.5 nm. The average volume per FSNPs was obtained from the TEM, see Eq. (4.2).

(Eq. 4.2)

Where V is the average volume and d is the diameter determined by TEM. The density of the FSNPs (corresponding to that of amorphous silica, ca. 2.2 g/cm3), allowed the estimation of the number of FSNPs obtained at the end of the synthesis per unit volume. Since the total number of the RBITC dye molecules in the suspension is known (determined by the absorption value, in comparison with the absorption of the free dye, multiplied by Avogadro’s number, 6.022x1023), the number of the RBITC dye per FSNP can be obtained from Eq. 4.3.

(Eq. 4.3)

Where Np is the number of the RBITC dye molecules per FSNP, NRBITC is the total

number of the RBITC dye molecules in the suspension and NNPs is the number of FSNPs in the suspension.

0.10 RBITC FSNPs

0.08

0.06 Abs 0.04

0.02

0.00 350 400 450 500 550 600 650 700 Wavelength (nm)

Figure 4.4 - UV-vis spectra of RBITC and RBITC fluorescent silica NPs (FSNPs) in ethanol at 25 ºC. UV-vis spectrum of FSNPs was fitted using a 2nd order exponential decay to remove silica scattering. Both samples were dissolved to a final concentration with almost the same absorbance (0.09). FCUP 109 Dye doped fluorescent silica nanoparticles

Figure 4.4 shows RBITC and RBITC FSNPs absorption spectra in ethanol at 25 ºC with a final concentration with almost the same absorbance value of approximately 0.09. Free RBITC molecules in solution and RBITC FSNPs show a similar broad absorption band. The absorption spectrum of RBITC presents a maximum at 542 nm wavelength while the one for RBITC FSNPs is at 555 nm wavelength. This small shift to a higher wavelength could be due to the silica matrix being a less polar medium than ethanol, causing a red shift of the spectrum maximum of RBITC dye molecules.[13, 14] Similar findings were reported for silica nanoparticles encapsulating tetramethylrhodamine-dextran (TMR-Dex); tetramethylrhodamine-5-isothhiocyanate (TRICT) modified with APTES and Rubpy when compared to the free dyes in water.[13, 14]

Regarding the possible effect that conjugation of APTES to RBITC dye could have in the emission properties of the dye, the UV-vis spectrum of a RBITC-APTES conjugate solution was recorded and compared to that of the RBITC dye in solution (Figure 4.5). Both solutions were prepared in absolute ethanol with a concentration of 6.7x10-7 M and the spectra were recorded at 25 ºC. As shown in Figure 4.5 the UV-vis spectrum of RBITC-APTES conjugate presents a broad absorption band with a maximum absorbance at 542 nm similar to UV-vis spectrum of RBITC. This indicates that the RBITC emission properties remain unchanged after conjugation with APTES

0.05 RBITC RBITC-APTES

0.04

0.03 Abs 0.02

0.01

0.00 350 400 450 500 550 600 650 700 Wavelength (nm)

Figure 4.5 - UV-vis spectra of RBITC and RBITC-APTES conjugate in ethanol at 25 ºC. 110 FCUP Dye doped fluorescent silica nanoparticles

The amount of RBITC dye in FSNPs was determined quantitatively by UV-Vis spectroscopy by comparison with the free dye in solution. The quantity of RBITC dye molecules per NP is equal to the RBITC dye weight per unit volume (derived through the absorption), divided by the number of NPs per unit volume (determined by TEM). Table 4.2 shows the results obtained for the determination of the amount of RBITC dye molecules per FSNP. These determinations were made for FSNP samples with similar size (a and b from the same batch; c and d from different batches relatively to the former one), and with different mass concentrations of NPs (0.8 mg/mL and 1.6 mg/mL).

Table 4.2 – Amount of RBITC dye molecules per fluorescent silica nanoparticle

Average NP NP RBITC RBITC Size [NPs] NPs in Sample volume density molecules in molecules (nm) (g/L) suspension (cm3) (g/cm3) suspension per NP

a 64.4 1.6 5.21x1015 7.88x1017 151

b 64.4 0.8 2.60x1015 3.79x1017 146 1.40x10-16 2.2 c 68.1 1.6 4.41x1015 6.02x1017 136

d 66.9 1.6 4.65x1015 4.52x1017 97

The concentration of NPs solutions was 1.6 g/L and 0.8 g/L. The number of silica NPs was derived based on the mass of dry samples, the average volume of the particles and the density of the NPs. An assumption was made that all the mass was attributed to the silica NPs. Based on the procedure described above in this section for calculation of the amount of RBITC molecules in FSNPs, and using the known values including the density of the silica matrix (2.2 g/cm3), the concentration and the size of the NPs the amount of RBITC molecules was determined. The results indicated an estimate of about 100 to 150 RBITC dye molecules per FSNP.

4.2.1.3. Fluorescence excitation and fluorescence emission spectra

Steady-state fluorescence excitation (Figure 4.6) and fluorescence emission (Figure 4.7) spectra of RBITC, RBITC-APTES conjugate and RBITC-APTES FSNPs were recorded. All measurements were performed at 25 ºC and all experimental solutions were prepared in absolute ethanol, dissolved to a final concentration of approximately FCUP 111 Dye doped fluorescent silica nanoparticles

6.3x10-7 M. The fluorescence excitation (Figure 4.6) and emission spectra (Figure 4.7) for RBITC and RBITC-APTES conjugate shows the presence of strong peaks around 540 and 565 nm respectively, typical for RBITC molecules in ethanol.[15, 16] In case of RBITC-APTES FSNPs the fluorescence excitation (Figure 4.6) and emission spectra (Figure 4.7) also shows the presence of strong peaks around 555 and 570 nm respectively slightly shifted compared to the free dye. The excitation and emission spectra of the free and silica encapsulated RBITC dye are identical, showing that the spectral properties of RBITC when doped inside the silica NPs do not change. Furthermore the encapsulation of RBITC within the silica matrix increases the fluorescence intensity signal compared to the free dye in solution. Fluorescence excitation spectra of RBITC and RBITC-APTES FSNPs match well with their corresponding absorption spectra. Both absorption and emission spectra of RBITC fluorescent silica nanoparticles clearly confirmed successful doping of RBITC molecules into silica nanoparticles.

1000 RBITC RBITC-APTES FSNPs 800

600

400 Intensity (a.u.)

200

0 360 400 440 480 520 560 600 Wavelength (nm)

Figure 4.6 - Fluorescence excitation spectra of RBITC, RBITC-APTES conjugate and FSNPs recorded at 25 ºC in absolute ethanol. 112 FCUP Dye doped fluorescent silica nanoparticles

1000 RBITC RBITC-APTES FSNPs 800

600

(a.u.)

400 Intensity

200

0 540 560 580 600 620 640 660 680 700 Wavelength (nm)

Figure 4.7 - Fluorescence emission spectra of RBITC, RBITC-APTES conjugate and FSNPs recorded at 25 ºC in absolute ethanol.

4.2.1.4. Fluorescence quantum yield

The efficiency of the fluorescence process is measured by the quantum yield. By

definition, the fluorescence quantum yield ΦF expresses the portion of excited molecules that deactivate by emitting a fluorescent photon. It is the ratio of the number of emitted photons to the number of absorbed photons per time unit[3]:

(Eq. 4.4)

Fluorescence quantum yield values for RBITC, RBITC-APTES conjugate and RBITC-APTES FSNPs with dye adsorbed or covalently bound to the silica matrix, were determined, following a comparative method where the quantum yield of an unknown dye molecule is obtained by comparison with a dye standard molecule having a known [17] quantum yield. Rhodamine b (ΦF = 0.49 in ethanol ) was chosen as the standard dye molecule. To minimize reabsorption effects, the absorbance sample values were kept below 0.1. In Table 4.3 are presented the results obtained for fluorescence quantum yield of RBITC, RBITC-ATES conjugate and RBITC-APTES FSNPs. In order to gain an insight into the nature of the binding between the dye and the silica matrix, namely determining if the dye was physically entrapped or chemically adsorbed onto silica matrix, two kinds of particles were used and compared with RBITC FSNPs synthesized previously. The two kinds of particles used were FSNPs with dye adsorbed to the silica

NPs surface (Ads:RBITC-APTES@SiO2 FSNPs) and FSNPs with dye covalently bound FCUP 113 Dye doped fluorescent silica nanoparticles

to the silica matrix (Shell:RBITC-APTES@SiO2 FSNPs). In both cases, we expect the dye to be more accessible to the solvent than in the standard NPs, allowing us to have a control, to determine the effects, of encapsulation, and solvent proximity on optical properties of the dye. To prepare these nanoparticles the following modifications to the procedure described in section 4.1.3 were made: in case of the FSNPs with dye adsorbed these were prepared in two steps, first bare silica NPs were prepared by hydrolysis and condensation of TEOS through a microemulsion method and secondly the obtained NPs were left to react with a solution of RBITC-APTES conjugate in order to allow dye adsorption to the silica matrix. After reaction NPs were washed by repeated cycles of centrifugation/resuspension in ethanol to remove any unreacted dye molecules, and dried in a desiccator. In this way the NPs obtained were silica NPs with a layer of dye physically adsorbed to their surface (Figure 4.8 B). For the FSNPs with dye covalently bound these were synthesized in a similar way of the former ones with the difference that in the second step the bare silica NPs reacted with RBITC-APTES conjugate in a microemulsion with hydrolysis and polymerisation of TEOS in order to obtain a NP with a silicon core and a surrounding layer containing dye entrapped into silica (Figure 4.8 C). Both quantum yields of Ads:RBITC-APTES@SiO2 FSNPs and

Shell:RBITC-APTES@SiO2 FSNPs were determined and compared to those of RBITC-

APTES@SiO2 FSNPs.

Figure 4.8 - (A) RBITC doped fluorescent silica NPs prepared by hydrolysis and polymerization of TEOS in a microemulsion method; (B) bare silica NPs with RBITC dye molecules adsorbed onto the nanoparticle’s surface; (C) fluorescent core-shell NPs with a silicon core and a shell of RBITC dye molecules and TEOS.

Table 4.3 - Fluorescence quantum yields of RBITC, RBITC-APTES conjugate and FSNPs 114 FCUP Dye doped fluorescent silica nanoparticles

Sample Quantum yield (Φ)a SD

RBITC 0.24 0.03

RBITC-APTES 0.23 0.04

RBITC-APTES@SiO2 FSNPs 0.34 0.04

Ads:RBITC-APTES FSNPs 0.09 0.02

Shell:RBITC-APTES@SiO2 0.34 --- FSNPs

a calculated from eq. 4.1 using data of rhodamine b as standard.

The relative fluorescence quantum yields of RBITC and of the RBITC-APTES conjugate were found to be 0.24 (± 0.03) and 0.23 (± 0.04) respectively while that for RBITC FSNPs was 0.34 (± 0.04). The results demonstrate an increase in quantum yield achieved by encapsulating the dye within a silica NP. On the other hand the determined relative fluorescence quantum yield for the FSNPs with dye adsorbed to

the silica matrix (Ads:RBITC-APTES@SiO2) is considerably lower (0.09 ± 0.02) comparatively to the former ones. This result indicates that the fluorophore is adsorbed to the nanoparticle’s surface and is exposed to the surrounding environment that is

responsible for its photobleaching. In case of Shell:RBITC-APTES@SiO2 FSNPs, which were left to react with RBITC-APTES in the same conditions used to produce the

RBITC-APTES@SiO2 FSNPs, meaning they undergo a reaction of hydrolysis and polymerization of TEOS in the presence of the dye, the quantum yield of the NPs is

again higher than the free dye in solution and similar to RBITC-APTES@SiO2 FSNPs. This is indicative that the dye is embedded in the silica matrix and it is protected from the outer environment since within the silica matrix the -O-Si-O- network can limit the

diffusion of atmospheric O2 and solvent and thus reduce the interaction of these components with the encapsulated dye molecules.[13] The silica matrix as an enhancement effect on the fluorescence quantum yield of the encapsulated dye

molecules and protects them against solvents and atmospheric O2 that can affect and degrade the photophysical properties of the dye. Similar results were obtained for other dyes, where increases of the quantum yields were noticed after encapsulation.[18, 19] FCUP 115 Dye doped fluorescent silica nanoparticles

4.2.1.5. Lifetime measurements

Fluorescent lifetime measurements for RBITC, the RBITC-APTES conjugate,

RBITC-APTES FSNPs (dye covalent bound - RBITC-APTES@SiO2; and dye adsorbed to the silica matrix – Ads:RBITC-APTES@SiO2) were recorded to investigate dye distribution in fluorescent silica nanoparticles. Once again all measurements were performed at room temperature and all experimental solutions were prepared in absolute ethanol. Fluorescence lifetime decay curves are presented in Figure 4.9 and lifetime data are compiled in Table 4.4. As shown in Table 4.4 RBITC has a single component with one lifetime value (fluorescence decay was fitted using a single exponential decay). However, in case of RBITC-APTES conjugate and RBITC-

APTES@SiO2 FSNPs (both dye adsorbed and covalently bound to the silica matrix) the fluorescence decay could only be fitted by a 2nd exponential decay, indicating two different microenvironments around RBITC molecules (two lifetimes values see Table 4.4). The lifetime of the free dye was measured to be 2.44 ns in absolute ethanol while RBITC-APTES conjugate and the NPs exhibit two-component lifetime behaviour with a low (LC) and a high (HC) lifetime components of 1.04 ns (LC) and 2.99 (HC) for RBITC-APTES conjugate; 0.87 ns (LC) and 2.66 ns (HC) for NPs with dye adsorbed

(Ads:RBITC-APTES@SiO2) and 1.27 ns (LC) and 3.44 ns (HC) for NPs with dye covalently bound (RBITC-APTES@SiO2).

Table 4.4 - Lifetime data of RBITC and fluorescent silica nanoparticles (FSNPs) in absolute ethanol

FIT Sample 2 Τ1 (ns) Τ2 (ns) Χ

RBITC 2.44 --- 1.16

RBITC-APTES 2.99 1.04 1.20

RBITC-APTES@SiO2 3.44 1.27 1.12 FSNPs

Ads:RBITC-APTES@SiO2 2.66 0.87 1.35 FSNPs

116 FCUP Dye doped fluorescent silica nanoparticles

A 1000 B

T (RBITC-APTES) = 2.99 T(RBITC) = 2.44 1 T (RBITC-APTES) = 1.04 1000 2 100

100

Log10 I Log10 Log10 I Log10 10

10

200 400 600 800 1000 200 400 600 800 1000 Time (ns) time (ns)

C D T (RBITC-APTES@SiO ) = 3.44 1000 1 2 T (Ads:RBITC-APTES@SiO ) = 2.66 T (RBITC-APTES@SiO ) = 1.27 1 2 1000 2 2 T (Ads:RBITC-APTES@SiO ) = 0.87 2 2

100

100

Log10 I Log10 Log10 I Log10

10 10

200 400 600 800 1000 200 400 600 800 1000 Time (ns) Time (ns)

Figure 4.9 - Fluorescence lifetime decay curves of RBITC (A), RBITC-APTES conjugate (B), RBITC-APTES FSNPs with

dye covalently bound to silica matrix (RBITC-APTES@SiO2) (C),and RBITC-APTES FSNPs with dye adsorbed to silica

surface (Ads:RBITC-APTES@SiO2) (D), all at ambient temperature (298 K) .The excitation was fixed at 370 nm and the emission was monitored at 550 nm.

The presence of the two components (high and low components are designated as

τ1 and τ2 respectively) has been seen before for silica nanoparticles encapsulating other dyes (NIR664 or FTIC) and were assumed to be related with dye distribution and microenvironment within the NP.[20, 21] For instance, Santra and coworkers[21] reported that for FTIC FSNPs two lifetime components were observed and the authors correlated them with two different microenvironments associated with different solvation conditions around FTIC dye molecules. Roy et al.[20] also reported a nonhomogeneous dye distribution inside silica NPs based on fluorescence lifetime measurements. Once again the studied NPs presented two lifetime components indicating that the dye was distributed in two domains that the authors identified as being a screened core region and a more solvent-accessible region near the surface. It is know that dye lifetime can be influenced by many parameters, including dye-solvent interaction and the quenching of the dye because of the interaction of adjacent molecules.[20] However, if the presence of two different lifetime components is related with dye-solvent interactions it would be expected to observe several lifetimes FCUP 117 Dye doped fluorescent silica nanoparticles

according to the different hydration spheres of the dye molecules within the silica particle, instead of just two. For this reason in the present work it is suggested that the presence of two components can be related to another factor, the RBITC-APTES conjugation. The commercial RBITC dye used in this work is a mixture of isomers (see Figure 4.10) and for this reason the conjugation of APTES to RBITC through the isothiocyanate functional group (NCS) in the dye with the amine group (NH2) from the silane, can occur in different positions of the aromatic ring (Figure 4.11). Depending on the position of NCS group the coupling with APTES can influence dye lifetime through quenching due to the interaction of the dye with the silane adjacent molecules. In this way the smaller decay component is suggested to be associated with APTES conjugate with RBITC 5-isomer, since at this position the electronic density is higher due to the proximity with the carboxylic acid functional group and the APTES molecules, which can quench dye fluorescence by effects of charge transfer. On the other hand the higher lifetime component is suggested to be related with APTES conjugation with RBITC 6-isomer. At this position the dye may not sense to the same extent the electronic density of APTES molecules and therefore the effects of charge transfer are minor compared to the former one. In the case of RBITC-APTES@SiO2 FSNPs the two lifetimes observed (1.27 ns for the small lifetime component and 3.44 ns for the higher lifetime component) suggests that the RBITC-APTES conjugate is encapsulated within the silica matrix and shielded from the solvent molecules through the silica net since both lifetimes are higher when compared to those of the free RBITC-APTES conjugate in solution (1.04 ns LC and 2.99 ns HC).

NCS

SCN

COOH COOH

H C N O N+ CH + 3 3 H3C N O N CH3

CH CH 3 3 CH3 CH3

Figure 4.10 – Structures of rhodamine b isothiocyanate (RBITC) isomers. Left: rhodamine b 5-isothiocyanate and right: rhodamine b 6-isothiocyanate.

118 FCUP Dye doped fluorescent silica nanoparticles

NH C NH O S Si O COOH O

+ H3C N O N CH3

CH3 CH3

O

O Si O

NH C NH

S COOH

+ H3C N O N CH3

CH3 CH3

Figure 4.11 – Structures of RBITC-APTES conjugate for RBITC 5-isomer (top) and RBITC 6-isomer (bottom)

For RBITC FSNPs with dye adsorbed to the silica surface (Ads:RBITC-

APTES@SiO2) the two-component lifetime (0.87 ns LC and 2.66 ns HC) are smaller compared to free RBITC-APTES conjugate in solution. In this case the smaller and higher lifetime components are also related to the RBITC-APTES conjugation. For the longer lifetime decay (2.66 ns) suggested to be related with APTES conjugate with RBITC 6-isomer, the value is similar to the free dye in solution (2.99 ns), indicating that the adsorbed RBITC 6-isomer presents a free RBITC-APTES conjugate behaviour. Regarding the smaller lifetime decay (0.87 ns) this is suggested to be related with APTES conjugate with RBITC 5-isomer and therefore with the effects of high electronic density near dye molecules and effects of charge transfer mentioned previously. The fact that these values are smaller compared to the free RBITC-APTES conjugate could be related to the fact that the dye is adsorbed to the surface and wobbling with the solvent around the NP.[22] This means that the adsorbed RBITC-APTES conjugate have a movement restriction and when associated with a high electronic density the dye FCUP 119 Dye doped fluorescent silica nanoparticles

molecules are less flexible to avoid the effects of charge transfer which translates into a higher lifetime decrease.

The results obtained suggest that the dye is encapsulated within the silica matrix corroborating the previous results on the quantum yield. Increase on dye lifetime after silica encapsulation has also been reported by others authors in similar systems including for dyes belonging to the rhodamine family. For instance Roy[20] and coworkers have reported an increase in the fluorescence lifetime of a near-infrared dye (NIR664) conjugated with 3-methacryloyloxypropyltriethoxysilane (MPTES) after encapsulation into silica NPs with approximately 105 nm. In the particular case of silica nanoparticles incorporating rhodamine dyes Ma et al.[14] have described that for TMR- APTES (reaction product of TRITC and APTES) an increase in dye lifetime from 1.94 ns to 3.67 ns after encapsulation within silica nanoparticles was observed. Also for a system described by Larson et al.[23] consisted of a homogeneous silica NP with TRITC dye molecules sparsely embedded within the silica matrix the same increasing behaviour in dye fluorescence lifetime was reported after silica encapsulation.

4.2.1.6. Fluorescence anisotropy

Anisotropy measurements can be exploited to obtain more information about the rigidity of the environment surrounding a fluorescent probe and the extent to which this rigid environment actually prevents the motional dynamics of the former. When a fluorescent molecule is excited with polarized light the resulting fluorescence is also polarized. Fluorescence depolarization is caused by rotational diffusion of the fluorophore during the excited lifetime and so fluorescence polarization measurements can be used to determine the rotational mobility of the fluorophore. Fluorescence anisotropy (r) is an experimental measure of the fluorescence depolarization. Depolarization by rotational diffusion of spherical rotors is described by the Perrin Equation (Eq. 4.5)

(Eq. 4.5)

Where r is the measured anisotropy, r0 is the fundamental anisotropy, τ is the fluorescence lifetime and Dr is the rotational diffusion coefficient. The lower the anisotropy value, the faster the rotational diffusion therefore as the fluorophore binds to the NP the rotational diffusion should decrease and the anisotropy should increase. Since the NP interior is far more rigid than the free fluorophore environment the bound 120 FCUP Dye doped fluorescent silica nanoparticles

fluorophore molecule should undergo more restricted movement when bound inside the NP.

Figure 4.12 shows the emission fluorescence anisotropy spectra as a function of wavelength for RBITC and the FSNPs (dye covalently bound - RBITC-APTES@SiO2;

and dye adsorbed – Ads:RBITC-APTES@SiO2, to the silica matrix). Based on these

results and in the fluorescence lifetime the rotational diffusion coefficient (Dr) was calculated for the free RBITC in solution and for the FSNPs. The values obtained are presented in Table 4.5.

0.25

RBITC-APTES@SiO FSNPs 2 0.20

Ads:RBITC-APTES@SiO FSNPs 0.15 2

0.10 Anisotropy r

0.05 RBITC

0.00 580 590 600 610 620 Wavelength (nm)

Figure 4.12 - Steady state emission fluorescence anisotropy of RBITC and RBITC FSNPs with dye adsorbed

(Ads:RBITC-APTES@SiO2) and dye covalently bound (RBITC-APTES@SiO2) to silica matrix. The excitation wavelength was 530 nm.

Table 4.5 - Anisotropy (r) and rotational diffusion coefficient (Dr) values of RBITC and fluorescent silica nanoparticles (FSNPs) adsorbed and covalently bound to silica NPs in absolute ethanol.

Sample r SD Τ (ns) Dr (ns-1)

RBITC 0.025 0.005 2.44 0.876

RBITC-APTES@SiO 2 0.187 0.009 2.76 0.048 FSNPs Ads:RBITC-APTES@SiO 2 0.172 0.007 2.19 0.074 FSNPs

Fluorescence lifetimes of RBITC-APTES@SiO2 and Ads:RBITC-APTES@SiO2 FSNPs are the weighted average of the multicomponent fit.

The free dye presents an anisotropy value of 0.025, much smaller than that of the encapsulated dye (0.187) or of the fluorophore adsorbed onto the silica surface (0.172). This value for the free dye indicates that the dye in solution has a great FCUP 121 Dye doped fluorescent silica nanoparticles

mobility since there is no restriction to the molecule rotation and consequently its rotational diffusion is high (0.876 ns-1). On the other hand when the dye is encapsulated inside the silica NP the anisotropy value increases (0.187) and the dye molecules undergo more rigid movement due to the confinement inside the NP. Thus the rotational diffusion coefficient of the encapsulated dye decreases to 0.048 ns-1. The fluorescence anisotropy emission spectra (Figure 4.12) indicate a strong polarization of the fluorescence in agreement with a very low mobility of the dyes, as expected because of their inclusion in the NPs. The slower rotation of the dye molecules is in accordance with a model of restricted rotational motion — the so called wobbling-in- cone model. This model assumes that the major axis of the dye wobbles uniformly [24] within a cone of semiangle θc (Figure 4.13). These findings are in accordance to those found for other dyes such as Rubpy, TMR-Dex or TRITC-APTES conjugate in which an increase in the emission anisotropy was also observed after dye encapsulation inside silica NPs.[14, 23] A similar behaviour is observed for the NPs with the dye adsorbed to the silica surface (Ads:RBITC-APTES@SiO2). The anisotropy value for these NPs is 1.72, which is bigger than the free dye in solution but smaller than the encapsulated dye, this is owing to the fact that the dye is adsorbed and wobbling with the solvent around the NP having some restriction to the movement but less than in case of the encapsulated one. For these NPs the rotational diffusion coefficient is 0.074 ns-1.

Figure 4.13 - Representation of the wobbling-in-cone model, where θc is the angle between the probe (dye) axis (direction of the optical transition moment) and the symmetry axis of the wobbling motion (cone axis).

122 FCUP Dye doped fluorescent silica nanoparticles

4.2.2 Characterization of RBITC-APTES@SiO2 NPs grafted with DNA

The oligonucleotide-modified RBITC-APTES FSNPs nanoparticles were prepared by covalent immobilization of thiolated oligonucleotides onto the silica nanoparticles surface functionalized with an epoxy silane (3-glycidoxypropyltrimethoxy silane - GPTES). A scheme of the immobilization strategy is presented in Figure 4.14.

Figure 4.14 - Strategy for immobilisation of thiolated oligonucleotides onto dye loaded silica nanoparticle surfaces.

GPTES was chosen to functionalize the NPs surface since it is widely used in the field of microarrays in order to attach oligonucleotide probes covalently to silicon based surfaces. The epoxy groups present in the GPTES structure are known to be extremely reactive.[25] The obtained functionalized NPs (FSNPs-GPTES) contained 0.20 mmol of GPTES per 1 g of material determined by C and H elemental analysis.

To check if immobilization occurred UV-vis spectroscopy was performed after the reaction of the functionalized NPs with DNA and compared with the spectra of NPs and DNA before immobilization. UV-vis spectra of DNA, functionalized NPs (FSNPS- GPTES) and functionalized NPs after DNA immobilization (FSNPS-GPTES-DNA) are present in Figure 4.15. Particle’s surface zeta potential ζ was also measured and the obtained results are presented in Table 4.6. FCUP 123 Dye doped fluorescent silica nanoparticles

0.10

0.05 DNA

0.50 Abs

0.00 FSNPs-GPTES

FSNPs-GPTES-DNA

-0.05 300 400 500 600 700 Abs 0.25 Wavelength (nm)

0.00 FSNPs-GPTES FSNPs-GPTES-DNA 300 400 500 600 700 Wavelength (nm)

Figure 4.15 - UV-vis spectra of DNA and functionalized FSNPS before (FSNPs-GPTES) and after (FSNPs-GPTES- DNA) DNA immobilization in potassium phosphate buffer (10 mM, pH = 8). Inset: zoom in the UV-vis spectra of FSNPS before and after DNA immobilization.

From the analysis of the former UV-vis spectra no direct conclusions can be found since the region of the spectra where the DNA band appear is noisy for both FSNPs- GPTES and FSNPs-GPTES-DNA spectra. From these results it cannot be concluded whether if immobilization occurred. UV-vis spectroscopy is not the best technique to prove immobilization and unfortunately, we were not able to find a better way to prove this reaction occurred.

In Table 4.6, the average and standard deviation of the zeta potentials for FSNPs- GPTES and FSNPs-GPTES-DNA are reported. These values were calculated using the average of five separate ensemble measurements.

Table 4.6 – Zeta potential ζ of FSNPs-GPTES and FSNPs-GPTES-DNA

Sample Zeta potential ζ (mV) Standard deviation

FSNPs-GPTMS -30.68 1.80

FSNPs-GPTMS-DNA -8.31 0.69 124 FCUP Dye doped fluorescent silica nanoparticles

The zeta potential ζ of FSNPs-GPTES presented a negative surface potential of - 30.68 mV that changed to -8.31 mV after reaction with ssDNA. This difference on surface charge could indicate DNA immobilization onto silica NPs surface but further experiments, such as agarose gel electrophoresis, will be needed to actually prove that immobilization took place.

4.2.3 Conclusions

RBITC-APTES FSNPs with a mean diameter of around 64 nm were synthesized by alkaline hydrolysis and polymerization of TEOS using a reverse microemulsion methodology to control the size of the particles. TEM and SEM analyses of the obtained FSNPs show the spherical morphology of the NPs as well the narrow polydispersity also confirmed by DLS. Furthermore there is a good agreement between the hydrodynamic diameters obtained by DLS and the ones obtained by TEM.

After encapsulation the maximum emission wavelength of RBITC shifted 13 nm to a higher wavelength probably due to the nature of the silica shell, which is less polar than ethanol. However this small red shift did not change to a great extent the spectral properties of RBITC when doped inside the silica NPs. Absorption and emission spectra of RBITC fluorescent silica nanoparticles clearly confirmed successful doping of RBITC molecules into the silica matrix. Through the absorption values it was possible to determine the amount of RBITC dye molecules encapsulated per NP. The prepared RBITC-APTES FSNPs have between 100 to 150 dye molecules doped in one particle.

Although absorption, excitation and emission spectra of RBITC-APTES FSNPs show only small red-shifts when compared to the free dye in ethanolic solution, fluorescence lifetime, quantum yield and anisotropy vary significantly when RBITC dyes are encapsulated. The quantum yield of RBITC-APTES FSNPS was determined to be approximately 1.4 times higher than the quantum yield of RBITC and RBITC- APTES molecules in ethanol. In such way RBITC FSNPs have significant fluorescence intensity improvement once the silica matrix that surrounds RBITC dye molecules effectively shields them from interaction with solvent molecules. Quantum yield determinations were also used to check if dye was physically entrapped or chemically adsorbed onto silica matrix. For that purpose RBITC-APTES FSNPs were compared to that of FSNPs with bare silica core surrounded by a layer of dye (FSNPs with dye FCUP 125 Dye doped fluorescent silica nanoparticles

adsorbed) and a layer containing dye and silica (FSNPs with dye covalently bound). Results obtained showed that when the dye is adsorbed to NPs surface (Ads:RBITC-

APTES@SiO2) the quantum yield decreases in comparison to RBITC-APTES FSNPs, supporting the hypothesis that dye is adsorbed and exposed to the surrounding environment that is responsible for its photobleaching. In the other hand quantum yield results on FSNPs with dye covalently bound to the silica matrix (Shell:RBITC-

APTES@SiO2) were shown to be similar to that of RBITC-APTES FSNPs suggesting that the dye is embedded in the silica matrix and it is protected from the outer environment. Furthermore these two types of FSNPS with dye covalently bound to the silica matrix (RBITC-APTES@SiO2 and Shell:RBITC-APTES@SiO2) were optically very similar.

Fluorescence lifetime decay and steady state fluorescence anisotropy of RBITC also increase with silica encapsulation. The results obtained suggest that the dye is encapsulated within the silica matrix corroborating the previous results on the quantum yield. Furthermore dye distribution inside the NP can be classified according to the interaction of dye isomers with the adjacent APTES molecules presenting two different environments each corresponding to a small and a high lifetime component. The reaction between RBITC and APTES to produce RBITC-APTES conjugate is accomplished through the isothiocyanate functional group (NCS) in the dye with the amine group (NH2) from the silane. Depending on the position of NCS group in the isomers, the coupling with APTES can occur in different positions of the aromatic ring and thus influence dye lifetime due to the interaction of the dye with the silane adjacent molecules that can quench the dye fluorescence. The smaller and largest lifetimes are associated with APTES conjugate with RBITC 5-isomer and RBITC 6-isomer respectively. In the case of APTES conjugated with RBITC 5-isomer there is a high electronic density around the dye molecules which can quench dye fluorescence by effects of charge transfer. Regarding the conjugation of APTES with RBITC 6-isomer the electronic density around dye molecules is lower compared to the former one and therefore the effects of charge transfer are minor. Moreover increase of dye lifetime after silica encapsulation again suggests dye shielding due to the silica matrix. With respect to steady state fluorescence anisotropy the large increase in emission anisotropy from RBITC in solution (0.025) to RBITC encapsulated in silica NPS (0.187), indicates that the motion of RBITC molecules in silica is strongly restricted due to the confinement inside the NP. The motion of the encapsulated dye molecules is described by a wobbling-in-cone model. A similar behaviour is observed for the NPs with the dye 126 FCUP Dye doped fluorescent silica nanoparticles

adsorbed to the silica surface. For these NPs the dye rotational motion is similar to that of the free dye in solution, suggesting that the dye is physically bound but wobbling with the solvent around the NPs. Lifetime increase value denotes the rigid environment of RBITC dye molecules within silica NPs in accordance with the obtain steady state fluorescence anisotropy values.

The particle surfaces were also modified with an organosilane (GPTES) in order to allow the biological binding of the NPs to oligonucleotides. C and H elemental analysis

revealed that the resulting functionalized NPs (RBITC-APTES@GPTEsSiO2) contain 0.20 mmol of GPTEs per 1 g of material. The oligonucleotide-modified silica nanoparticles were prepared by covalent immobilization of thiolated oligonucleotides onto the silica nanoparticles surface. UV-vis spectroscopy was used to check if immobilization occurred but the technique seems not to be sensitive enough to prove that the binding occurred. Zeta potential measurements shown a difference in particle’s surface after reaction with DNA that could indicate immobilization, however to prove that reaction actually took place further experiments will be needed.

FCUP 127 Dye doped fluorescent silica nanoparticles

4.3. References

1. Chen, G.W., Song, F.L., Wang, X., Sun, S.G., Fan, J.L. and Peng, X.J., Bright and stable Cy3-encapsulated fluorescent silica nanoparticles with a large Stokes shift. Dyes and Pigments, 2012. 93(1-3): p. 1532-1537. 2. He, X., Chen, J., Wang, K., Qin, D. and Tan, W., Preparation of luminescent Cy5 doped core-shell SFNPs and its application as a near-infrared fluorescent marker. Talanta, 2007. 72(4): p. 1519-26. 3. Fery-Forgues, S. and Lavabre, D., Are fluorescence quantum yields so tricky to measure? A demonstration using familiar stationery products. Journal of Chemical Education, 1999. 76(9): p. 1260-1264. 4. Shi, H., He, X.X., Wang, K.M., Yuan, Y., Deng, K., Chen, J.Y. and Tan, W.H., Rhodamine B isothiocyanate doped silica-coated fluorescent nanoparticles (RBITC-DSFNPs)-based bioprobes conjugated to Annexin V for apoptosis detection and imaging. Nanomedicine-Nanotechnology Biology and Medicine, 2007. 3(4): p. 266-272. 5. Santra, S., Zhang, P., Wang, K.M., Tapec, R. and Tan, W.H., Conjugation of biomolecules with luminophore-doped silica nanoparticles for photostable biomarkers. Analytical Chemistry, 2001. 73(20): p. 4988-4993. 6. He, X., Duan, J., Wang, K., Tan, W., Lin, X. and He, C., A novel fluorescent label based on organic dye-doped silica nanoparticles for HepG liver cancer cell recognition. Journal of Nanoscience and Nanotechnology, 2004. 4(6): p. 585-9. 7. He, X.X., Wang, K.M., Tan, W.H., Li, J., Yang, X.H., Huang, S.S., Li, D. and Xiao, D., Photostable luminescent nanoparticles as biological label for cell recognition of system lupus erythematosus patients. Journal of Nanoscience and Nanotechnology, 2002. 2(3-4): p. 317-320. 8. Rosi, N.L. and Mirkin, C.A., Nanostructures in biodiagnostics. Chemical Reviews, 2005. 105(4): p. 1547-62. 9. Knopp, D., Tang, D.P. and Niessner, R., Bioanalytical applications of biomolecule-functionalized nanometer-sized doped silica particles. Analytica Chimica Acta, 2009. 647(1): p. 14-30. 10. Santra, S., Wang, K., Tapec, R. and Tan, W., Development of novel dye-doped silica nanoparticles for biomarker application. Journal of Biomedical Optics, 2001. 6(2): p. 160-6. 11. Pereira, C., Biernacki, K., Rebelo, S.L.H., Magalhaes, A.L., Carvalho, A.P., Pires, J. and Freire, C., Designing heterogeneous oxovanadium and copper 128 FCUP Dye doped fluorescent silica nanoparticles

acetylacetonate catalysts: Effect of covalent immobilisation in epoxidation and aziridination reactions. Journal of Molecular Catalysis a-Chemical, 2009. 312(1- 2): p. 53-64. 12. Mahajan, S., Sethi, D., Seth, S., Kumar, A., Kumar, P. and Gupta, K.C., Construction of Oligonucleotide Microarrays (Biochips) via Thioether Linkage for the Detection of Bacterial Meningitis. Bioconjugate Chemistry, 2009. 20(9): p. 1703-1710. 13. Liang, S., Shepard, K., Pierce, D.T. and Zhao, J.X., Effects of a nanoscale silica matrix on the fluorescence quantum yield of encapsulated dye molecules. Nanoscale, 2013. 5: p. 9365-9373. 14. Ma, D.L., Kell, A.J., Tan, S., Jakubek, Z.J. and Simard, B., Photophysical Properties of Dye-Doped Silica Nanoparticles Bearing Different Types of Dye- Silica Interactions. Journal of Physical Chemistry C, 2009. 113(36): p. 15974- 15981. 15. Ferrie, M., Pinna, N., Ravaine, S. and Vallee, R.A.L., Wavelength-dependent emission enhancement through the design of active plasmonic nanoantennas. Optics Express, 2011. 19(18): p. 17697-17712. 16. Tsou, C.J., Chu, C.Y., Hung, Y. and Mou, C.Y., A broad range fluorescent pH sensor based on hollow mesoporous silica nanoparticles, utilising the surface curvature effect. Journal of Materials Chemistry B, 2013. 1: p. 5557-5563. 17. Casey, K.G. and Quitevis, E.L., Effect of solvent polarity on nonradiative processes in xanthene dyes: Rhodamine B in normal alcohols. J. Phys. Chem., 1988. 92(23): p. 6590–6594 18. Cohen, B., Martin, C., Iyer, S.K., Wiesner, U. and Douhal, A., Single Dye Molecule Behavior in Fluorescent Core-Shell Silica Nanoparticles. Chemistry of Materials, 2012. 24(2): p. 361-372. 19. Rampazzo, E., Bonacchi, S., Montalti, M., Prodi, L. and Zaccheroni, N., Self- organizing core-shell nanostructures: Spontaneous accumulation of dye in the core of doped silica nanoparticles. Journal of the American Chemical Society, 2007. 129(46): p. 14251-14256. 20. Roy, S., Woolley, R., MacCraith, B.D. and McDonagh, C., Fluorescence lifetime analysis and fluorescence correlation spectroscopy elucidate the internal architecture of fluorescent silica nanoparticles. Langmuir, 2010. 26(17): p. 13741-6. 21. Santra, S., Liesenfeld, B., Bertolino, C., Dutta, D., Cao, Z.H., Tan, W.H., Moudgil, B.M. and Mericle, R.A., Fluorescence lifetime measurements to FCUP 129 Dye doped fluorescent silica nanoparticles

determine the core-shell nanostructure of FITC-doped silica nanoparticles: An optical approach to evaluate nanoparticle photostability. Journal of Luminescence, 2006. 117(1): p. 75-82. 22. Yip, P., Karolin, J. and Birch, D.J.S., Fluorescence anisotropy metrology of electrostatically and covalently labelled silica nanoparticles. Measurement Science & Technology, 2012. 23(8). 23. Larson, D.R., Ow, H., Vishwasrao, H.D., Heikal, A.A., Wiesner, U. and Webb, W.W., Silica nanoparticle architecture determines radiative properties of encapsulated fluorophores. Chemistry of Materials, 2008. 20(8): p. 2677-2684. 24. Kinosita, K., Jr., Ikegami, A. and Kawato, S., On the wobbling-in-cone analysis of fluorescence anisotropy decay. Biophysical Journal, 1982. 37(2): p. 461-4. 25. Escorihuela, J., Banuls, M.J., Puchades, R. and Maquieira, A., Development of oligonucleotide microarrays onto Si-based surfaces via thioether linkage mediated by UV irradiation. Bioconjugated Chemistry, 2012. 23(10): p. 2121-8. FCUP 130

FCUP 131

5. Europium polyoxometalates encapsulated into silica nanoparticles

The incorporation of europium-polyoxometalates into silica nanoparticles can lead to a biocompatible nanomaterial with suitable luminescent properties for applications in biosensors, biological probes and imaging.

Europium Keggin-type polyoxometalates Eu(PW11)x (x = 1 and 2) with different europium coordination environments were prepared using simple methodologies and no expensive reactants. These luminescent compounds were then encapsulated into silica nanoparticles for the first time through the water-in-oil microemulsion methodology with a non-ionic surfactant.

The europium-polyoxometalates and the nanoparticles were characterized using several techniques (FT-IR, FT-Raman, 31P MAS NMR, TEM-EDS, AFM, DLS and ICP analysis). The stability of the material and the integrity of the europium compounds incorporated were also examined.

Furthermore, the photo-luminescence properties of nanomaterials Eu(PW11)x@SiO2 were evaluated and compared with the free europium-polyoxometalates. The silica surface of the most stable nanoparticles was successfully functionalized with appropriate organosilanes to enable covalent binding of oligonucleotides.

5.1. Materials and Methods

5.1.1. Chemicals

Sodium tungstate ( 99 % Sigma), sodium hydrogen phosphate ( 97 % Sigma), europium chloride (99.9% Sigma), hydrochloric acid (37% Panreac), potassium chloride (> 99.5 % Merck), tetraethoxysilane (99% Aldrich), Triton X-100 (Aldrich), 1- hexanol (98% Merck), cyclohexane (99% Aldrich), ammonia (25% Merck), ethanol (99.5% Panreac), acetone (99.9% Fluka), acetonitrile (99.9% ROMIL), (3- glycidyloxypropyl)trimethoxysilane (99% Aldrich) and (3-chloropropyl)trimethoxysilane (99% Aldrich), were used as received.

132 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

5.1.2. Instrumentation and methodologies

5.1.2.1. Elemental analysis

Elemental analysis for K, W, Eu and P was performed by ICP-MS on a Varian 820- MS and C and H analysis was executed on a Leco CHNS-932, both techniques were carried out in the University of Santiago de Compostela. Hydration water contents were determined by thermogravimetric analysis performed in air between 30 °C and 700 °C, with a heating speed of 5 ºC/min, using a TGA-50 Shimadzu thermobalance. Thermogravimetry technique was carried out in the center for research in ceramics and composite materials (CICECO) associated laboratory from the University of Aveiro by Dr. Duarte Ananias.

5.1.2.2. Vibrational Spectroscopy

Fourier transform Infrared (FT-IR) absorption spectra were obtained on a Mattson 7000 FT-IR spectrometer. Spectra were collected in the 400–4000 cm-1 range, using a resolution of 4 cm-1 and 64 scans. Fourier transform Raman (FT-Raman) spectra were recorded on a RFS-100 Bruker FT spectrometer, equipped with a Nd:YAG laser with excitation wavelength at 1064 nm, with laser power set to 350 mW. The FT-Raman studies were carried out in the CICECO associated laboratory by Dr. Carlos Granadeiro

5.1.2.3. Solid state NMR

31P MAS NMR spectra were recorded for liquid solutions using a Bruker Avance III

400 spectrometer and chemical shift are given with respect to external 85% H3PO4. The 31P NMR MAS solid-state measurements were performed in a 7 T (300 MHz) AVANCE III Bruker spectrometer under a magic angle spinning of 15 KHz at room temperature. The spectra were obtained by a solid echo sequence with an echo delay of 15 microseconds, a 90 degree pulse of 10.5 microseconds at a power of 20 W and a

relaxation delay of 30 seconds. Potassium phosphate (K3PO4) was used as reference. This technique was carried out in the department of science materials (CENIMAT/I3N) of the Faculty of Sciences from University Nova de Lisboa by Dr. Gabriel Feio. FCUP 133 Europium polyoxometalates encapsulated into silica nanoparticles

5.1.2.4. Transmission electron microscopy

TEM images were obtained using a HITACHI H-8100 instrument operating at an acceleration voltage of 200 kV and energy dispersive X-ray spectroscopy (EDS) analysis was performed on a ThermoNoran spectrometer. Samples for TEM analysis were prepared by depositing ethanol suspensions of the nanoparticles on carbon coated copper grids and allowing them to completely dry. TEM images were analysed using image J software (version 1.44p), available through http://imagej.nih.gov/ij. TEM was carried out in the institute of materials and surfaces science and engineering (ICEMS) from Instituto Superior Técnico (IST).

5.1.2.5. Scanning electron microscopy

SEM analysis and EDS elemental mapping were performed on a Hitachi SU-70 instrument operating at an acceleration voltage of 30 kV. Samples for SEM analysis were prepared by depositing ethanol suspensions of the nanoparticles on carbon coated copper grids and allowing them to completely dry. Both techniques were carried out in CICECO associated laboratory by Dr. Duarte Ananias.

5.1.2.6. Dynamic light scattering

DLS measurements were performed at 25ºC, using a Malvern ZetasizerNanoZS compact scattering spectrometer with a 4.0 mW He-Ne laser (633 nm wavelength) at a scattering angle of 173º. The average hydrodynamic diameter and the size distribution of the samples were determined using Malvern Dispersion Technology Software 5.10. All measurements were repeated five times to verify the reproducibility of the results.

Stable samples were prepared by dissolving the potassium salts of Eu(PW11)x in Millipore water.

5.1.2.7. Atomic force microscopy

Samples for atomic force microscopy (AFM) were prepared by drying onto freshly cleaved mica substrates from dilute aqueous solutions. AFM measurements were made using an AFM Workshop TT-AFM instrument in vibrating (intermittent contact) mode. A small (15 µm) scanner and low gains were used to ensure high resolution. Probes from AppNano (ACT) with resonant frequency of around 300 kHz were used. Images of around 2 µm x 2µm were acquired, and analysed using Gwyddion software 134 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

and a custom routine to measure accurately the height of nanoparticles. AFM measurements were carried out by Dr. Peter Eaton.

5.1.2.8. X-ray crystallography

Crystalline K11[Eu(PW11O39)2.xH2O materials suitable for single-crystal X-ray diffraction analysis were harvested and mounted in a CryoLoop using viscous oil.[1] Diffraction data were collected at 150 K on a Bruker X8 Kappa APEX II charge-coupled device (CCD) area-detector diffractometer (Mo K graphite-monochromated radiation) using the APEX2 software[2], and equipped with an Oxford Cryosystems Series 700 cryo stream controlled by the Cryopad interface.[3] Images were processed with SAINT+ software[4], and the absorption corrections were performed by the multi-scan method implemented in SADABS.[5] The structure was solved by direct methods implemented in SHELXS-97[6, 7], allowing the immediate identification of most of the heaviest elements, namely Eu and W atoms, while the remaining atoms were positioned through successive full-matrix least squares refinement cycles on F2 using SHELXL-97.[6, 8] All atoms of the Europium-phosphotungstate anions and the K+ cations were refined using anisotropic displacement parameters, while the oxygen atoms of crystallization water molecules were refined with isotropic parameters. Although the H- atoms of the water molecules were not located from difference Fourier maps or positioned in calculated positions, they were added to the molecular formula of the compound. X-ray crystallography analysis was carried out by Dr. Luis Cunha Silva.

5.1.2.9. Photoluminescence and lifetime measurements

Emission and excitation spectra were recorded at 298 K and 14 K using a Fluorolog-2® Horiba Scientific (Model FL3-2T) spectroscope, with a modular double grating excitation spectrometer (fitted with a 1200 grooves/mm grating blazed at 330 nm) and a TRIAX 320 single emission monochromator (fitted with a 1200 grooves/mm grating blazed at 500 nm, reciprocal linear density of 2.6 nm∙mm-1), coupled to a R928 Hamamatsu photomultiplier, using the front face acquisition mode. The excitation source was a 450 W Xe arc lamp. Emission spectra were corrected for detection and optical spectral response of the spectrofluorimeter and the excitation spectra were corrected for the spectral distribution of the lamp intensity using a photodiode reference detector. Lifetime measurements were carried out using a 1934D3 phosphorimeter coupled to the Fluorolog®-3, and a Xe-Hg flash lamp (6 μs/pulse half width and 20-30 μs tail) was used as the excitation source. The variable pressure measurements were FCUP 135 Europium polyoxometalates encapsulated into silica nanoparticles

performed using a helium-closed cycle cryostat with vacuum system measuring ~5×10- 6 mbar and a Lakeshore 330 auto-tuning temperature controller with a resistance heater. The temperature can be adjusted from 14 to 450 K. Photoluminescence experiments were carried out in CICECO associated laboratory by Dr. Duarte Ananias.

5.1.2.10. Quantum efficiency

5 Based on the emission spectra, D0 life times and empirical radiative and non- 5 [9-11] radiative transition rates, the D0 quantum efficiency, q, was determined for

(Eu(PW11)2) and Eu(PW11)2@SiO2. Assuming that only non-radiative and radiative 5 processes are involved in the depopulation of the D0 state, q is given by:

k q r kr knr

where kr and knr are the radiative and non-radiative transition probabilities, respectively, –1 and kexp=τexp (kr + knr) is the experimental transition probability. The emission intensity, 5 7 I, taken as the integrated intensity S of the emission lines for the D0 F0-6 transitions, is given by:

I w A N S i j i j i j i i j

5 7 wi j where i and j represent the initial ( D0) and final ( F0-6) levels, respectively, is the A N transition energy, i j the Einstein coefficient of spontaneous emission and i the

5 [9-11] 5 7 population of the D0 emitting level. Because the D0 F5,6 transitions are not 5 observed experimentally, their influence on the depopulation of the D0 excited state may be neglected and, thus, the radiative contribution is estimated based only on the 5 7 relative intensities of the D0 F0-4 transitions. The emission integrated intensity, S, of 5 7 the D0 F0-4 transitions has been measured for compounds Eu(PW11)2 and 5 7 Eu(PW11)2@SiO2 at 298 K. Because the D0 F1 transition does not depend on the local ligand field seen by the Eu3+ ions (due to its dipolar magnetic nature) it may be 5 7 –1 [12] used as a reference for the whole spectrum, in vacuum A( D0 F1)=14.65 s , and kr is given by:

4  0 1 S0 J kr A0 1 S  0 1 J 0 0 J 136 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

5 where A0-1 is the Einstein coefficient of spontaneous emission between the D0 and the 7 F1 levels. An average index of refraction of 1.5 was considered for both samples, 5 7 –1 [13] leading to A( D0 F1) 50 s . Quantum efficiency measurements were performed by Dr. Duarte Ananias.

5.1.3. Synthesis of europium polyoxometalates Eu(PW11)x (x = 1 and 2)

The potassium salts of europium polyoxometalates K4[PW11Eu(H2O)3O39]∙4H2O

(EuPW11) and K11[Eu(PW11O39)2]∙5H2O (Eu(PW11)2), were prepared by Dr. Salete Balula and Dr. Carlos Granadeiro, following a modified procedure from the literature.[14- 16] The potassium salt of the precursor ligand (K7[PW11O39]·7H2O; PW11) was prepared [17, 18] by reported procedures. Then, a solution of EuCl3.6H2O was added dropwise to 3+ the aqueous solution of PW11. The mixture was stirred for 1 h at 90 ºC. PW11 and Eu were solubilized in the minimum amount of water, and the solutions were added in the 3+ rigorously stoichiometric amounts of PW11 and Eu to prepare PW11Eu (1:1) or

Eu(PW11)2 (1:2). Both europium polyoxometalates, as well as the precursor PW11 were characterized by elemental and thermal analysis, FT-IR, FT Raman and 31P NMR spectroscopy to check the authenticity and purity of the desired compounds.

5.1.4. Encapsulation of Eu(PW11)x (x = 1 and 2) into silica nanoparticles

The encapsulation of Eu(PW11)x in silica nanoparticles was performed by hydrolysis and polymerization of tetraethoxysilane (TEOS) with aqueous ammonia in a water-in-oil (W/O) reverse microemulsion.[19-21] Briefly, a W/O microemulsion containing Triton X- 100 (2.22 mL), 1-hexanol (1.83 mL), cyclohexane (9.31 mL), TEOS (200 µL) and a

Eu(PW11)x aqueous solution (50 mg or of Eu(PW11)x in 1 mL of H2O) was mixed with a W/O microemulsion containing Triton X-100 (2.22 mL), 1-hexanol (1.83 mL), cyclohexane (9.31 mL) and ammonia (solution at 25%, 200 µL). The mixture was stirred for 24 h at room temperature. The nanoparticles were precipitated out of the microemulsion by addition of acetone, and recovered by centrifugation. The precipitate obtained was then washed by repeated cycles of centrifugation/ressuspension in ethanol and water to remove any surfactant or unreacted molecules, and dried in a desiccator. The nanoparticles obtained were characterized by FT-IR and FT Raman spectroscopy, 31P MAS-NMR spectroscopy, ICP-MS analysis and TEM. The amount of

EuPW11 loaded in EuPW11@SiO2, as determined by ICP-MS was 44 µmol (9 wt.% of

W) and 4 µmol into Eu(PW11)2@SiO2 (2 wt.% of W), per gram of material. FCUP 137 Europium polyoxometalates encapsulated into silica nanoparticles

5.1.5. Functionalization of Eu(PW11)2@SiO2

The surface of Eu(PW11)2@SiO2 was modified by a grafting methodology described [22] by Balula et al. The dried Eu(PW11)2@SiO2 nanoparticles (68 mg) were dissolved in acetonitrile (7 ml) and then each organosilane (3-glycidyloxypropyl)-trimethoxysilane, GPTEs or (3-chloropropyl)-trimethoxysilane, CPTEs), each at a concentration of 2 mmol was added. The reaction with GPTEs was completed by refluxing the mixture under argon for 24 h. For CPTEs the mixture was treated using two different procedures: (i) under reflux and under argon for 24 h at 80 ºC and (ii) using a CEM Discover microwave device, using irradiation with a power of 100 W, at maximum pressure 100 psi, and 80 ºC for 2 h. The resulting functionalized nanoparticles were then centrifuged, washed with acetonitrile several times and dried under vacuum for further use. The amount of organosilane grafted on the particle surface was determined by C and H elemental analysis. The resulting Eu(PW11)2@GPTEsSiO2 contains 0.80 mmol of GPTEs per 1 g of material. The Eu(PW11)2@CPTEsSiO2 has 2.0 mmol and 1.3 mmol of CPTEs per 1 g of material, when prepared by the refluxing and microwave procedures, respectively.

5.2. Results and Discussion

In this work photoluminescent core/shell nanoparticles were prepared using LnPOMs as the core, and using a reverse microemulsion technique for the alkaline hydrolysis of TEOS around them. Silica encapsulation provides a protective layer around the LnPOMs molecules, reducing the interaction with the surrounded media which can adversely affect the properties of LnPOMs. Furthermore core-shell silica nanoparticles are easily functionalized and chemically stable. The LnPOMs used were 4- 11- the mono-substituted [PW11O39Eu(H2O)3] and the sandwich-type [Eu(PW11O39)2] Keggin derivatives. Both compounds were synthesized by a modified procedure that, in comparison with published procedures, is simpler, less expensive and less sensitive to pH. In addition, it was possible to determine for the first time the crystal structure of the 11- potassium salt of [Eu(PW11O39)2] . The nanocomposites obtained by encapsulation of the two Keggin derivatives exhibit a well-defined core/shell structure with an LnPOM core surrounded by a silica shell, with mean diameters of approximately 16 nm and 51 nm for the mono-substituted and sandwich-type LnPOMs nanocomposites, respectively. The nanocomposites obtained were functionalized with organosilane 138 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

linkers, namely (3-glycidyloxypropyl)- and (3-chloropropyl)-trimethoxysilanes, in order to enable their subsequent binding to biologically active molecules. Photoluminescence studies of the nanocomposites and the LnPOM salts were also performed to evaluate the effect of the encapsulation on the luminescence properties of the LnPOMs.

5.2.1. Characterization of Eu(PW11)x compounds

The major structural difference between the two compounds synthesized is the

coordination sphere of europium metal centre. In Eu(PW11)2, the europium is

coordinated to eight oxygen atoms from the two PW11 units (four oxygens from the

lacunary region of each unit), and in the case of EuPW11 the europium is coordinated to

four lacunary oxygen atoms from PW11 and three water ligands, as previously described in the literature (Figure 5.1).[14]

11− Figure 5.1 - (a) The structures of the sandwich type europium-phosphotungstate anion, [Eu(PW11O39)2] ; (b) its {EuO8} coordination center displaying a square-antiprismatic geometry and (c) the mono-substituted europium- 4- phosphotungstate anion, [PW11Eu(H2O)3O39] drawn in polyhedral and ball-and-stick mixed model.

5.2.1.1. X-ray crystallography

Information concerning crystallographic data collection and structure refinement details are summarized in Table 5.1. Crystallographic information (excluding structure factors) can be obtained free of charge via http://www.fiz- FCUP 139 Europium polyoxometalates encapsulated into silica nanoparticles

karlsruhe.de/obtaining_crystal_structure_data.html or from the Inorganic Crystal Structure Database (ICSD, FIZ Karlsruhe, Hermann-von-Helmholtz-Platz 1, Eggenstein-Leopoldshafen, 76344, Germany; phone: +49 7247808555, fax: +49 7247808259; e-mail: [email protected]), on quoting the depository number CSD - 425262.

Crystalline material of the potassium salt of Eu(PW11)2 suitable for single-crystal X- ray diffraction analysis could be grown and the crystal structure of this salt was [23] determined for the first time. Eu(PW11)2 crystallized in the monoclinic system with the respective structure solved in the P21/c space group, and revealed an asymmetric unit 11- + with one [Eu(PW11O39)2] anion, 11 K cations distributed over 14 positions and a large number of crystallization water molecules. The europium-polyoxometalate anion 3+ 7- reveals one Eu centre linked to two mono-lacunary Keggin units [PW11O39] (Figure 5.1). As reported for similar lanthano-polyoxometalate anions based compounds, 11- 3+ [Ln(PW11O39)2] , the Eu is coordinated by eight lacunary oxygen atoms belonging to [14, 24-28] two Keggin fragments leading to an eight coordinated centre, {EuO8}. The coordination environment resembles a square-antiprismatic geometry (pseudo-D4d 7- symmetry), due to the relative rotation of the two [PW11O39] moieties. In fact, the relative orientation between the two idealized squares defined by the coordination oxygen atoms is ca. 40º. The angle between the normal vectors of the oxygen-based square planes is 3.71(1)º and the interplanar distance (distance between the two square planes) is 2.592(1) Å. The closer and longer O∙∙∙O distances within the oxygen- based square planes are in the 2.7233(2)-2.9415(2) Å and 4.0339(2)-4.1220(2) Å 11- ranges, respectively. In fact, the main structural characteristics of the [Eu(PW11O39)2] anion are comparable to those already reported for related compounds.[14, 24]

In the extended crystalline arrangement, the sandwich-type europium- 11− + polyoxometalate anions [Eu(PW11O39)2] are surrounded by charge balancing K cations and a large number of water molecules, being involved in an extensive network of ionic and intermolecular (hydrogen bonds) interactions.

140 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

Table 5.1 - Crystal and structure refinement data for Eu(PW11)2

Formula EuH50K11O103P2W22

Formula weight 6387.10

Crystal description colourless needles

Crystal size (mm) 0.20 0.02 0.02

Temperature (K) 150.0(2)

Crystal system monoclinic

Space group P21/c

a (Å) 18.7852(13)

b (Å) 37.6161(3)

c (Å) 14.0578(12)

α (°) 90

β (°) 92.400(4)

γ (°) 90

Volume (Å3) 9924.9(12)

Z 4

−3 ρcalc (g cm ) 4.275

μ (mm−1) 26.614

θ range (°) 3.64 to 23.28

Index ranges -20 h 20, -41 k 35,-13 l 15

Reflections collected 72894

Independent reflections 14161 (Rint = 0.0730)

Final R indices [I>2 (I)] R1 = 0.0550, wR2 = 0.1136

Final R indices (all data) R1 = 0.0898, wR2 = 0.1264

Largest diff. peak and hole (eÅ–3) 3.615 and -2.177

5.2.1.2. Thermogravimetry

The number of crystallization water molecules was determined by thermogravimetry

(TG) of EuPW11 and Eu(PW11)2 compounds (Figure 5.2). The weight loss is compatible FCUP 141 Europium polyoxometalates encapsulated into silica nanoparticles

with the formulas K4[PW11Eu(H2O)3O39)].4H2O and K11[Eu(PW11O39)2].5H2O for EuPW11 and Eu(PW11)2 respectively.

100

95

90

85 Weight loss (%) loss Weight

80 0 200 400 600 800 Temperature ºC

Figure 5.2 - Thermogravimetric curves of EuPW11 (in blue) and Eu(PW11)2 (in red).

Both compounds exhibit one step of weight loss, but over different temperature ranges. The TG of Eu(PW11)2 shows weight loss in the range of 50 – 150 ºC attributed to the release of crystal water: 1.4% (the calculated value for 5 water molecules is

1.5%). A more extensive weight loss was found for EuPW11, 4.3% (the calculated value for 7 water molecules is 4.0%), in the range 50 - 230 ºC. The weight loss in the range 150 – 230 ºC (experimental result: 1.2%; calculated for 3 water molecules: 1.5%) is typical of coordinated water molecules whereas the remaining weight loss is due to crystallization water.

5.2.1.3. 31P NMR spectroscopy

31P NMR spectroscopy was also used to identify and to characterize the mono- substituted EuPW11 and the sandwich-type Eu(PW11)2 structures. Figure 5.3 shows the spectra of the potassium salts of both europium-polyoxometalates in D2O solution as well as the spectrum of the monovacant precursor PW11. 142 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

-10.10

PW

11 5.58

EuPW

11 0.36

Eu(PW ) 11 2

40 30 20 10 0 -10 -20 -30 -40

(ppm)

31 Figure 5.3 - P NMR spectra of monovacant precursor PW11 and Eu(PW11)x in D2O solution.

As expected, a singlet is observed for each compound at different chemical shifts: -

10.10 ppm for the PW11, 5.58 ppm for the mono-substituted EuPW11 and 0.36 ppm for

the sandwich-type Eu(PW11)2. These results are in accordance with the literature data for similar compounds[14], indicating that the distinct 1:1 and 1:2 europium- polyoxometalates were successfully prepared.

5.2.2. Characterization of Eu(PW11)x@SiO2 nanoparticles

The previously prepared europium compounds were incorporated in silica nanoparticles for the first time.[23] The encapsulation procedure was carried out by hydrolysis and polymerization of tetraethoxysilane (TEOS), in the presence of

appropriate amounts of either Eu(PW11)x using a reverse microemulsion methodology.[19-21] The preparation of materials was performed under two conditions: using the same weight amount, and using the same molar amount of the europium-

polyoxometalates. The materials obtained for EuPW11 and Eu(PW11)2 were designated

EuPW11@SiO2 and Eu(PW11)2@SiO2, respectively. Furthermore, the europium- FCUP 143 Europium polyoxometalates encapsulated into silica nanoparticles

encapsulated nanoparticles Eu(PW11)2@SiO2 were functionalized with two organosilanes: (3-glycidyloxypropyl)-trimethoxysilanes (GPTEs) and (3-chloropropyl)- trimethoxysilanes (CPTEs), by reaction of the hydroxyl groups of their surface, in a post-synthesis step.

5.2.2.1. Transmission Electron Microscopy

TEM images of the nanocomposites of silica doped with Eu(PW11)x (x = 1 or 2) show uniform nanosized spheres with a core-shell structure (Figure 5.4). As expected, the EDS analysis revealed that the imaged nanoparticles have europium tungsten and silica in their constitution (Figure 5.5). The EuPW11@SiO2 and Eu(PW11)2@SiO2 nanoparticles prepared using equal weight of europium-polyoxometalates have a mean diameter of around 16 ± 1.9 nm and 51 ± 5.8 nm, respectively (calculated from more than 100 randomly selected nanoparticles on the TEM grid).

Figure 5.4 - TEM images of (a,b) EuPW11@SiO2 and (d,e) Eu(PW11)2@SiO2 nanoparticles showing the core/shell structure (both materials prepared using 50 mg of corresponding europium compounds); (c,f) Size distribution histograms of EuPW11@SiO2 and Eu(PW11)2@SiO2 nanoparticles respectively. 144 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

500 Si Si Eu(PW ) @SiO 500 EuPW @SiO 11 2 2 11 2 C 400 400 Cu 300

300 Counts

Counts Cu 200 200 C O O 100 100 W Cu W Cu W Cu P K Cu P K EuEuEu W W W Eu Eu W W W 0 0 0.0 2.5 5.0 7.5 10.0 12.5 0.0 2.5 5.0 7.5 10.0 12.5 KeV KeV

Figure 5.5 - EDS spectra of silica nanoparticles of mono-substituted compound EuPW11@SiO2 and the sandwich-type

Eu(PW11)2@SiO2 (both materials prepared using 50 mg of corresponding europium compounds). The copper peak comes from the support grid.

To investigate if the size of the europium-polyoxometalate compound has some

influence on the final silica nanoparticle size, DLS measurement of the EuPW11 and

Eu(PW11)2 aqueous solutions were carried out. The hydrodynamic diameter found for

EuPW11 was 1.80 ± 0.04 nm and for Eu(PW11)2 was 1.70 ± 0.30 nm, which indicates

the hydrodynamic size of the EuPW11 and Eu(PW11)2 are similar and should not be the

main reason for the difference of size found for Eu(PW11)x@SiO2 particles. Analysis of AFM images (Figure 5.6) of the POMs showed features with mean heights of 1.0 ± 0.6

for EuPW11 and 1.8 ± 0.7 for Eu(PW11)2. These figures are rather more in line with the crystal structures of the POMs than the DLS data, which indicate a maximum

dimension of Eu(PW11)2 roughly double that of EuPW11. However, this data included the presence of a significant proportion of features with dimensions rather larger than expected for single POMs (i.e. larger than 2 nm), which may indicate the presence of small clusters of molecules. Nevertheless, the majority of the feature heights measured was appropriate for the diameter of single POMS (i.e. between 0.8 to 1.8 nm). The presence of larger clusters in the AFM data may be a drying artifact. Taken altogether, the size measurements suggest that the majority of seeds in the synthesis procedure may be single POM molecules, although it is likely that during subsequent silica growth a large number of secondary POMs become trapped in each nanoparticle.

FCUP 145 Europium polyoxometalates encapsulated into silica nanoparticles

Figure 5.6 - AFM topography and amplitude images respectively of EuPW11 (a,b) and Eu(PW11)2 (c,d). Topography images show the presence of features with dimensions larger than expected for single POMs (i.e. larger than 2 nm).

The formation of monodisperse colloidal nanoparticles involves two sequential steps: nucleation and growth. According to Vanblaaderen and co-workers,[35, 36] particle growth occurs through monomer addition, with the growth rate being controlled by the rate of alkoxide hydrolysis. Polydispersity and final particle size can be determined by the balance between monomer addition and nucleation.[37] Increasing the ratio of europium-polyoxometalate (nuclei) / TEOS (monomer), increases the concentration of seeds competing for the monomer in the growing process and could thus lead to a reduction of the final particle size. To clarify the relation between the nanoparticle size and the molar quantity of Eu(PW11)x present in the reaction medium, another preparation of europium-polyoxometalate nanoparticles was performed with the sandwich-type Eu(PW11)2. Silica nanoparticles were synthesized following the same reverse microemulsion methodology but using higher amount (95 mg) of Eu(PW11)2, which corresponds to the same molar amount (16 µmol) used before for the preparation of EuPW11@SiO2. In this case, the Eu(PW11)2@SiO2 nanocomposites obtained had a mean diameter of 28 nm, as determined by TEM (Figure 5.7). 146 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

Compared to Eu(PW11)2@SiO2 nanoparticles prepared using 8 µmol of Eu(PW11)2 these results suggest that the molar quantity of europium-phosphotungstate does have a relevant influence on the silica nanoparticle final size, with larger concentrations of POMs leading to greater numbers of seeds, and thus smaller final particle size.

Figure 5.7 - TEM images of (a,b) Eu(PW11)2@SiO2 NPs prepared using 95 mg (16 µmol) of corresponding europium

polyoxometalate.; (c) Size distribution histogram of the mentioned EuPW11@SiO2 NPs. For direct comparison size

distribution the histogram of EuPW11@SiO2 (d) NPs prepared using 8 µmol of the same europium polyoxometalate is also presented.

5.2.2.1. Scanning Electron Microscopy

Elemental mapping of NPs by scanning electron microscopy-energy dispersive X- ray spectrometry (SEM-EDS) was performed to evaluate the distribution of the encapsulated POMs into the NPs. Figure 5.8 and Figure 5.9 present the SEM-EDS

mapping images of EuPW11@SiO2 and Eu(PW11)2@SiO2 respectively.

The mapping identified the presence of tungsten (W) and silicon (Si) through the NPs. Although it was not possible to identify the lanthanide ion Eu3+ by EDS (Figure 5.9 b), the results show that the W (one of the major elements that constitute POMs structure) is well distributed in the samples and that it is surrounded by Si.

FCUP 147 Europium polyoxometalates encapsulated into silica nanoparticles

Figure 5.8 - (a) STEM image of EuPW11@SiO2 NPs; (b) overlapping of EDS mapping for Si (red) and W (green), (c, d) separated EDX mapping for Si and W respectively

Figure 5.9 - (a) STEM image of Eu(PW11)2@SiO2 NPs; (b) EDS spectra of Eu(PW11)2@SiO2, (c, d) separated EDS mapping for Si and W respectively. The Copper (Cu), aluminium (Al) and tin (Sn) peaks come from the support grid. 148 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

5.2.2.2. Characterization by vibrational spectroscopy

Spectroscopic methods including FT-IR, FT-Raman and solid state 31P NMR were used to characterize the nanoparticles and to analyse the integrity of the incorporated

Eu(PW11)x.

The FT-IR spectra of these nanomaterials as well as those from the corresponding

europium- polyoxometalates are presented in Figure 5.10. The spectra of the EuPW11

and Eu(PW11)2 display four characteristic strong asymmetrical vibration bands for the -1 Keggin-type frameworks: as(P-O) between 1100-1040 cm , terminal as(W-Ot) near -1 -1 950 cm , corner-sharing as(W-Ob-W) near 850 cm and edge-sharing as(W-Oc-W) near 800 cm-1.[29-32] The silica material displays its main bands in the same region as -1 the Keggin derivative compounds (400-1100 cm , Figure 5.10): as(Si-O-Si), s(Si-O- [33] Si) and (O-Si-O). Thus, most of the Eu(PW11)x bands in the nanocomposite spectra are overlapped by the strong silica bands. However, comparing the spectra of silica

and Eu(PW11)x@SiO2 it is possible to find at least one extra small band between 900 and 700 cm-1 (highlighted in Figure 5.10) which can be attributed to the edge-sharing

as(W-Oc-W) stretching modes, indicating the presence of the polyoxometalate compound.

Figure 5.10 - FT-IR spectra for EuPW11 (left) and for Eu(PW11)2 (right) and its corresponding core/shell nanoparticles with and without functionalization prepared using equal weight of europium-polyoxometalate.

FCUP 149 Europium polyoxometalates encapsulated into silica nanoparticles

A B

Eu(PW ) 11 2

Eu(PW ) @SiO 11 2 2 EuPW 11

Eu(PW ) @CPTEsSiO 11 2 2

EuPW @SiO 11 2 Eu(PW ) @GPTEsSiO 11 2 2

1800 1600 1400 1200 1000 800 600 400 1800 1600 1400 1200 1000 800 600 400 -1 -1 Wavenumber (cm ) Wavenumber (cm )

Figure 5.11 - FT-Raman spectra for EuPW11 (A) and for Eu(PW11)2 (B) and the same particles in silica-coated core:shell form, with and without functionalization (both materials prepared using 50 mg of corresponding europium compound).

The incorporation of Eu(PW11)x was confirmed by FT-Raman spectroscopy (Figure

5.11). The FT-Raman spectra of the encapsulates EuPW11@SiO2 and

Eu(PW11)2@SiO2 materials are more elucidative than the FT-IR data because this technique is extremely sensitive to the Eu(PW11)x compounds and the shell of silica does not show any significant band on the Raman region of these materials. The potassium salts of Eu(PW11)x and the Eu(PW11)x@SiO2 composites show two strong bands at 970 – 1000 cm-1 range, which are attributed to s(W-Od) at the higher and to

as(W-Od) at the lower wavenumber. Near 900 cm-1 a weaker band is observed [30, 32] corresponding to the corner-sharing as(W-Ob-W) stretches. Upon functionalization of the nanoparticles, the FT-IR and FT-Raman spectra of

Eu(PW11)2@GPTEsSiO2 and Eu(PW11)2@CPTEsSiO2 materials showed the same characteristic bands from silica and from the Keggin derivatives (Figure 5.10 and Figure 5.11 B), showing that the surface modification procedure does not significantly affect the encapsulated compounds The absence of any bands from the chemical functionalities grafted in the surface (GPTEs and CPTEs) in both materials is probably due to their low amount, as shown by the results obtained by elemental analysis of C and H (see section 5.1.5). 150 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

31 The solid state P NMR spectra of EuPW11 and Eu(PW11)2 potassium salts and corresponding silica-coated core:shell nanoparticles are shown in Figure 5.12.

A B

Eu(PW ) 11 2

EuPW 11

Eu(PW ) @SiO 11 2 2

EuPW @SiO 11 2

Eu(PW ) @CPTEsSiO 11 2 2

Eu(PW ) @GPTEsSiO 11 2 2 PW 11

70 60 50 40 30 20 10 0 -10 -20 -30 -40 -50 -60 -70 70 60 50 40 30 20 10 0 -10 -20 -30 -40 -50 -60 -70 (ppm) (ppm)

31 Figure 5.12 - Solid state P MAS NMR spectra of the monovacant precursor PW11, potassium salt EuPW11 and its

corresponding silica-coated core/shell nanoparticles (A), and of potassium salt Eu(PW11)2 and their corresponding silica- coated core/shell nanoparticles with and without functionalization (B). All nanoparticles prepared using 50 mg of corresponding europium compound.

The spectrum of EuPW11 exhibits a single peak at 0.32 ppm while the spectrum of

Eu(PW11)2 shows one signal at -3.77 with a shoulder at -4.92 ppm, which could be 7- caused by the slight asymmetry of the two [PW11O39] units that surround the europium [29, 34] ion in the sandwich compound. After encapsulation of EuPW11 into silica nanoparticles, the main peak is shifted to 1.09 ppm and two shoulders are observed at -11.13 and -14.98 ppm. This small shift could be due to the interaction between the compound and the silica, since the 31P nucleus is highly sensitive to its local environment. The two shoulders could be due to the presence of uncoordinated 7- [PW11O39] anions in different environments resulting from partial EuPW11

decomposition. This hypothesis is supported by the spectrum of the precursor PW11 (Figure 5.12 A - bottom), which contains a single peak at -14.47 ppm. On the other

hand, the spectra of the material Eu(PW11)2@SiO2 shows a single peak around -5 ppm,

slightly shifted in comparison with the potassium salt Eu(PW11)2. These results indicate

that the stability of EuPW11@SiO2 was lower than that of the Eu(PW11)2@SiO2. It appears that the europium cation was separated from the POM structure during the FCUP 151 Europium polyoxometalates encapsulated into silica nanoparticles

silica shell deposition process. For this reason, only the more stable Eu(PW11)2@SiO2 was further functionalized. Upon functionalization of Eu(PW11)2@SiO2 nanoparticles with GPTEs and CPTEs, the main peak is observed around -5 ppm with a small shoulder around -15 ppm, which may indicate that the functionalization process did not affect the structure of the Eu(PW11)2.

5.2.2.3. Photoluminescence properties

The excitation spectra of EuPW11 and Eu(PW11)2 recorded at room-temperature (298 K) (Figure 5.13) display a series of sharp lines from 355 to 550 nm, characteristics 3+ 6 7 5 5 5 of the Eu intra-4f transitions, namely F0,1 D4-1, L6 and G2-6. However at low temperature (14 K) the corresponding spectra also present a structured broad UV band ranging from 240 to 355 nm which may be attributed to a LMCT transition of the type O

W in the monovacant PW11 Keggin units. These excitation spectra, and corresponding temperature behaviour, are similar to those recently reported for the [38] related Eu(SiW11)2 compound. In particular, the two related sandwich compounds,

Eu(PW11)2 and Eu(SiW11)2, have almost identical spectra. After encapsulation into the silica nanoparticles the UV broad bands of EuPW11@SiO2 partially appear at ambient temperature and for both compounds a blue shift relative to the low temperature UV broad bands.

B A 7 5 7F 5L F L 0 6 0 6

EuPW Eu(PW )

11 11 2

4

4

2-6

2-6

D

D

5

G

5

G 5 5 LMCT LMCT 0 0 7 5

7 5 0

F

0 F

7 F D F 7 F D 0 2

F 0 2 7 7 7 5 7F 5D F D 0 1 0 1

EuPW @ SiO Eu(PW ) @ SiO 11 2 11 2 2

250 300 350 400 450 500 550 250 300 350 400 450 500 550 Wavelength (nm) Wavelength (nm)

Figure 5.13 - Excitation spectra of EuPW11 (A) and Eu(PW11)2 (B) and their corresponding core:shell nanoparticles at ambient temperature (298 K, black lines) and 14 K (red lines) while monitoring the emission at 614 nm.

The emission spectra of EuPW11 and Eu(PW11)2, and their corresponding core:shell nanoparticles, EuPW11@SiO2 and Eu(PW11)2@SiO2 recorded at ambient temperature 152 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

with and without a high vacuum (ca. 5×10-6 mbar) are shown in Figure 5.14. Lifetime measurements of the mentioned compounds and corresponding nanoparticles were also performed and are presented in Figure 5.15.

B 5D 7F 0 2 A 5D 7F 0 2

5D 7F 0 1 Eu(PW ) 5 7 11 2 5D 7F D F EuPW 0 4 0 1 11 5D 7F 0 4 5 7 D F 5D 7F 5 7 0 0 0 3 D F 5D 7F 0 0 0 3

Eu(PW ) @ SiO EuPW @ SiO 11 2 2 11 2

580 600 620 640 660 680 700 580 600 620 640 660 680 700 Wavelength (nm) Wavelength (nm)

Figure 5.14 - Ambient temperature (298 K) emission spectra of EuPW11 (A) and Eu(PW11)2 (B) and their corresponding core:shell nanoparticles at ambient conditions (black lines, pressure of 1 bar) and with a high vacuum (red lines, pressure of ca. 5×10-6 mbar). The excitation was fixed at 394 nm.

The sharp lines are assigned to transitions between the first excited non- 5 7 3+ degenerate D0 state and the F0-4 levels of the fundamental Eu septet. Except for 5 7 3+ D0 F1, which has a predominant magnetic-dipole character independent of the Eu crystal site, the observed transitions are mainly of electric-dipole nature. As happened

with the excitation mode, the emission spectra of Eu(PW11)2 and their behavior with and [38] without high vacuum was very similar to that of Eu(SiW11)2. In particular the splitting 5 7 of the D0 F1 transition into three Stark components increased greatly when the 7 sample was exposed to the high vacuum. For this compound, the splitting of the F0,1 levels into one and three Stark components, respective, the predominance of the 5 7 5 7 D0 F2 transition relative to the D0 F1 and the measurement of a single lifetime (Figure 5.15 B) unequivocally indicates the presence of a single Eu3+ environment as could be expected from their crystal structure determination. The changes observed for

the emission spectra of Eu(PW11)2@SiO2 show the influence of the encapsulation by silica on the Eu3+ emission properties. The silica shell has the ability to protect the

Eu(PW11)2 units from the effects of their environment, and thus the effect of the high vacuum on the core/shell compounds was almost eliminated. This is clear proof of the successful encapsulation of the two europium phosphotungstates into the silica nanoparticles. Moreover, the emission spectra for a single Eu3+ is extremely sensitive to small modifications in the first Eu3+coordination sphere, such as the variation of the FCUP 153 Europium polyoxometalates encapsulated into silica nanoparticles

number and type of coordinated moieties. For instance, the ratio between the

5 7 5 7 integrated intensities of the D0→ F2 and D0→ F1 transitions, I 5 7 I 5 7 , also ( D0 F2 ) ( D0 F1 ) known as the asymmetric ratio (R), is 1.52, 1.60, and 2.20 for Eu(PW11)2 at ambient pressure and at high vacuum and for Eu(PW11)2@SiO2 at ambient pressure, respectively. These values, typical of relatively symmetrical local environments, suggest a slightly more distorted environment of the Eu3+ coordination in vacuum and, to a greater extent, with the encapsulation process (a smaller value indicates a lower distortion of Eu3+ local environment, approaching to the ideal case of an inversion 5 7 centre for which the D0 F2 transition is absent). Thus, the observed changes in the room-temperature emission spectrum with the application of a high vacuum of ca. -6 5×10 mbar (Figure 5.14) clearly indicates a structural change on the Eu(PW11)2 compound probably due to the release of some solvent water molecules, and to the connection of the silanol groups around the phosphotungstate unities. This can also explain the slight decrease of the Eu3+ emission lifetime from 2.46 ± 0.02 ms

(Eu(PW11)2) to 2.30 ± 0.02 ms (Eu(PW11)2@SiO2), under ambient conditions (Figure

5.15 B). The effect of the encapsulation into the silica of the EuPW11 is similar to the one discussed above for Eu(PW11)2 and Eu(PW11)2@SiO2. However, the EuPW11 compound is more influenced by the application of the high vacuum (see the top of Figure 5.14 B), probably due to the partial or complete release of coordinated water molecules. In addition, lifetime measurements both for EuPW11 and EuPW11@SiO2 are only appropriately fitted using a bi-exponential function (Figure 5.15 A), which clearly demonstrates the presence of at least an impurity phase, probably due to the presence of a small amount of monovacant PW11 precursor.

EuPW S = 0.33 ± 0.01 ms 11 A EuPW L = 2.22 ± 0.03 ms 11

EuPW @ SiO S = 0.28 ± 0.01 ms 11 2 EuPW @ SiO L = 2.39 ± 0.07 ms

11 2

LnI

0 5 10 15 20 25 Time (ms) 154 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

B

Eu(PW ) = 2.46 ± 0.02 ms 11 2 Eu(PW ) @ SiO = 2.30 ± 0.02 ms

11 2 2

LnI

0 5 10 15 20 25 Time (ms)

3+ 5 Figure 5.15 - Eu D0 decay curves of EuPW11 (A) and Eu(PW11)2 (B) and its corresponding core/shell nanoparticles at ambient temperature (298 K) and pressure (1 bar). The excitation was fixed at 394 nm and the emission was monitored at ca. 614 nm.

5 Based on the emission spectra, D0 lifetimes and empirical radiative and non- radiative transition rates, and assuming that only non-radiative and radiative processes 5 5 are involved in the depopulation of the D0 state, the D0 quantum efficiency, q was

determined for Eu(PW11)2 and Eu(PW11)2@SiO2 (Table 5.2) following the methodology presented by Wu et al.[39] Note that these calculations assume the presence of a single 3+ Eu environment and thus are not strictly applicable to the EuPW11 based compounds.

The results demonstrate that, the relative high quantum efficiency of Eu(PW11)2 increases from 49% to 55% with the incorporation of the POM complex into the silica nanoparticles, mostly due to an increase of the radiative transition rate.

5 5 Table 5.2 - Experimental D0 lifetime, τ, radiative, kr, and non-radiative, knr, transition rates and D0 quantum efficiency,

q, for compounds Eu(PW11)2 and Eu(PW11)2@SiO2. The data have been obtained at room temperature (296 K).

-1 -1 Compound τ [ms] kr [s ] Knr [s ] q [%]

Eu(PW11)2 2.46±0.02 200 207 49

Eu(PW11)2@SiO2 2.30±0.02 241 194 55

FCUP 155 Europium polyoxometalates encapsulated into silica nanoparticles

5.2.3. Conclusions

Potassium salts of europium-polyoxometalates with distinct europium coordination

(Eu(PW11)x, x = 1 and 2) were successfully encapsulated in silica nanoparticles using a reverse microemulsion methodology. The distinct coordination of europium in

Eu(PW11)x compounds seems to influence the stability of the polyoxometalate structure during the silica nanoparticle preparation. By 31P solid NMR it was possible to confirm that Eu(PW11)2 containing europium coordinated with two units of monovacant precursor, yields Eu(PW11)2@SiO2 nanoparticles where the polyoxometalate maintains its structural integrity. On the other hand, the EuPW11 structure having labile water ligands in the europium coordination sphere, seemed to be less stable during the process of encapsulation in the silica shell. In fact, the analysis by 31P NMR suggests a partial structural decomposition of EuPW11 into PW11 may have occurred during

EuPW11@SiO2 preparation. Uniform nanosized spheres with core-shell structure with

51 and 28 nm was observed for Eu(PW11)2@SiO2 using 8 and 16 µmol, respectively.

EuPW11@SiO2 with uniform size of 16 nm were obtained using 16 µmol of compound. The amount of polyoxometalate used in the nanocomposites preparation procedure appears to have a strong influence on the nanoparticle final size, typical of a seed [35, 36] mediated growth mechanism. Eu(PW11)2@SiO2 nanoparticles were successfully functionalized using different functional groups suitable for further immobilization of biomolecules to design novel biosensors.

The variable pressure Eu3+ photoluminescence studies clearly demonstrated the encapsulation of the europium-polyoxometalates compounds and the effect of the capping silica shell on the protection from the outer environment. The photoluminescence properties of the silica encapsulated europium compounds at ambient conditions are very similar to the ones of the uncapped compounds. However, contrary to the former compounds, their photoluminescence are not significantly changed with application of extreme environment conditions such as high vacuum. The calculations performed on the Eu(PW11)2 and Eu(PW11)2@SiO2 demonstrated an 5 improvement of their D0 quantum efficiency for the capped silica compound.

156 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

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1. Kottke, T. and Stalke, D., Crystal Handling at Low-Temperatures. Journal of Applied Crystallography, 1993. 26: p. 615-619. 2. APEX2, Data Collection Software Version 2.1-RC13, Bruker AXS. 2006: Delft, The Netherlands. 3. Cryopad, Remote monitoring and control, Version 1.451. 2006, Oxford Cryosystems: Oxford, United Kingdom. 4. SAINT+, Data Integration Engine v. 7.23a ©. 1997-2005, Bruker AXS: Madison, Wisconsin, USA. 5. Sheldrick, G.M., SADABS v.2.01, Bruker/Siemens Area Detector Absorption Correction Program. 1998, Bruker AXS: Madison, Wisconsin, USA. 6. Sheldrick, G.M., A short history of SHELX. Acta Crystallogr A, 2008. 64(Pt 1): p. 112-22. 7. Sheldrick, G.M., SHELXS-97, Program for Crystal Structure Solution. 1997: University of Göttingen. 8. Sheldrick, G.M., SHELXL-97, Program for Crystal Structure Refinement. 1997: University of Göttingen. 9. Carlos, L.D., Messaddeq, Y., Brito, H.F., Ferreira, R.A.S., Bermudez, V.D. and Ribeiro, S.J.L., Full-color phosphors from europium(III)-based organosilicates. Advanced Materials, 2000. 12(8): p. 594-598. 10. Malta, O.L., Brito, H.F., Menezes, J.F.S., Silva, F.R.G.E., Alves, S., Farias, F.S. and de Andrade, A.V.M., Spectroscopic properties of a new light-converting device Eu(thenoyltrifluoroacetonate)(3) 2(dibenzyl sulfoxide). A theoretical analysis based on structural data obtained from a sparkle model. Journal of Luminescence, 1997. 75(3): p. 255-268. 11. Malta, O.L., dos Santos, M.A.C., Thompson, L.C. and Ito, N.K., Intensity parameters of 4f-4f transitions in the Eu(dipivaloylmethanate)(3) 1,10- phenanthroline complex. Journal of Luminescence, 1996. 69(2): p. 77-84. 12. Werts, M.H.V., Jukes, R.T.F. and Verhoeven, J.W., The emission spectrum and the radiative lifetime of Eu3+ in luminescent lanthanide complexes. Physical Chemistry Chemical Physics, 2002. 4(9): p. 1542-1548. 13. Hazenkamp, M.F. and Blasse, G., Rare-Earth Ions Adsorbed onto Porous- Glass - Luminescence as a Characterizing Tool. Chemistry of Materials, 1990. 2(2): p. 105-110. FCUP 157 Europium polyoxometalates encapsulated into silica nanoparticles

14. Zhang, C., Howell, R.C., Scotland, K.B., Perez, F.G., Todaro, L. and Francesconi, L.C., Aqueous speciation studies of europium(III) phosphotungstate. Inorganic Chemistry, 2004. 43(24): p. 7691-7701. 15. Griffith, W.P., Moreea, R.G.H. and Nogueira, H.I.S., Lanthanide complexes as oxidation catalysts for alcohols and alkenes. Polyhedron, 1996. 15(20): p. 3493- 3500. 16. Haraguchi, N., Okaue, Y., Isobe, T. and Matsuda, Y., Stabilization of Tetravalent Cerium Upon Coordination of Unsaturated Heteropolytungstate Anions. Inorganic Chemistry, 1994. 33(6): p. 1015-1020. 17. Brevard, C., Schimpf, R., Tourne, G. and Tourne, C.M., W-183 Nmr - a Complete and Unequivocal Assignment of the Tungsten-Tungsten Connectivities in Heteropolytungstates Via Two-Dimensional W-183 Nmr Techniques. Journal of the American Chemical Society, 1983. 105(24): p. 7059- 7063. 18. Tézé, A. and Hervé, G., α-, β-, and γ-Dodecatungstosilicic Acids: Isomers and Related Lacunary Compounds. Inorganic syntheses, 1992. 27: p. 85-96. 19. Granadeiro, C.M., Ferreira, R.A.S., Soares-Santos, P.C.R., Carlos, L.D., Trindade, T. and Nogueira, H.I.S., Lanthanopolyoxotungstates in silica nanoparticles: multi-wavelength photoluminescent core/shell materials. Journal of Materials Chemistry, 2010. 20(16): p. 3313-3318. 20. Ye, Z.Q., Tan, M.Q., Wang, G.L. and Yuan, J.L., Novel fluorescent europium chelate-doped silica nanoparticles: preparation, characterization and time- resolved fluorometric application. Journal of Materials Chemistry, 2004. 14(5): p. 851-856. 21. Balula, M.S.S., Nogueira, H.I.S. and Cavaleiro, A.M.V., New polyoxotungstates with Ln(III) and Co(II) and their immobilization in silica particles, in Advanced Materials Forum Iii, Pts 1 and 2, P. Vilarinho, M., Editor. 2006, Trans Tech Publications Ltd: Zurich-Uetikon. p. 1206-1210. 22. Balula, S.S., Santos, I.C.M.S., Cunha-Silva, L., Carvalho, A.P., Pires, J., Freire, C., Cavaleiro, J.A.S., de Castro, B. and Cavaleiro, A.M.V., Phosphotungstates as catalysts for monoterpenes oxidation: Homo- and heterogeneous performance. Catalysis Today, 2013. 203: p. 95-102. 23. Neves, C.S., Granadeiro, C.M., Cunha-Silva, L., Ananias, D., Gago, S., Feio, G., Carvalho, P.A., Eaton, P., Balula, S.S. and Pereira, E., Europium Polyoxometalates Encapsulated in Silica Nanoparticles Characterization and 158 FCUP Europium polyoxometalates encapsulated into silica nanoparticles

Photoluminescence Studies. European Journal of Inorganic Chemistry, 2013(16): p. 2877-2886. 24. Cao, J., Liu, S., Cao, R., Xie, L., Ren, Y., Gao, C. and Xu, L., Organic-inorganic hybrids assembled by bis(undecatungstophosphate) lanthanates and dinuclear copper(II)-oxalate complexes. Dalton Transactions, 2008(1): p. 115-120. 25. Du, D.Y., Qin, J.S., Li, S.L., Wang, X.L., Yang, G.S., Li, Y.G., Shao, K.Z. and Su, Z.M., A series of inorganic-organic hybrid compounds constructed from bis(undecatungstophosphate) lanthanates and copper-organic units. Inorganica Chimica Acta, 2010. 363(14): p. 3823-3831. 26. Li, B., Zhao, J.W., Zheng, S.T. and Yang, G.Y., Two Novel 1-D Organic- Inorganic Composite Phosphotungstates Constructed from [Ln(alpha- PW11O39)(2)](11-) Units and [Cu(en)(2)](2+) Bridges (Ln = Ce-III/Er-III). Journal of Cluster Science, 2009. 20(3): p. 503-513. 27. Naruke, H., Iijima, J. and Sanji, T., Enantioselective Resolutions and Circular Dichroism Studies of Lanthanide-Containing Keggin-Type [Ln(PW11O39)(2)](11-) Polyoxometalates. Inorganic Chemistry, 2011. 50(16): p. 7535-7539. 28. Niu, J.Y., Zhang, S.W., Chen, H.N., Zhao, J.W., Ma, P.T. and Wang, J.P., 1-D, 2-D, and 3-D Organic-Inorganic Hybrids Assembled from Keggin-type Polyoxometalates and 3d-4f Heterometals. Crystal Growth & Design, 2011. 11(9): p. 3769-3777. 29. Sousa, F.L., Pillinger, M., Ferreira, R.A.S., Granadeiro, C.M., Cavaleiro, A.M.V., Rocha, J., Carlos, L.D., Trindade, T. and Nogueira, H.I.S., Luminescent polyoxotungstoeuropate anion-pillared layered double hydroxides. European Journal of Inorganic Chemistry, 2006(4): p. 726-734. 30. Rocchicciolideltcheff, C., Fournier, M., Franck, R. and Thouvenot, R., Vibrational Investigations of Polyoxometalates .2. Evidence for Anion Anion Interactions in Molybdenum(Vi) and Tungsten(Vi) Compounds Related to the Keggin Structure. Inorganic Chemistry, 1983. 22(2): p. 207-216. 31. Niu, J.Y., Wang, K.H., Chen, H.N., Zhao, J.W., Ma, P.T., Wang, J.P., Li, M.X., Bai, Y. and Dang, D.B., Assembly Chemistry between Lanthanide Cations and Monovacant Keggin Polyoxotungstates: Two Types of Lanthanide Substituted Phosphotungstates [{(alpha-PW11O39H)Ln(H2O)(3)}(2)](6-) and [{(alpha- PW11O39)Ln(H2O)(eta(2),mu-1,1)-CH3COO}(2)](10-). Crystal Growth & Design, 2009. 9(10): p. 4362-4372. FCUP 159 Europium polyoxometalates encapsulated into silica nanoparticles

32. Iijima, J. and Naruke, H., Structural characterization of Keggin sandwich-type [Ln(III)(alpha-PW11O39)(2)](11-) (Ln = La and Ce) anion containing a pseudo- cubic Ln(III)O(8) center. Inorganica Chimica Acta, 2011. 379(1): p. 95-99. 33. Luo, X.J. and Yang, C., Photochromic ordered mesoporous hybrid materials based on covalently grafted polyoxometalates. Physical Chemistry Chemical Physics, 2011. 13(17): p. 7892-7902. 34. Griffith, W.P., Morley-Smith, N., Nogueira, H.I.S., Shoair, A.G.F., Suriaatmaja, M., White, A.J.P. and Williams, D.J., Studies on polyoxo and polyperoxo- metalates - Part 7. Lanthano- and thoriopolyoxotungstates as catalytic oxidants with H2O2 and the X-ray crystal structure of Na-8[ThW10O36]center dot 28H(2)O. Journal of Organometallic Chemistry, 2000. 607(1-2): p. 146-155. 35. Vanblaaderen, A., Vangeest, J. and Vrij, A., Monodisperse Colloidal Silica Spheres from Tetraalkoxysilanes - Particle Formation and Growth-Mechanism. Journal of Colloid and Interface Science, 1992. 154(2): p. 481-501. 36. Vanblaaderen, A. and Kentgens, A.P.M., Particle Morphology and Chemical Microstructure of Colloidal Silica Spheres Made from Alkoxysilanes. Journal of Non-Crystalline Solids, 1992. 149(3): p. 161-178. 37. Guo, J.J., Lu, X.H., Cheng, Y.C., Li, Y., Xu, G.J. and Cui, P., Size-controllable synthesis of monodispersed colloidal silica nanoparticles via hydrolysis of elemental silicon. Journal of Colloid and Interface Science, 2008. 326(1): p. 138-142. 38. Juliao, D., Fernandes, D.M., Cunha-Silva, L., Ananias, D., Balula, S.S. and Freire, C., Sandwich lanthano-silicotungstates: Structure, electrochemistry and photoluminescence properties. Polyhedron, 2013. 52: p. 308-314. 39. Qi, W., Li, H.L. and Wu, L.X., A novel, luminescent, silica-sol-gel hybrid based on surfactant-encapsulated polyoxometalates. Advanced Materials, 2007. 19(15): p. 1983-1987. FCUP 160

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6. Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

Silica-based materials including silica nanoparticles have attracted attention for biological applications due to their unique characteristics. Silica nanoparticles are mechanically robust, stable and transparent and they can protect and stabilize the embedded fluorophores.[1] Furthermore, silica is considered to be a biocompatible material and it has been used to enhance the biocompatibility of various materials such as QDs, gold or iron oxide nanoparticles.[2]

According to Williams[3], biocompatibility refers to the ability of a biomaterial to perform its desired function with respect to a medical therapy, without eliciting any undesirable local or systemic effects in the recipient or beneficiary of that therapy, but generating the most appropriate beneficial cellular or tissue response in that specific situation, and optimising the clinically relevant performance of that therapy. In this context, toxicity of nanoparticles refers to the ability of the particles to adversely affect the normal physiology as well as to directly interrupt the normal structure of organs and tissues of humans and animals.[4]

There is increasing interest and applicability in the biomedical and pharmacological fields in the application of silica nanoparticles. Therefore, there is a necessity to investigate the influence of silica nanoparticles on cells as their uptake implies a close contact between nanoparticles and cells when they are used in biological systems.

Studies of cytotoxicity and genotoxicity of silica nanoparticles have been performed by looking at the integrity of DNA[5-7], cell proliferation rate and cell death[7-9] after exposure to silica nanoparticles. However the question about the possible toxicity of these nanomaterials has not been fully answered since there are studies highlighting the biocompatible nature of silica[10, 11] while others demonstrate significant toxic effects caused by silica nanoparticles, namely induction of reactive oxygen species (ROS)[7, 12, 13], hepatotoxicity[14, 15] or inflammation[16]. Therefore, a detailed evaluation of the cytotoxic potential of silica nanoparticles is required.

This chapter reports cytotoxicity studies of the fluorescent silica nanoparticles synthesized in this work conducted in three different human cell models. Nanoparticle cytotoxicity was evaluated by assessing cell viability using the Calcein-AM assay. 162 FCUP Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

Phase contrast microscopy was used to evaluate cell morphology and integrity after exposure to the nanoparticles. For the fluorescent silica nanoparticles encapsulating the organic dye RBITC, a cellular uptake experiment was also performed.

6.1. Cytotoxicity assays

Cytotoxicity is the degree to which an agent has specific destructive action on certain cells. When exposed to a cytotoxic compound cells can respond in different ways, for example they may undergo necrosis, in which they lose membrane integrity and die rapidly as a result of cell lysis (breaking down of a cell). Cells can also stop growing and start dividing, or they can undergo apoptosis (programmed cell death).

Cytotoxicity assays are widely used in research not only to screen for cytotoxicity of newly developed compounds but also to help understanding the normal and abnormal biological processes that control cell growth, division, and death. The most common endpoint to measure cell viability and cytotoxic effects is by assessing cell membrane integrity. Compounds that have cytotoxic effects often compromise cell membrane integrity. Cell viability can be evaluated by using vital dyes, by protease biomarkers, with MTT or MTS redox potential assays, or by measuring ATP content. Vital dyes (dyes capable of penetrating living cells and not inducing immediate evident degenerative changes) such as trypan blue or propidium bromide are commonly used to evaluate cell membrane integrity. These dyes are normally excluded from the inside of healthy cells but when cell membrane has been compromised they freely cross the membrane and stain intracellular components.[17]

Protease biomarker assays are used to provide relative numbers of live and dead cells within the same cell population. The assay is based on measurement of a conserved and constitutive protease activity within live cells and therefore serves as a biomarker of cell viability. When cells have a healthy cell membrane they present active live-cell protease, but once the cell loses its membrane integrity the live-cell protease becomes inactive and leakage into the surrounding culture medium. Live-cell protease substrate can cross the cell membrane while dead-cell protease substrate cannot and therefore can only be measured in culture media after cells have lost their membrane integrity.[18]

Cytotoxicity can also be monitored using an assay utilizing 3-(4, 5-dimethyl-2- thiazolyl)-2, 5-diphenyl-2H-tetrazolium bromide - the so called MTT assay. MTT is a FCUP 163 Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

colorimetric assay for accessing cell viability that measures the reducing potential of the cell. In living cells MTT, a yellow tetrazolium dye, is reduced to its insoluble formazan product which has a purple colour.[19] An alternative to the colorimetric assay is a luminescent assay based on Adenosine TriPhosphate (ATP) for the quantitative evaluation of cell proliferation and cytotoxicity.[17] ATP is a marker for cell viability because it is present in all metabolically active cells and the concentration declines very rapidly when the cells undergo necrosis or apoptosis. The ATP assay system is based on the production of light caused by the reaction of ATP with added luciferase and D-luciferin being the emitted light proportional to the ATP concentration.

Alternatively to the described assays, cytotoxicity can be assessed by the calcein AM assay. This assay provides a simple, rapid and accurate method to measure cell viability and/or cytotoxicity. Calcein-AM is a non-fluorescent, hydrophobic compound that easily permeates intact live cells. In live cells the non-fluorescent calcein-AM is hydrolysed by intracellular esterases and converted to calcein, a strongly green fluorescent compound that is retained in the cell cytoplasm.

In the present work the calcein AM assay was the chosen methodology to evaluate cell viability after treatment of the three cellular lines with the fluorescent silica nanoparticles.

6.2. Materials and Methods

Studies of cell viability, cellular uptake and phase contrast microscopy were performed in the laboratory of toxicology of the Faculty of Pharmacy from University of Porto. All measurements were carried out by Dr. Sónia Fraga.

6.2.1. Chemicals

Human neuroblastoma SH-SY5Y cells (ACC 209) and human intestinal epithelial Caco-2 cells (ACC 169) were obtained from the German Collection of Microorganisms and Cell Cultures (DSMZ). Human hepatoma Hepa RG cells were obtained from Life Technologies. Antibiotic antimycotic solution (100x) stabilized with 10,000 units penicillin, 10 mg streptomycin and 25 µg amphotericin B per mL (Sigma), 0,25% trypsin-EDTA solution (Sigma), Calcein-AM (Sigma-Aldrich), 96-well plates (BD, Biosciences), were used as received. 164 FCUP Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

6.2.2. Cellular culture

SH-SY5Y and Caco-2 cells were grown in Dulbecco’s Modified Eagle’s Medium (DMEM), 4.5 g/L glucose, 25 mM sodium bicarbonate, 25 mM HEPES, supplemented with 10% fetal bovine serum (FBS), 100 U/mL penicillin, 100 μg/mL streptomycin, 0.25

μg/mL amphotericin B, and maintained at 37ºC in a 5% CO2–95% air atmosphere. Hepa RG were grown in William’s E medium, 2g/L glucose, 25 mM sodium bicarbonate, 25 mM HEPES, supplemented with 10% fetal bovine serum (FBS), 50 µM hydrocortisone hemissucinate, 5 µg/mL bovine insulin 100 U/mL penicillin, 100 μg/mL

streptomycin, 0.25 μg/mL amphotericin B, and maintained at 37ºC in a 5% CO2–95% air atmosphere. Cultures were passage at approximately 80% confluence using a 0.05% trypsin/0.53 mM EDTA solution to a maximum of 10 passages. For the viability assay, the cells were plated in 96-well plates at density of 2.5x104 cells/well for SH- SY5Y, 2x104 cells/well for Caco-2 and 1x106 cells/well for HEPA RG.

6.2.3. Nanoparticle uptake

The cultured cells were incubated with increasing concentrations of the nanoparticles (0.1-3.2 µg/mL; 200 µL/well), freshly prepared by direct dilution in serum-

and phenol red-free culture media, for 24 and 48 h at 37 ºC in a 5% CO2–humidified environment. After incubation culture media was carefully discarded and 150 µL of Hank’s balanced salt solution (HBSS with Ca2+/Mg2+) was added. Afterwards fluorescence emission was measured in a microplate reader (Synergy HT, BioTek) with an excitation and emission wavelength of 530 ± 25 nm and 530 ± 25 nm respectively.

6.2.4. Cell viability by Calcein-AM assay

The effect of silica nanoparticles on cell viability was determined by calcein-AM assay. The procedure is briefly described here. The cultured cells were incubated with increasing concentrations of the compounds of interest (0.1-3.2 µg/mL; 200 µL/well) for

24 and 48 h at 37 ºC in a 5% CO2 humidified environment. After incubation culture media was carefully discarded and 150 µL of calcein-AM 1 µM was added to each well and incubate for 60 min at room temperature. Then the calcein-AM was carefully discarded and 150 µL of HBSS (with Ca2+/Mg2+) was added to each well. Afterwards fluorescence emission was measured in a microplate reader (Synergy HT, BioTek) with excitation and emission wavelengths of 485 ± 10 nm and 530 ± 12.5 nm respectively. Results are presented as percentage of vehicle-treated control cells. FCUP 165 Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

6.2.5. Phase contrast microscopy

Evaluation of potential cell morphological changes was assessed by phase contrast microscopy analysis at 24 h after incubation of the cultured cell lines with different concentrations of the compounds of interest (from 0.1 µg/ml to 3.2 µg/ml) using an inverted microscope (Nikon Eclipse TS100).

6.2.6. Fluorescence spectroscopy

Fluorescence measurements were performed in a Varian Cary Eclipse spectrofluorometer, equipped with a constant-temperature cell holder (PeltierMulticell Holder).

6.2.7. Transmission electron microscopy

TEM images were obtained using a HITACHI H-8100 instrument operating at an acceleration voltage of 200 kV. Samples for TEM analysis were prepared by depositing suspensions of the nanoparticles in high glucose, phenol red-free DMEM on carbon coated copper grids and allowing them to completely dry.

6.2.8. Statistical analysis

Data are presented as mean ± SEM (standard error of the mean). Statistical analysis was performed using the GraphPad Prism 6.02 software (San Diego, USA). Data were analysed by a non-parametric method the Kruskal–Wallis one-way analysis of variance by ranks followed by Dunn’s test. Significance was accepted at a p value ≤ 0.05.

6.3. Results and Discussion

In the studies of cell viability and cell morphology not only the effect of the fluorescent silica nanoparticles but also the effect of bare silica nanoparticles (with approximately 67 nm in diameter) and the fluorophores encapsulated in each type of particle (RBITC and LnPOMs) were evaluated. In the particular case of silica nanoparticles encapsulating LnPOMs, only the compound with higher stability was used. As described previously in chapter 5, the compound that leads to the more stable particles was the sandwich POM (Eu(PW11O39)2). The europium salt (europium 166 FCUP Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

chloride) used in the synthesis of the Eu(PW11O39)2 was also tested. In summary for the studies of cell viability and morphology the following particles and compounds were

used, RBITC, RBITC-APTES FSNPs (RBITC-APTES@SiO2), bare silica nanoparticles 3+ (SiO2 NPs), europium salt (Eu ), Eu(PW11O39)2 and Eu(PW11O39)2@SiO2.

The three different types of human cell lines, namely Caco-2, Hepa RG and SH- SY5Y cells, were incubated with the compounds of interest. The Caco-2 cell line is a continuous cell of heterogeneous human epithelial colorectal adenocarcinoma cells. Hepa RG cells are terminally differentiated hepatic cells derived from a human hepatic progenitor cell line that retains many characteristics of primary human hepatocytes, and SH-SY5Y is a neuroblastoma cell line.

After incubation of cells with the nanoparticles and compounds described above, cell integrity was evaluated by phase contrast microscopy. In the case of the fluorescent silica nanoparticles encapsulating the organic dye RBITC, a cellular uptake experiment was also performed.

6.3.1. Cellular uptake of silica nanoparticles

When investigating the cytotoxicity of the fluorescent silica nanoparticles, it is also important to understand the interaction of the nanoparticles during cell proliferation. In principle if nanoparticles could be taken into cells this would significantly affect their cytotoxicity.[5] Cellular uptake of silica NPs by the three different cell lines was evaluated by fluorescence emission of the RBITC-APTES FSNPs. After 24 and 48 h incubation, no fluorescence emission of RBITC-APTES FSNPs was observed even in cells exposed to the highest tested concentration (3.2 µg/mL). This finding could be due to the low concentration of NPs and consequently of RBITC, insufficient for detection of cellular uptake. According to the concentration of the silica NPs stock solution (1.6 mg/ml in ethanol), 3.2 µg/mL was the maximum concentration tested to avoid interference of ethanol in cellular viability (0.2% ethanol).

Since nanoparticles can be unstable in biological media, an additional experiment was perform to ensure that excessive aggregation was not occurring when the nanoparticles were dispersed in the medium required for the cell culture experiments. To ensure that NPs were present in solution, TEM imaging of a sample prepared from a 3.2 µg/ml NP solution in culture media (high glucose DMEM) was prepared. The excitation and emission fluorescence of the same sample was measured. As can be seen by TEM images (Figure 6.1) and fluorescence excitation and emission spectra FCUP 167 Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

(Figure 6.2), NPs are present and still show fluorescence but the signal is too small compared to the one of the stock solution. For this reason the fluorescence emission of the cells exposed to RBITC-APTES FSNPs was not detected through the microplate reader measurements.

Figure 6.1 - TEM images of RBITC-APTES FSNPs dried from high glucose DMEM at a concentration of 3.2 µg/ml.

A Excitation 1000 Emission RBITC-APTES@SiO RBITC-APTES@SiO 2 2 stock solution 800 stock solution

600

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200 RBITC-APTES@SiO 2 RBITC-APTES@SiO 2 in DMEM in DMEM 0 350 400 450 500 550 600 650 700 750 Wavelength (nm)

168 FCUP Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

B 20 RBITC-APTES@SiO 3.2 ug/mL in DMEM 2

Excitation Emission 15

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5

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Figure 6.2 - (A) Fluorescence excitation and emission spectra of stock solution of RBITC-APTES FSNPs (1.6mg/ml) in ethanol and RBITC-APTES FSNPs solution (3.2 µg/ml) in high glucose, phenol red-free DMEM ; (B) zoom of the fluorescence excitation and emission spectra of RBITC-APTES FSNPs solution (3.2 µg/ml) in high glucose, phenol red- free DMEM.

It would be necessary to make NPs which can be dispersed in aqueous solutions so that the uptake assays can be done with higher concentrations of nanoparticles. One way to overcome this issue is to synthesize NPs with water soluble groups such APTES on the surface. This approach has been already used in the group to prepare more water dispersible fluorescent silica NPs.

6.3.2. Cell esterase activity (Calcein-AM assay)

To investigate the potential cytotoxicity of the different fluorescent silica nanoparticles synthesized in this work, cell viability was measured after treatment with both kinds of silica NPs (RBITC-APTES FSNPs and LnPOMs silica NPs) at different

concentrations from 0.1 µg/ml to 3.2 µg/ml. Bare silica nanoparticles (SiO2 NPs), 3+ RBITC, europium salt (Eu ) and Eu(PW11O39)2 were also incubated with cells to test cell viability in presence of these compounds at the same concentrations. Sections 6.3.1.1 and 6.3.1.2 shows the results obtained in the three cell lines used (human intestinal epithelial Caco-2, human neuroblastoma SH-SY5Y and human hepatoma Hepa RG cells) for RBITC-APTES FSNPs and LnPOMs silica NPs respectively. FCUP 169 Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

6.3.2.1. Effect of RBITC@SiO2 NPs on Caco-2, SH-SY5Y and Hepa RG cells viability

Figures 6.3, 6.4 and 6.5 present the results of the effect of RBITC@SiO2 NPs,

RBITC and SiO2 NPs on cell esterase activity, as assessed by the calcein-AM assay on Caco-2, SH-SY5Y and HEPA RG cells respectively. The assays were conducted after 24 h and 48 h exposure of the cells to the different compounds.

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Figure 6.3 - Effect of RBITC-APTES@SiO2 NPs, RBITC and SiO2 NPs on esterase activity of human intestinal epithelial Caco-2 cells, as assessed by the calcein-AM assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage of control (vehicle-treated cells) and data are presented as mean ± SEM (n=4-28 per group).

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Figure 6.4 - Effect of RBITC-APTES@SiO2 NPs, RBITC and SiO2 NPs on esterase activity of human neuroblastoma SH-SY5Y cells, as assessed by the calcein-AM assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage of control (vehicle-treated cells) and data are presented as mean ± SEM (n=4-28 per group). 170 FCUP Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

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C C 5 0 S iO 2 N P s 2 5 S iO 2 N P s

0 .0 0 .6 1 .2 1 .8 2 .4 3 .0 3 .6 0 .0 0 .6 1 .2 1 .8 2 .4 3 .0 3 .6

C o n c e n tra tio n C o n c e n tra tio n ( g /m L ) ( g /m L )

Figure 6.5 - Effect of RBITC-APTES@SiO2 NPs, RBITC and SiO2 NPs on esterase activity of human hepatoma Hepa RG cells, as assessed by the calcein-AM assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage of control (vehicle-treated cells) and data are presented as mean ± SEM (n=8-28 per group).

From figures 6.3 to 6.5 it can be seen that for Caco-2 and Hepa RG cells

RBITC-APTES@SiO2 have a proliferative effect on cells meaning that the cells grow or multiply by rapidly producing cells and thus no cytotoxic behaviour is observed. The

same effect is observed for RBITC dye and silica NPs (SiO2) by their own. In the case of the SH-SY5Y cell line no differences were observed on the cell esterase activity up

to 48 h between RBITC, RBITC-APTES@SiO2 and SiO2 NPs, indicating that this cell line is less sensitive to the presence of the nanoparticles and RBITC dye. These results are to some extent expected since for similar systems (silica NPs with identical sizes on the same or similar cell lines) the cytotoxic effects were only observed at higher concentrations of NPs (from 25 µg/mL up).[7, 9, 20]

Among other factors such as size or shape, NPs effects upon cell viability has also been described as concentration-dependent. For instance, Foldbjerg et al.[20] have evaluated silica NPs toxicity in human (Caco-2 included) and murine cell lines using the Cell Counting Kit-8 (CCK-8) assay (which is based on the tetrazolium salt) after 24 h exposure to a range of NPs concentration from 0 to 300 µg/mL. In this study the authors investigated the toxicity of two kinds of silica NPs that they called unmodified and bovine serum albumin-stabilized (BSA) silica NPs. The results reported in their study have shown a concentration-dependent decrease in cell viability in all cell lines for both types of NPs. In the particular case of Caco-2 cells treated with unmodified silica NPs the decrease in cell viability was observed only from approximately 25 FCUP 171 Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

mg/mL. A small increase in cell viability was observed at concentrations lower than 25 µg/mL. This suggests that the results obtained in the present work are due to the lower concentrations used. In another study reported by Malvindi and coworkers[21], a non- toxic behaviour of silica NPs was observed in five different cell lines (including Caco-2 cells) when exposed to small concentrations over a period of 96 h. In this study, the authors investigated the possible cytotoxicity of three different sized silica NPs (25, 60 and 115 nm) on five cell lines using the water soluble tetrazolium salt-8 (WST-8) cell viability assay. For this purpose cells were incubated with different concentrations of NPs (from 2.5 pM up to 2500 pM) over a period of 96 h. Cell viability was evaluated in terms of concentration and time-dependence and the results reported have shown that upon exposure to increasing concentrations of any of the different sized silica NPs, viability of all cell lines was not altered up to 96 h. The authors suggested that the results obtained were likely due to the low NPs concentrations used.[21]

Regarding the results obtained for Hepa RG cell line in the present study, similar findings were previously reported in literature for another hepatic cell model, the human hepatoma HepG2 cells (a similar hepatic cell line to that used in the present work). Li et al.[7] have investigated the effect of three different sized silica nanoparticles (19, 43 and 68 nm) on HepG2 cell viability after incubation with increasing NPs concentrations (12.5 up to 200 µg/mL) for a period of 24 h. They reported that cell viability decreased as function of NPs concentration for all the three types of NPs and that for the 68 nm NPs (NPs with similar size to those synthesized in this work) the decreased in cell viability was observed at the concentration of 100 µg/mL.

Similar results to those reported here for SH-SY5Y cells, were described by other authors using small concentrations of silica NPs. Kim et al.[22] have investigated the cytotoxicity of LUDOX silica NPs (commercial colloidal silica NPs in aqueous phase) in the human neuronal SH-SY5Y cell line. In this study, SH-SY5Y cells were exposed to silica NPs at 10, 100 and 1000 ppm during periods of 6, 24 and 48 h. The authors reported that cell viability was concentration and time-dependent but for a dosage level of 10 ppm no significant cytotoxicity was observed up to 48 h. In a later study from the same authors[23] they reported that concentrations of polygon silica NPs <100 ppm had no significant impact on the viability of SH-SY5Y neuronal cells.

The results reported in the mentioned studies are relative to synthesized[7, 21] and commercial[20, 22, 23] plain silica NPs. As far as concerns there are no studies on the cytotoxic effects of fluorescent silica NPs in the cellular models used in the present 172 FCUP Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

study. In this way, the findings presented in here are a novelty and can be a starting point for future studies regarding these kinds of fluorescent nanomaterials.

However, as in the uptake experiments there would be a need to perform these studies with higher concentrations of NPs to obtain a broader picture of their cytotoxicity if they are to be used in greater concentrations. For that purpose it would be necessary to produce more water dispersible NPs either by functionalizing the particle’s surface with water soluble groups in a post synthesis step or by making them water soluble during the synthesis process.

6.3.2.2. Effect of Eu(PW11O39)2@SiO2 NPs on Caco-2, SH-SY5Y and Hepa RG cells viability

Figures 6.6, 6.7 and 6.8 show the results of the effect of Eu(PW11O39)2@SiO2 NPs, 3+ Eu(PW11O39)2, Eu salt and SiO2 NPs on cell esterase activity assessed by the calcein- AM assay on Caco-2, SH-SY5Y and HEPA RG cells respectively. The results were obtained after 24 h and 48 h exposure of the cells to the different compounds.

A B

2 0 0 2 0 0

y y

t t

i i

s s

n n e

e 1 7 5 1 7 5

t t

n n

i i

) )

l l

e e o

o 1 5 0 1 5 0

c c

r r

t t

n n

n n

e e

o o

c c

c c s

s 1 2 5 1 2 5

f f

e e

r r

o o

o o

u u

% % l

l 1 0 0 1 0 0

( (

f f

E u (P O M s )@ S iO 2 N P s E u (P O M s )@ S iO 2 N P s

n n

i i e

e E u 3 + E u 3 + c

c 7 5 7 5

l l a

a P O M s P O M s

C C 5 0 S iO 2 N P s 5 0 S iO 2 N P s

0 .0 0 .6 1 .2 1 .8 2 .4 3 .0 3 .6 0 .0 0 .6 1 .2 1 .8 2 .4 3 .0 3 .6

C o n c e n tra tio n C o n c e n tra tio n ( g /m L ) ( g /m L )

3+ Figure 6.6 - Effect of Eu(POMs)@SiO2 NPs, Eu , POMs and and SiO2 NPs on esterase activity of human intestinal epithelial Caco-2 cells, as assessed by the calcein-AM assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage of control (vehicle-treated cells) and data are presented as mean ± SEM (n=8-24 per group). FCUP 173 Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

A B

2 0 0 2 0 0

y y

t t

i i s

s E u (P O M s )@ S iO 2 N P s E u (P O M s )@ S iO 2 N P s n

n 3 + e

e 1 7 5 3 + 1 7 5 E u t

t E u

n n i

i P O M s

) ) l

l P O M s

e e o

o 1 5 0 1 5 0

c c r

r S iO 2 N P s t

t S iO N P s n

n 2

n n

e e

o o

c c

c c s

s 1 2 5 1 2 5

f f

e e

r r

o o

o o

u u

% % l

l 1 0 0 1 0 0

( (

f f

n n

i i

e e c

c 7 5 7 5

l l

a a

C C 5 0 5 0

0 .0 0 .6 1 .2 1 .8 2 .4 3 .0 3 .6 0 .0 0 .6 1 .2 1 .8 2 .4 3 .0 3 .6

C o n c e n tra tio n C o n c e n tra tio n ( g /m L ) ( g /m L )

3+ Figure 6.7 - Effect of Eu(POMs)@SiO2 NPs, Eu , POMs and SiO2 NPs on esterase activity of human neuroblastoma SH-SY5Y cells, as assessed by the calcein-AM assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage of control (vehicle-treated cells) and data are presented as mean ± SEM (n=8-24 per group).

A B

2 0 0 2 0 0

y y t

t E u (P O M s )@ S iO N P s i

i E u (P O M s )@ S iO 2 N P s 2 s

s 3 + 3 + n

n E u E u e

e 1 7 5 1 7 5 t

t P O M s n

n P O M s

i i

) )

l l e

e S iO N P s S iO N P s o

o 1 5 0 2 1 5 0 2

c c

r r

t t

n n

n n

e e

o o

c c

c c s

s 1 2 5 1 2 5

f f

e e

r r

o o

o o

u u

% % l

l 1 0 0 1 0 0

( (

f f

n n

i i

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c 7 5 7 5

l l

a a

C C 5 0 5 0

0 .0 0 .6 1 .2 1 .8 2 .4 3 .0 3 .6 0 .0 0 .6 1 .2 1 .8 2 .4 3 .0 3 .6

C o n c e n tra tio n C o n c e n tra tio n ( g /m L ) ( g /m L )

3+ Figure 6.8 - Effect of Eu(POMs)@SiO2 NPs, Eu , POMs and and SiO2 NPs on esterase activity of human hepatoma Hepa RG cells, as assessed by the calcein-AM assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage of control (vehicle-treated cells) and data are presented as mean ± SEM (n=8-28 per group).

Similar to the results obtained for RBITC@SiO2, RBITC and SiO2 NPs (presented in 3+ the previous section 6.3.2.1) are the results of Eu(PW11O39)2, Eu salt

Eu(PW11O39)2@SiO2 NPs, and SiO2 NPs on cell esterase activity in the three cell lines used. 174 FCUP Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

From figures 6.6 to 6.8 it can be seen that for Caco-2 and Hepa RG cells

Eu(PW11O39)2@SiO2 NPs have a proliferative effect on cells as well as Eu(PW11O39)2, 3+ Eu and SiO2 NPs. In case of Caco-2 cellular line the proliferative effect is more evident at 48h. For SH-SY5Y cell line no differences are observed on the cell esterase activity up to 48 h between the different compounds tested.

In principle is expected that POMs by themselves won’t cause cytotoxic effects since they are considered to exhibit low toxicity.[24] For instance, cytotoxic effects were investigated by Geisberger et al.[25] for a wells-Dawson type POM and no significant impact on the cell viability of HeLa cancer cells was observed for POM concentrations up to 100 µg/mL. The same authors also reported the potential cytotoxic effect on HeLa 10- cells of a POM ([Co4(H2O)2(PW9O34)2] ) encapsulated into a carboxymethyl chitosan (CMC) matrix.[26] The authors found that at the higher concentration used (2 mg/mL) the POM/CMC nanocomposites did not display cytotoxicity. However, regarding silica NPs encapsulating POMs, to the extent that it is known no results related with the possible cytotoxicity of these nanomaterials are available.

Once again, the results presented in here are a novelty and give important information for further studies regarding biological applications of LnPOMs based fluorescent silica nanoparticles.

6.3.3. Morphological analysis by phase contrast microscopy

Potential morphological changes of Caco-2, SH-SY5Y and Hepa RG cells in response to the silica NPs were investigated by phase contrast microscopy. Sections 6.3.2.1 and 6.3.2.2 present the observed results for the three cell lines incubated with RBITC FSNPs and LnPOMs silica NPs respectively.

6.3.3.1. RBITC-APTES FSNPS

Figures 6.9, 6.10 and 6.11 show the images of phase contrast microscopy of

RBITC, RBITC-APTES@SiO2 and SiO2 NPs incubated with Caco-2, SH-SY5Y and HEPA RG cells respectively. The images were collected after 24 h incubation at a 100x magnification.

FCUP 175 Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

Figure 6.9 - Representative phase contrast microscopy images of Caco-2 cells at 24 hours after incubation with SiO2 nanoparticles, RBITC dye and RBITC-APTES@SiO2 nanoparticles (100x magnification).

176 FCUP Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

Figure 6.10 - Representative phase contrast microscopy images of SH-SY5Y cells at 24 hours after incubation with SiO2

nanoparticles, RBITC dye and RBITC-APTES@SiO2 nanoparticles (100x magnification).

FCUP 177 Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

Figure 6.11 - Representative phase contrast microscopy images of Hepa RG cells at 24 hours after incubation with SiO2 nanoparticles, RBITC dye and RBITC-APTES@SiO2 nanoparticles (100x magnification).

Phase contrast microscopy images presented in figures 6.9 to 6.11 shows normal cell morphology after incubation with RBITC-APTES@SiO2, RBITC and SiO2 NPs. This indicates that no morphological changes had occurred in the three cell lines after incubation with the RBITC dye and the NPs. Results on cell morphology are in agreement with those obtained on cell viability measurements since no cytotoxic effects were observed by either technique.

6.3.3.2. Eu(PW11O39)2@ SiO2 NPs

Figures 6.12, 6.13 and 6.14 show the images of phase contrast microscopy of Eu+3,

Eu(PW11O39)2, Eu(PW11O39)2@SiO2 and SiO2 NPs incubated with Caco-2, SH-SY5Y and HEPA RG cells respectively. The images were collected after 24 h incubation at a 100x magnification. 178 FCUP Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

Figure 6.12 - Representative phase contrast microscopy images of Caco-2 cells at 24 hours after incubation with SiO2

nanoparticles, Eu salt, Eu(PW11O39)2 and Eu(PW11O39)2@SiO2 nanoparticles (100x magnification).

Figure 6.13 - Representative phase contrast microscopy images of SH-SY5Ycells at 24 hours after incubation with SiO2

nanoparticles, Eu salt, Eu(PW11O39)2 and Eu(PW11O39)2@SiO2 nanoparticles (100x magnification). FCUP 179 Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

Figure 6.14 - Representative phase contrast microscopy images of Hepa RGcells at 24 hours after incubation with SiO2 nanoparticles, Eu salt, Eu(PW11O39)2 and Eu(PW11O39)2@SiO2 nanoparticles (100x magnification).

Regarding the potential changes in cell morphology by phase contrast microscopy it can be observed from figures 6.12 to 6.14 that for the three cellular lines used there were no morphological changes after cell treatment with Eu3+,

Eu(PW11O39)2, Eu(PW11O39)2@SiO2 or SiO2 NPs. Similar to the results presented in the previous section (6.3.3.1), phase contrast microscopy images show normal cell morphology after cell incubation with the compounds of interest after 24 h. Again results on cell morphology are in agreement with those obtained on cell viability measurements.

6.4. Conclusions

Cellular uptake, cellular viability and cellular morphology experiments were performed to evaluate possible cytotoxicity of the fluorescent silica NPs synthesized in the present work. For this purpose Caco-2, SH-SY5Y and Hepa RG cellular lines were incubated with increasing concentrations from 0.1 µg/mL up to 3.2 µg/mL of the 180 FCUP Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

fluorescent silica NPs and their corresponding fluorophores. Solutions of bare silica NPs with similar size and at the same concentrations were also tested.

Cellular uptake experiments were performed for the fluorescent silica NPs encapsulating RBITC dye. Cellular uptake of these NPs was evaluated by

measurement of the fluorescence emission of the RBITC-APTES@SiO2 after incubation with the different cell lines for 24 and 48 h using a microplate reader. After 24 and 48 h incubation, no fluorescence was detected. However, the concentrations tested were very low when compared to the stock solution (1.6 mg/mL) as shown by

the fluorescence excitation and emission spectra of the RBITC-APTES@SiO2 solutions in DMEM (3.2 µg/mL) and in ethanol (1.6 mg/ml). Fluorescence and emission spectra

of RBITC-APTES@SiO2 solution in DMEM present a much lower signal compared to

that of RBITC-APTES@SiO2 stock solution in ethanol. These results show that NPs still present fluorescence but the signal is too small to be detected by the microplate reader suggesting that higher concentrations need to be test in order to check cellular uptake of NPs.

The cytotoxicity of bare silica NPs, fluorescent silica NPs and their corresponding fluorophores was examined by the calcein-AM assay. In case of the Caco-2 and Hepa RG cellular lines, the calcein-AM assay showed that cells can survive and proliferate in the presence of all the components tested. For the SH-SY5Y cellular line, no significant changes were observed in cellular viability after incubation with all the compounds indicating a less sensitivity of this cellular line in the presence of the NPs or the fluorophores. At the concentrations tested, all the compounds presented a non-toxic behaviour. Nevertheless, since in the literature there is evidence of cytotoxicity of plain silica NPs at higher concentrations (> 25 µg/mL), testing the NPs developed in this study in higher concentrations could be a subject for further investigation.

Cellular integrity was evaluated by phase contrast microscopy after incubation of cells with NPs and the fluorophores. Microscopic analysis showed normal cell morphology after treatment with the fluorescent silica NPs and their corresponding fluorophores meaning that treated cells have similar morphology to the control ones. Results on cell morphology are in agreement with those of cell viability.

FCUP 181 Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

6.5. References

1. Ruedas-Rama, M.J., Walters, J.D., Orte, A. and Hall, E.A.H., Fluorescent nanoparticles for intracellular sensing: A review. Analytica Chimica Acta, 2012. 751: p. 1-23. 2. Erathodiyil, N. and Ying, J.Y., Functionalization of inorganic nanoparticles for bioimaging applications. Accounts of Chemical Research, 2011. 44(10): p. 925- 35. 3. Williams, D.F., On the mechanisms of biocompatibility. Biomaterials, 2008. 29(20): p. 2941-2953. 4. Li, X.M., Wang, L., Fan, Y.B., Feng, Q.L. and Cui, F.Z., Biocompatibility and Toxicity of Nanoparticles and Nanotubes. Journal of Nanomaterials, 2012. 5. Jin, Y.H., Kannan, S., Wu, M. and Zhao, J.X.J., Toxicity of luminescent silica nanoparticles to living cells. Chemical Research in Toxicology, 2007. 20(8): p. 1126-1133. 6. Duan, J.C., Yu, Y.B., Li, Y., Yu, Y., Li, Y.B., Zhou, X.Q., Huang, P.L. and Sun, Z.W., Toxic Effect of Silica Nanoparticles on Endothelial Cells through DNA Damage Response via Chk1-Dependent G2/M Checkpoint. Plos One, 2013. 8(4). 7. Li, Y., Sun, L., Jin, M.H., Du, Z.J., Liu, X.M., Guo, C.X., Li, Y.B., Huang, P.L. and Sun, Z.W., Size-dependent cytotoxicity of amorphous silica nanoparticles in human hepatoma HepG2 cells. Toxicology in Vitro, 2011. 25(7): p. 1343-1352. 8. Miletto, I., Gilardino, A., Zamburlin, P., Dalmazzo, S., Lovisolo, D., Caputo, G., Viscardi, G. and Martra, G., Highly bright and photostable cyanine dye-doped silica nanoparticles for optical imaging: Photophysical characterization and cell tests. Dyes and Pigments, 2010. 84(1): p. 121-127. 9. Soenen, S.J., Manshian, B., Doak, S.H., De Smedt, S.C. and Braeckmans, K., Fluorescent non-porous silica nanoparticles for long-term cell monitoring: cytotoxicity and particle functionality. Acta Biomaterialia, 2013. 9(11): p. 9183- 93. 10. Akbar, N., Mohamed, T., Whitehead, D. and Azzawi, M., Biocompatibility of amorphous silica nanoparticles: Size and charge effect on vascular function, in vitro. Biotechnology and Applyed Biochemistry, 2011. 58(5): p. 353-62. 11. Barandeh, F., Nguyen, P.L., Kumar, R., Iacobucci, G.J., Kuznicki, M.L., Kosterman, A., Bergey, E.J., Prasad, P.N. and Gunawardena, S., Organically 182 FCUP Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

Modified Silica Nanoparticles Are Biocompatible and Can Be Targeted to Neurons In Vivo. Plos One, 2012. 7(1). 12. Lin, W.S., Huang, Y.W., Zhou, X.D. and Ma, Y.F., In vitro toxicity of silica nanoparticles in human lung cancer cells. Toxicology and Applied Pharmacology, 2006. 217(3): p. 252-259. 13. Ahmad, J., Ahamed, M., Akhtar, M.J., Alrokayan, S.A., Siddiqui, M.A., Musarrat, J. and Al-Khedhairy, A.A., Apoptosis induction by silica nanoparticles mediated through reactive oxygen species in human liver cell line HepG2. Toxicology and Applied Pharmacology, 2012. 259(2): p. 160-168. 14. Liu, T., Li, L., Fu, C., Liu, H., Chen, D. and Tang, F., Pathological mechanisms of liver injury caused by continuous intraperitoneal injection of silica nanoparticles. Biomaterials, 2012. 33(7): p. 2399-407. 15. Nishimori, H., Kondoh, M., Isoda, K., Tsunoda, S., Tsutsumi, Y. and Yagi, K., Silica nanoparticles as hepatotoxicants. European Journal of Pharmaceutics and Biopharmaceutics, 2009. 72(3): p. 496-501. 16. Park, E.J. and Park, K., Oxidative stress and pro-inflammatory responses induced by silica nanoparticles in vivo and in vitro. Toxicology Letters, 2009. 184(1): p. 18-25. 17. Riss, T.L. and Moravec, R.A., Use of multiple assay endpoints to investigate the effects of incubation time, dose of toxin, and plating density in cell-based cytotoxicity assays. Assay and Drug Development Technologies, 2004. 2(1): p. 51-62. 18. Niles, A.L., Moravec, R.A., Hesselberth, P.E., Scurria, M.A., Daily, W.J. and Riss, T.L., A homogeneous assay to measure live and dead cells in the same sample by detecting different protease markers. Analytical Biochemistry, 2007. 366(2): p. 197-206. 19. Mosmann, T., Rapid Colorimetric Assay for Cellular Growth and Survival - Application to Proliferation and Cyto-Toxicity Assays. Journal of Immunological Methods, 1983. 65(1-2): p. 55-63. 20. Foldbjerg, R., Wang, J., Beer, C., Thorsen, K., Sutherland, D.S. and Autrup, H., Biological effects induced by BSA-stabilized silica nanoparticles in mammalian cell lines. Chemico-Biological Interactions, 2013. 204(1): p. 28-38. 21. Malvindi, M.A., Brunetti, V., Vecchio, G., Galeone, A., Cingolani, R. and Pompa, P.P., SiO2 nanoparticles biocompatibility and their potential for gene delivery and silencing. Nanoscale, 2012. 4(2): p. 486-495. FCUP 183 Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles

22. Kim, Y.J., Yu, M., Park, H.O. and Yang, S.I., Comparative study of cytotoxicity, oxidative stress and genotoxicity induced by silica nanomaterials in human neuronal cell line. Molecular & Cellular Toxicology, 2010. 6(4): p. 337-344. 23. Kim, Y.J. and Yang, S.I., Neurotoxic effects by silica TM nanoparticle is independent of differentiation of SH-SY5Y cells. Molecular & Cellular Toxicology, 2011. 7(4): p. 381-388. 24. Hungerford, G., Hussain, F., Patzke, G.R. and Green, M., The photophysics of europium and terbium polyoxometalates and their interaction with serum albumin: a time-resolved luminescence study. Physical Chemystry Chemical Physics, 2010. 12(26): p. 7266-75. 25. Geisberger, G., Gyenge, E.B., Hinger, D., Bosiger, P., Maake, C. and Patzke, G.R., Synthesis, characterization and bioimaging of fluorescent labeled polyoxometalates. Dalton Transactions, 2013. 42(27): p. 9914-9920. 26. Geisberger, G., Paulus, S., Carraro, M., Bonchio, M. and Patzke, G.R., Synthesis, Characterisation and Cytotoxicity of Polyoxometalate/Carboxymethyl Chitosan Nanocomposites. Chemistry - A European Journal, 2011. 17(16): p. 4619-4625.

184 FCUP

III Concluding Remarks and Perspectives

FCUP 187

Concluding Remarks and Perspectives

Fluorescent silica nanoparticles show unique chemical and optical properties such as bright fluorescence, higher photostability and biocompatibility compared to classical fluorophores. Moreover, fluorescent silica nanoparticles are quite easy to prepare, exhibit good monodispersity and their surface can be easily functionalized for further bioconjugation. Due to these attractive features, fluorescent silica nanoparticles have received an increasing interest for biological applications in the past few years. However, despite the increasing applicability in biological fields the biocompatibility of these nanoparticles is not completely clear and remains an issue of debate. Therefore, nanotoxicology research has been intensified in order to obtained detailed information about biocompatibility of silica based nanomaterials

The purpose of this research work was to prepare and optimize the synthesis of two different kinds of fluorescent silica nanoparticles incorporating organic (rhodamine b isothiocyanate – RBITC) and inorganic (lanthanide-based polyoxometalates – LnPOMs) fluorophores. It was also an aim of this study the characterization of these nanomaterials in order to evaluate the effects of silica encapsulation on the stability, fluorescence emission and quantum yield of the fluorophores. In addition, the evaluation of the potential cytotoxicity of these nanoparticles was also a study purpose.

In this work it was clearly demonstrated that using a reverse microemulsion methodology for the alkaline hydrolysis of TEOS it was possible to produce fluorescent silica nanoparticles incorporating the different fluorophores. All the samples of the fluorescent nanoparticles synthesized show a narrow polydispersity. Furthermore, in the synthesis of fluorescent silica NPs encapsulating the inorganic fluorophores (LnPOMs) the ratio of LnPOM/TEOS used has a strong influence on the nanoparticle final size indicating that LnPOMs act as seeds for the growth of the silica shell. In this work, it was found that the reverse-microemulsion-based methodology gives superior results compared to the Stöber method regarding size and shape control and reproducibility and also in terms of fluorescent yield and stability.

After encapsulation of the different fluorophores within the silica NPs the structural integrity, stability and quantum yield were evaluated. RBITC silica NPs show similar spectral properties (absorption, fluorescence excitation and emission) to the free dye, indicating the successful doping of dye molecules into silica matrix. One of the 188 FCUP Concluding Remarks and Perspectives

advantages of encapsulating fluorophores in silica frameworks is brightness enhancement of fluorophore molecules due to the fact that one nanoparticle can incorporate more than one molecule. In the work described here the fluorescent

RBITC-APTES@SiO2 nanoparticles encapsulated between 100 and 150 dye molecules per particle, on average. The silica matrix also acts as a shield, protecting the encapsulated fluorophores from the outer environment that can be responsible for their photobleaching. This capability was observed by the increase of fluorescence quantum yield, fluorescence lifetime and steady state fluorescence anisotropy of

RBITC-APTES@SiO2 NPs compared to the free dye. Fluorescence anisotropy also suggested that the RBITC dye was covalently bound to the silica matrix due to the restricted motion of RBITC molecules within the silica matrix. This motion of the encapsulated dye molecules was described as being a wobbling-in-cone model.

Regarding the encapsulation of LnPOMs in silica NPs two europium

polyoxometalates, with distinct europium coordination spheres (Eu(PW11)x, x = 1 and 2), were successfully encapsulated into a silica matrix for the first time. Silica

incorporation of Eu(PW11)x was confirmed by FT-IR and FT-Raman spectroscopy. The

distinct coordination of the europium centre in Eu(PW11)x compounds seems to influence the stability of the polyoxometalate structure during silica encapsulation as 31 showed by P solid NMR. Eu(PW11)2 containing europium coordinated with two units

of monovacant precursor yields Eu(PW11)2@SiO2 NPs where the polyoxometalate

maintains its structural integrity, while the EuPW11 structure, seemed to be less stable during the process of silica encapsulation. Furthermore, 31P NMR analysis suggests

some structural decomposition of EuPW11 into PW11 during EuPW11@SiO2 NPs

preparation. Photoluminescence studies were also used to demonstrate Eu(PW11)x encapsulation and the effect of the silica shell on the protection from the outer environment. These studies showed that at ambient conditions (298 K; pressure of 1 bar) the photoluminescence properties of the silica encapsulated europium compounds are very similar to the parental POMs. However, in high vacuum, the

photoluminescence properties of Eu(PW11)x compounds change whereas those of

Eu(PW11)x@SiO2 remained unaltered. The quantum efficiency of Eu(PW11)2 was determined before and after encapsulation and the results show an increase of

quantum efficiency of the Eu(PW11)2 upon incorporation within the silica matrix.

To be able to employ the fluorescent silica NPs in biological systems for the attachment of a specific target, surface modification of these NPs is required. The fluorescent silica NPs were functionalized with organosilanes (GPTMs and CPTEs) and FCUP 189 Concluding Remarks and Perspectives

functionalization was confirmed by C and H elemental analysis. The readily available thiol groups on biomolecules, such as SH-terminated DNA are good candidates to couple to the fluorescent silica NPs. Based on this an attempt to conjugate the NPs with oligonucleotides was made for the RBITC-APTES@SiO2 NPs system functionalized with GPTMs. UV-vis spectroscopy was used to check if immobilization occurred but the technique seems not to be sensitive enough to prove that the binding occurred. Particle’s surface zeta potential ζ was also measured and a change on zeta potential ζ was observed after reaction of FSNPs-GPTES with ssDNA. The difference on surface charge could indicate DNA immobilization onto silica NPs surface but further experiments, such as agarose gel electrophoresis, will be needed to actually prove that immobilization took place.

Cytotoxicity of the fluorescent silica NPs was evaluated by cell uptake, cell viability and cell morphology of three different cell lines (Caco-2, SH-SY5Y and Hepa RG) after incubation with the fluorescent silica NPs. The strong optical properties of the nanoparticles can help in their easy tracking within the cells. However, the emission of many fluorophores and cell stains lies in the range from 500 to 600 nm and due to a strong overlapping it becomes difficult to distinguish between the fluorophores and the cells. The low concentration of NPs in the present work (3.2 µg/mL) hindered NPs cellular uptake measurements. Cell viability by the calcein-AM assay after exposure of the three cell lines to increasing concentrations of both RBITC and LnPOMs silica NPs, ranging from 0.1 µg/mL up to 3.2 µg/mL, showed that after 48 h the NPs don’t inhibit cell viability. Microscopic analysis also showed normal cell morphology after treatment with the fluorescent silica NPs and their corresponding fluorophores, meaning that treated cells have similar morphology to the control ones. Results on cell morphology are in agreement with those of cell viability showing that in the concentration range tested all the NPs presented a non-toxic behaviour.

The requirements for development of fluorescent silica nanoparticles incorporating organic and inorganic fluorophores were met and led to the synthesis of bright fluorescent silica NPs. Silica encapsulation of both kinds of fluorophores proved to be an excellent route not only for protecting the fluorophores from the outer environment but also to enhance their photophysical properties such as fluorescence emission and quantum yield. Silica surface has a high affinity for conjugation with derivatized silanes and a procedure has been applied to conjugate these nanomaterials with DNA. However, the technique used in here was not sensitive enough to prove bioconjugation 190 FCUP Concluding Remarks and Perspectives

of DNA to NPs and therefore it is necessary to further characterize the product obtained. Cytotoxic measurements reveal that in the concentration range tested the synthesized fluorescent silica NPs present a non-toxic behaviour in all the cell lines studied. Concerning the future bioapplications of these NPs, the toxicological tests gave important information, especially in case of NPs incorporating LnPOMs since as far as we know no cytotoxic studies were yet reported. However, these tests were applied with low concentrations (< 3.5 µg/mL) of NPs and since in literature there is evidence of cytotoxic effects of bare silica NPs at higher concentrations (> 25 µg/mL) testing the NPs developed in this study in higher concentrations could be a subject of further investigation. Apart from that, further studies to confirm that there is no significant leaking of the fluorophores, either in solution or in cells, as well as studying the cytotoxic effect with longer incubation times should also be aim of future work.

Fluorescent silica NPs have been successfully used in biological fields but are far from reaching their full potential. The applications of these NPs are evolving rapidly as researchers improve their ability to manipulate and apply them. Improvements in the NP dispersion should prevent agglomeration, decrease background noise, and reduce nonspecific adhesion to surfaces. As soon as the photostable and highly fluorescent silica NPs are better implemented into the complex biological field, they will have a great impact and applicability in areas such as bioanalysis, molecular imaging, and biotechnology. Although there are still many challenges related with the biological applications of fluorescent silica NPs, the results described in this thesis give an idea of the potential of these nanomaterials.