development of fluorescent silica ddevelopment of fluorescent silica nanoparticles encapsulating...

Development of

fluorescent silica

nanoparticles

encapsulating

organic and

inorganic

fluorophores:

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

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FCUP

FCT/UNL

2014

3.º

CICLO

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

Cristina Sofia dos Santos Neves Doutoramento em Química Sustentável Departamento de Química e Bioquímica 2014

Orientador Peter Eaton, Investigador Auxiliar, Faculdade de Ciências

Coorientador Eulália Pereira, Professora Auxiliar, Faculdade de Ciências

“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.

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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.

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Acknowledgments

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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

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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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5

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.

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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.

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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

[PW11O39Eu(H2O)3]4- and the sandwich-type [Eu(PW11O39)2]

11- 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

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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

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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.

Foram utilizados LnPOMs do tipo Keggin, um monosubstituído [PW11O39Eu(H2O)3]4- e

outro do tipo sanduíche [Eu(PW11O39)2]11-. 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

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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

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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

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

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General Index

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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

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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


5.2.1. 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

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15

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


6.3.1. 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

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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


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

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

rates and 5D0 quantum efficiency, q, for compounds Eu(PW11)2 and Eu(PW11)2@SiO2.

The data have been obtained at room temperature (296 K). ..................................... 154


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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

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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

fundamental unit MO6. (Source: Fernandez[16]) ........................................................... 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

([M6O19)n-) isopolyanion; B) Anderson-Evans ([XM6O24]

n-); C) Keggin ([XM12O40]n-); D)

Wells-Dawson ([X2M18O62]n-) and E) Preyssler ([XP5W30O110]

n-) 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

anion. The rotated M3O13 groups are highlighted (dark blue). (source: Lopez et al.[22]) 68

Figure 3.6 - Ball and stick (left) and polyhedral representation (right) for the α-

[XM12O40]n- Keggin anion showing the different classification of the oxygen atoms...... 68

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

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

1:2 [M’(XM11O39)2]n- (right). .......................................................................................... 69

Figure 3.9 - Formation scheme of the monolacunar (A) and sandwich type (B)

lanthanide-substituted Keggin anion [Ln(XM11O39)x]n-. ................................................. 71

Figure 3.10 – Photoluminescence emission spectra of the Eu3+ ion in water. The

radiative transitions take place from the 5D0 level. (Adapted from Werts[50]) ................ 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

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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

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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,

[Eu(PW11O39)2]11−; (b) its {EuO8} coordination center displaying a square-antiprismatic

geometry and (c) the mono-substituted europium-phosphotungstate anion,

[PW11Eu(H2O)3O39]4- 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

Figure 5.3 - 31P 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

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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

Figure 5.12 - Solid state 31P 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

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(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

Figure 5.15 - Eu3+ 5D0 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


activity of human neuroblastoma SH-SY5Y cells, as assessed by the calcein-AM



group). ...................................................................................................................... 169


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

Figure 6.6 - Effect of Eu(POMs)@SiO2 NPs, Eu3+, 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

Figure 6.7 - Effect of Eu(POMs)@SiO2 NPs, Eu3+, POMs and SiO2 NPs on esterase

activity of human neuroblastoma SH-SY5Y cells, as assessed by the calcein-AM

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group). ...................................................................................................................... 173

Figure 6.8 - Effect of Eu(POMs)@SiO2 NPs, Eu3+, POMs and and SiO2 NPs on

esterase activity of human hepatoma Hepa RG cells, as assessed by the calcein-AM



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



Figure 6.14 - Representative phase contrast microscopy images of Hepa RGcells at 24




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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

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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

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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

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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

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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]

)

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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).

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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).

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N

X

N

Y

R R

n

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.

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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]

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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

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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.

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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.

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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]

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]

)

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,

DNA, etc. (Figure 1.9). Moreover pH

changes do not lead to swelling and

porosity changes, and silica particles

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.

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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.

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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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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

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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

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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

controlled by the water-to-surfactant molar ratio (W0).[2]

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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

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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

possible. Recently Wang et al.[17] 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]

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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]

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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]

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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.

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

)

Organosilanes used on surface

modification Structure

Functional

group on NPs

Target

biomolecules Bioconjugation method

3-mercaptopropyltrimethoxysilane

(MPTS) Si

OCH3

OCH3

OCH3

HS

-SH

Dissulfide modified

oligonucleotides

(-S-S-)

Thiol-dissulfide exchange

(3-aminopropyl)-triethoxysilane

(APTES)

Si NH2

O CH3

OH3C

O

CH3

-NH2

Antibodies

(-NCS)

Amine-thiocyanate coupling

Carboxyethylsilanetriol

(CTES) +NaO Si

OH

OH

OH

O

-COOH

Proteins or other

amine containing

biomolecules

(-NH2)

Carbodiimide chemistry

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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

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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].

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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)-

tris(4,7-diphenyl-1,10-phenantroline) ([Ru(dpp)3]2+) and the analyte-insensitive

reference dye Oregon Green 488 dextran into silica NPs called PEBBLEs (probes

encapsulated by biologically localized embedding). The ([Ru(dpp)3]2+ 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

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57

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


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.

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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.

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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.

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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.

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751: p. 1-23.

25. Santra, S., Zhang, P., Wang, K.M., Tapec, R. and Tan, W.H., Conjugation of



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.

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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.

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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]

)

POMs can be classified in two main classes, the isopolyanions ([MmOy]p-) and the

heteropolyanions ([XxMmOy]q-). 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

http://en.wikipedia.org/wiki/Molybdenum

http://en.wikipedia.org/wiki/Tungsten

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65

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.

Figure 3.3 - Polyhedral representation of common polyoxoanions: A) Lindqvist ([M6O19)n-) isopolyanion; B) Anderson-

Evans ([XM6O24]n-); C) Keggin ([XM12O40]

n-); D) Wells-Dawson ([X2M18O62]

n-) and E) Preyssler ([XP5W30O110]

n-)

heteropolyanions. (Source: Lopez et al. [22])

3.1.2. Historical context

In 1826 Berzelius[23] reported the discovery of the first POM, the phosphomolybdate,

of formula [PMo12O40]3-. 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

understand the structure of POMs, proposing that the anion [PMo12O40]3- 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

[XM12O40]n- 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

[XM6O24]n- and the isopolyanions [Mo7O24]

6- 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

[TeMo6O24]6- 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

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heptamolybdate anion ([Mo7O24]6-) proving that the geometry of the heteropolyanions

was non-planar. Three years later, Dawson[30] reported the structure of another anion

the heteropolyanion of formula [P2W18O62]6- 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

dissolving [MOn]m− 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

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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

characterization in 1934[26] has a general formula [XM12O40]n- (M = Mo6+, W6+; X = P5+,

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

corner-shared oxygen atoms and through the central XO4 tetrahedron.[5]

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]

)

The Keggin anion has several geometrical isomers (rotational isomers or isomers

Baker-Figgis)[36], 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]

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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

designated as Ob and Oc respectively and Od represents a terminal oxygen.[37-39] 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-

[XM12O40]n-.

Figure 3.6 - Ball and stick (left) and polyhedral representation (right) for the α-[XM12O40]n- Keggin anion showing the

different classification of the oxygen atoms.

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69

From the Keggin anion it is possible to obtain several lacunar structures by

removing one or more MOx octahedrons.[5, 21] The monolacunar anion [XM11O39](n+4)-

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).

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

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).

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

n- (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).

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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]

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

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]

Lacunar anion [PW11O39]7- is formed when one of the W(VI) metals and its

terminally bound oxo group are missing. Lanthanide substituted [PW11Ln(H2O)3O39]4- 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).

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Figure 3.9 - Formation scheme of the monolacunar (A) and sandwich type (B) lanthanide-substituted Keggin anion

[Ln(XM11O39)x]n-.

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

[Ln(XW11O39)2]n- type with X = P and Ln = Ce3+, Ce4+, Pr3+ and Nd3+ (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

[Ln(XM11O39)2]m- 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

describe the synthesis and crystal structures of the 1:2 [Ln(PMo11O39)2]11– 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

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crystal structures of the silicotungstates [Ln(SiW11O39)(H2O)n]5- with Ln3+ = La3+, Ce3+,

Eu3+, Yb3+. Zhang[46] and co-workers have also reported the crystalline structure of the

phosphotungstate [Ce(PW11O39)(H2O)2]4-. 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:

[{Ln(H2O)3(α-PW11O39)}2]6- (Ln3+ = Nd3+ and Gd3+) and [{Ln(H2O)(α-

PW11O39)(CH3COO}2]10- (Ln3+ = Sm3+, Eu3+, Gd3+, Tb3+, Ho3+ and Er3+).

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

http://en.wikipedia.org/wiki/Metal

http://en.wikipedia.org/wiki/Chemical_element

http://en.wikipedia.org/wiki/Atomic_number

http://en.wikipedia.org/wiki/Scandium

http://en.wikipedia.org/wiki/Yttrium

http://en.wikipedia.org/wiki/Rare_earth_elements

http://en.wikipedia.org/wiki/Lanthanum

http://en.wikipedia.org/wiki/Electronic_configuration

http://en.wikipedia.org/wiki/Main_group_element

http://en.wikipedia.org/wiki/Main_group_element

http://en.wikipedia.org/wiki/Transition_metal

https://www.boundless.com/chemistry/definition/oxidation-state--2/

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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

5D0 level. (Adapted from Werts

[50])

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

http://en.wikipedia.org/wiki/Nuclear_charge

http://en.wikipedia.org/wiki/Chemical_series

http://en.wikipedia.org/wiki/Ionic_radii

http://en.wikipedia.org/wiki/Lanthanide_contraction

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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)

Eu3+

5D0 →

7F0 580

Nd3+

4F3/2 →

4F9/2 880

5D0 →

7F1 590

4F3/2 →

4F11/2 1060

5D0 →

7F2 613

4F3/2 →

4F13/2 1330

5D0 →

7F3 650

5D0 →

7F4 690

5D0 →

7F5

710 Er

3+

4I13/2 →

4I15/2 1150

Tb3+

5D4 →

7F6

490

5D4 →

7F5

545

5D4 →

7F4

590 Yb

3+

2F5/2 →

2F7/2 980

5D4 →

7F3

620

5D4 →

7F2

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]

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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] 4f65d

0 – 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

([Eu(SiMoW10O39)2]13-) into silica nanoparticles was performed by Green et al. in

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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

([Ln(CoW11O39)(H2O)3]7- with Ln3+ = Ce3+, La3+, Eu3+, Sm3+ and Tb3+) 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

the preparation of core/shell nanoparticles using [Ln(W5O18)2]9- (Ln3+ = Eu3+, Tb3+, Gd3+)

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

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77

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

the use of lanthanophosphopolyoxotungstates ([Ln(PW11O39)2)]11- with Ln3+ = La3+, Pr3+,

Sm3+ and Tb3+) 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

(Ce) containing POM. The [CeSiW11O39]4- 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

catalytic property of [Nd(SiMo7W4)]13- 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

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photocatalysts [Ln(PW11O39)2]/PVA with Ln3+ = La3+, Ce3+, Pr3+, Nd3+ and Sm3+) 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

use of two gadolinium (Gd) containing POMs ([GdW10O36]9- and [Gd(PW11O39)2]

11-) 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

commercial and widely used MRI contrast agent Gd-DTPA. Furthermore [GdW10O36]9-

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

[Gd(SiW11O39)2]13- and [Gd3O3(SiW9O34)2]

11-. 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

single molecular magnets (SMM), particularly for complexes of the type [Ln(W5O18)2]9-

with Ln3+ = Ho3+, Er3+ and [Ln(SiW11O39)2]13- with Ln3+ = Dy3+, Ho3+, Er3+, Yb3+. 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.

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79

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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

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49. Binnemans, K., Lanthanide-Based Luminescent Hybrid Materials. Chemical

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51. Eliseeva, S.V. and Bunzli, J.C., Lanthanide luminescence for functional

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53. Faustino, W.M., Malta, O.L. and de Sa, G.F., Intramolecular energy transfer

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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

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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-

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Eu(SiW11O39)2 13- and Eu(BW11O39)2 15-. Inorganica Chimica Acta, 1984.

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63. Griffith, W.P., Moreea, R.G.H. and Nogueira, H.I.S., Lanthanide complexes as

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II

Research work

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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)

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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.

300 400 500 600 700 800

0.6

0.7

0.8

0.9

1.0

1.1

Ab

s

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]

.

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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.

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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

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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.

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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

7.50 mL cyclohexane

140 µL RBITC aqueous

solution (0.1 M)

0.1 mL APTES

27.5 mg RBITC

2.5 mL anhydrous ethanol

RBITC aqueous solution

(0.1 M), with equimolar

quantity of APTES in water

Homogenize for 30 min React 24h in dark React overnight

Add:

100 µL TEOS

60 µL ammonium hydroxide


5.40 mL n-hexanol

22.50 mL cyclohexane

3 mg RBITC-APTES

(dissolved in deionized water

- 1.5 mL)

300 µL ammonium hydroxide


1.80 mL n-hexanol

7.50 mL cyclohexane

400 µL water

100 µL RBITC-APTES

aqueous solution

24h reaction Homogenize for 30 min Homogenize for 60 min

Washing Add:

300 µL TEOS

Add:

200 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.

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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).

550 575 600 625 650 675 700

0

200

400

600

800

1000

Inte

nsit

y (

a.u

)

Wavelength (nm)

A

C B

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.

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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.




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.





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.

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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

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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

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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

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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


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

http://imagej.nih.gov/ij

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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-

APTES@SiO2 was modified by a grafting methodology adapted from literature.[11] 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


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


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).

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105

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


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).

Sample Measurement Hydrodynamic

diameter (nm)

Polydispersity

index

Standard

deviation (±)

RBITC-APTES

FSNPs

1 95.83 0.238 46.75

2 102.9 0.206 46.70

3 96.65 0.211 44.39

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

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DLS measurement is often much larger than the TEM size and can have a large

variability in the particle size (polydispersity index >0.1).

0 20 40 60 80 100 120 140 160 180 200

0

5

10

15

20

25

Nu

mb

er

of

pa

rtic

les

(%

)

Hydrodynamic diameter (nm)

Size: 78.82 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]

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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.

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).

350 400 450 500 550 600 650 700

0.00

0.02

0.04

0.06

0.08

0.10

Ab

s

Wavelength (nm)

RBITC

FSNPs

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109

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

350 400 450 500 550 600 650 700

0.00

0.01

0.02

0.03

0.04

0.05

Ab

s

Wavelength (nm)

RBITC

RBITC-APTES

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

110 FCUP


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

Sample Size

(nm)

Average NP

volume

(cm3)

NP

density

(g/cm3)

[NPs]

(g/L)

NPs in

suspension

RBITC

molecules in

suspension

RBITC

molecules

per NP

a 64.4

1.40x10-16 2.2

1.6 5.21x1015 7.88x1017 151

b 64.4 0.8 2.60x1015 3.79x1017 146

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

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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.

360 400 440 480 520 560 600

0

200

400

600

800

1000 RBITC

RBITC-APTES

FSNPs

Inte

nsit

y (

a.u

.)

Wavelength (nm)

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

absolute ethanol.

112 FCUP


540 560 580 600 620 640 660 680 700

0

200

400

600

800

1000 RBITC

RBITC-APTES

FSNPs

Inte

nsit

y (

a.u

.)

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

quantum yield. Rhodamine b (ΦF = 0.49 in ethanol[17]) 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

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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


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

FSNPs 0.34 ---

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]

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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

Sample

FIT

Τ1 (ns) Τ2 (ns) Χ2

RBITC 2.44 --- 1.16

RBITC-APTES 2.99 1.04 1.20

RBITC-APTES@SiO2

FSNPs 3.44 1.27 1.12

Ads:RBITC-APTES@SiO2

FSNPs

2.66 0.87 1.35

116 FCUP


200 400 600 800 1000

10

100

1000L

og

10 I

Time (ns)

A

T(RBITC) = 2.44

200 400 600 800 1000

10

100

1000T

2(RBITC-APTES) = 1.04

Lo

g10 I

time (ns)

B

T1(RBITC-APTES) = 2.99

200 400 600 800 1000

10

100

1000 T2(RBITC-APTES@SiO

2) = 1.27

Lo

g10 I

Time (ns)

C

T1(RBITC-APTES@SiO

2) = 3.44

200 400 600 800 1000

10

100

1000

T2(Ads:RBITC-APTES@SiO

2) = 0.87

Lo

g10 I

Time (ns)

D

T1(Ads:RBITC-APTES@SiO

2) = 2.66

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

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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).

O N+

CH3

CH3

NH3C

CH3

NCS

COOH

O N+

CH3

CH3

NH3C

CH3

COOH

SCN

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

rhodamine b 6-isothiocyanate.

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O N+

CH3

CH3

NH3C

CH3

NH

COOH

C NH

Si

O

O

O

S

Si O

O

O

NH C NH

O N+

CH3

CH3

NH3C

CH3

COOH

S

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

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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

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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.

580 590 600 610 6200.00

0.05

0.10

0.15

0.20

0.25

An

iso

tro

py r

Wavelength (nm)

RBITC

Ads:RBITC-APTES@SiO2 FSNPs

RBITC-APTES@SiO2 FSNPs

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@SiO2

FSNPs 0.187 0.009 2.76 0.048

Ads:RBITC-APTES@SiO2 FSNPs

0.172 0.007 2.19 0.074

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

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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

within a cone of semiangle θc (Figure 4.13).[24] 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).

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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.

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300 400 500 600 700

0.00

0.25

0.50

300 400 500 600 700

-0.05

0.00

0.05

0.10

Ab

s

Wavelength (nm)

FSNPs-GPTES-DNA

FSNPs-GPTES

Ab

s

Wavelength (nm)

FSNPs-GPTES

FSNPs-GPTES-DNA

DNA

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

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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

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125

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


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.

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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.





2007. 3(4): p. 266-272.

5. Santra, S., Zhang, P., Wang, K.M., Tapec, R. and Tan, W.H., Conjugation of



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

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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

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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).




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.

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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.

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5.1.2.4. Transmission electron microscopy


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


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

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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

Based on the emission spectra, 5D0 life times and empirical radiative and non-

radiative transition rates, the 5D0 quantum efficiency, q,[9-11] was determined for

(Eu(PW11)2) and Eu(PW11)2@SiO2. Assuming that only non-radiative and radiative

processes are involved in the depopulation of the 5D0 state, q is given by:

where kr and knr are the radiative and non-radiative transition probabilities, respectively,

and kexp=τexp–1 (kr + knr) is the experimental transition probability. The emission intensity,

I, taken as the integrated intensity S of the emission lines for the 5D07F0-6 transitions,

is given by:

where i and j represent the initial (5D0) and final (7F0-6) levels, respectively, is the

transition energy, the Einstein coefficient of spontaneous emission and the

population of the 5D0 emitting level.[9-11] Because the 5D0 7F5,6 transitions are not

observed experimentally, their influence on the depopulation of the 5D0 excited state

may be neglected and, thus, the radiative contribution is estimated based only on the

relative intensities of the 5D0 7F0-4 transitions. The emission integrated intensity, S, of

the 5D07F0-4 transitions has been measured for compounds Eu(PW11)2 and

Eu(PW11)2@SiO2 at 298 K. Because the 5D07F1 transition does not depend on the

local ligand field seen by the Eu3+ ions (due to its dipolar magnetic nature) it may be

used as a reference for the whole spectrum, in vacuum A(5D07F1)=14.65 s–1,[12] and kr

is given by:

nrr

r

kk

kq

jiijijiji SNAwI

jiw

jiAiN

4

0 0

0

10

1010

J J

Jr

S

SAk

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where A0-1 is the Einstein coefficient of spontaneous emission between the 5D0 and the

7F1 levels. An average index of refraction of 1.5 was considered for both samples,

leading to A(5D07F1) 50 s–1.[13] 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

by reported procedures.[17, 18] Then, a solution of EuCl3.6H2O was added dropwise to

the aqueous solution of PW11. The mixture was stirred for 1 h at 90 ºC. PW11 and Eu3+

were solubilized in the minimum amount of water, and the solutions were added in the

rigorously stoichiometric amounts of PW11 and Eu3+ 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.

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137

5.1.5. Functionalization of Eu(PW11)2@SiO2

The surface of Eu(PW11)2@SiO2 was modified by a grafting methodology described

by Balula et al.[22] 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.


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

the mono-substituted [PW11O39Eu(H2O)3]4- and the sandwich-type [Eu(PW11O39)2]

11-

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

potassium salt of [Eu(PW11O39)2]11-. 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


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]

Figure 5.1 - (a) The structures of the sandwich type europium-phosphotungstate anion, [Eu(PW11O39)2]11−

; (b) its {EuO8}

coordination center displaying a square-antiprismatic geometry and (c) the mono-substituted europium-

phosphotungstate anion, [PW11Eu(H2O)3O39]4- 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-

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139

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

determined for the first time.[23] Eu(PW11)2 crystallized in the monoclinic system with the

respective structure solved in the P21/c space group, and revealed an asymmetric unit

with one [Eu(PW11O39)2]11- anion, 11 K+ cations distributed over 14 positions and a large

number of crystallization water molecules. The europium-polyoxometalate anion

reveals one Eu3+ centre linked to two mono-lacunary Keggin units [PW11O39]7- (Figure

5.1). As reported for similar lanthano-polyoxometalate anions based compounds,

[Ln(PW11O39)2]11-, the Eu3+ is coordinated by eight lacunary oxygen atoms belonging to

two Keggin fragments leading to an eight coordinated centre, {EuO8}.[14, 24-28] The

coordination environment resembles a square-antiprismatic geometry (pseudo-D4d

symmetry), due to the relative rotation of the two [PW11O39]7- 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

ranges, respectively. In fact, the main structural characteristics of the [Eu(PW11O39)2]11-

anion are comparable to those already reported for related compounds.[14, 24]

In the extended crystalline arrangement, the sandwich-type europium-

polyoxometalate anions [Eu(PW11O39)2]11− 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


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

ρcalc (g cm−3

) 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

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141

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

and Eu(PW11)2 respectively.

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.

80

85

90

95

100

0 200 400 600 800

Weig

ht lo

ss (

%)

Temperature ºC

142 FCUP


Figure 5.3 - 31

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-

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

0.3

6

-10

.10

PW11

EuPW11

Eu(PW11

)2

(ppm)

5.5

8

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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.

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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.

0.0 2.5 5.0 7.5 10.0 12.5

0

100

200

300

400

500

W

CuW

EuEu W

Co

un

ts

KeV

CO

CuW P EuK

Si

Cu

EuPW11

@SiO2

0.0 2.5 5.0 7.5 10.0 12.5

0

100

200

300

400

500

CuW WEuW

Cu

W

Co

un

ts

KeV

C

O

Si

CuW

P K Eu

Eu(PW11

)2@SiO

2

mailto:[email protected]

mailto:[email protected]

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145

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).

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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.

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147

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


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

Keggin-type frameworks: as(P-O) between 1100-1040 cm-1, terminal as(W-Ot) near

950 cm-1, corner-sharing as(W-Ob-W) near 850 cm-1 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

the Keggin derivative compounds (400-1100 cm-1, Figure 5.10): as(Si-O-Si), s(Si-O-

Si) and (O-Si-O).[33] 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.

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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

corresponding to the corner-sharing as(W-Ob-W) stretches.[30, 32] 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).

1800 1600 1400 1200 1000 800 600 400

Wavenumber (cm-1)

EuPW11

EuPW11

@SiO2

1800 1600 1400 1200 1000 800 600 400

Eu(PW11

)2@GPTEsSiO

2

Wavenumber (cm-1)

Eu(PW11

)2

Eu(PW11

)2@SiO

2

Eu(PW11

)2@CPTEsSiO

2

B A

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The solid state 31P NMR spectra of EuPW11 and Eu(PW11)2 potassium salts and

corresponding silica-coated core:shell nanoparticles are shown in Figure 5.12.

Figure 5.12 - Solid state 31

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

caused by the slight asymmetry of the two [PW11O39]7- units that surround the europium

ion in the sandwich compound.[29, 34] 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

[PW11O39]7- 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

70 60 50 40 30 20 10 0 -10 -20 -30 -40 -50 -60 -70

(ppm)

EuPW11

EuPW11

@SiO2

PW11

70 60 50 40 30 20 10 0 -10 -20 -30 -40 -50 -60 -70

(ppm)

Eu(PW11

)2

Eu(PW11

)2@SiO

2

Eu(PW11

)2@GPTEsSiO

2

Eu(PW11

)2@CPTEsSiO

2

A B

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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

of the Eu3+ intra-4f6 transitions, namely 7F0,15D4-1,

5L6 and 5G2-6. 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

related Eu(SiW11)2 compound.[38] 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.

250 300 350 400 450 500 550

LMCT

7F

0

5D

4

7F

0

5D

1

7F

0

5D

2

7F

0

5L

6

7F

0

5G

2-6

EuPW11

@ SiO2

Wavelength (nm)

EuPW11

A

250 300 350 400 450 500 550

LMCT

7F

0

5D

4

7F

0

5D

1

7F

0

5D

2

7F

0

5L

6

7F

0

5G

2-6

Eu(PW11

)2@ SiO

2

Wavelength (nm)

Eu(PW11

)2

B

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


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.

580 600 620 640 660 680 700

5D

0

7F

4

5D

0

7F

3

5D

0

7F

2

5D

0

7F

1

EuPW11

@ SiO2

EuPW11

5D

0

7F

0

A

Wavelength (nm)

580 600 620 640 660 680 700

Eu(PW11

)2@ SiO

2

Eu(PW11

)2

5D

0

7F

4

5D

0

7F

3

5D

0

7F

2

5D

0

7F

1

5D

0

7F

0

B

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-

degenerate 5D0 state and the 7F0-4 levels of the fundamental Eu3+ septet. Except for

5D07F1, which has a predominant magnetic-dipole character independent of the Eu3+

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

without high vacuum was very similar to that of Eu(SiW11)2.[38] In particular the splitting

of the 5D07F1 transition into three Stark components increased greatly when the

sample was exposed to the high vacuum. For this compound, the splitting of the 7F0,1

levels into one and three Stark components, respective, the predominance of the

5D07F2 transition relative to the 5D0

7F1 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

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

integrated intensities of the 5D0→7F2 and 5D0→

7F1 transitions, )()( 1

70

52

70

5 FDFDII , also

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

centre for which the 5D07F2 transition is absent). Thus, the observed changes in the

room-temperature emission spectrum with the application of a high vacuum of ca.

5×10-6mbar (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.

0 5 10 15 20 25

EuPW11

L = 2.22 ± 0.03 ms

EuPW11

S = 0.33 ± 0.01 ms

Ln I

Time (ms)

EuPW11

@ SiO2S = 0.28 ± 0.01 ms

EuPW11

@ SiO2L = 2.39 ± 0.07 ms

A

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0 5 10 15 20 25

B

Eu(PW11

)2 = 2.46 ± 0.02 ms

Ln I

Time (ms)

Eu(PW11

)2@ SiO

2 = 2.30 ± 0.02 ms

Figure 5.15 - Eu3+

5D0 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.

Based on the emission spectra, 5D0 lifetimes and empirical radiative and non-

radiative transition rates, and assuming that only non-radiative and radiative processes

are involved in the depopulation of the 5D0 state, the 5D0 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

Eu3+ 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.

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

5D0 quantum efficiency,

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

Compound τ [ms] kr [s-1] Knr [s

-1] q [%]

Eu(PW11)2 2.46±0.02 200 207 49

Eu(PW11)2@SiO2 2.30±0.02 241 194 55

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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

mediated growth mechanism.[35, 36] 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

improvement of their 5D0 quantum efficiency for the capped silica compound.

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33. Luo, X.J. and Yang, C., Photochromic ordered mesoporous hybrid materials

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34. Griffith, W.P., Morley-Smith, N., Nogueira, H.I.S., Shoair, A.G.F., Suriaatmaja,

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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.

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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

http://www.moleculardevices.com/Applications/Cellular-Function/Cell-Health.html

http://en.wikipedia.org/wiki/MTT_assay

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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.

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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.

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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


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.


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

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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

(SiO2 NPs), europium salt (Eu3+), 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

http://en.wikipedia.org/wiki/Heterogeneous

http://en.wikipedia.org/wiki/Human

http://en.wikipedia.org/wiki/Epithelium

http://en.wikipedia.org/wiki/Colon_%28anatomy%29

http://en.wikipedia.org/wiki/Adenocarcinoma

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(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.

350 400 450 500 550 600 650 700 750

0

200

400

600

800

1000

RBITC-APTES@SiO2

in DMEM

RBITC-APTES@SiO2

in DMEM

RBITC-APTES@SiO2

stock solution

Inte

nsity (

a.u

.)

Wavelength (nm)

Excitation Emission

RBITC-APTES@SiO2

stock solution

A

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500 525 550 575 600 625 650

0

5

10

15

20

Inte

nsity (

a.u

.)

Wavelength (nm)

RBITC-APTES@SiO2 3.2 ug/mL in DMEM

Excitation Emission

B

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),

RBITC, europium salt (Eu3+) 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.

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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.

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).

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).

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

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A

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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

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

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R B IT C

S iO 2 N P s

A

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

2 5

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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

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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,

Eu(PW11O39)2, Eu3+ 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.

Figure 6.6 - Effect of Eu(POMs)@SiO2 NPs, Eu3+

, 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).

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

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A

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

E u3 +

P O M s

S iO 2 N P s

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, 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



, 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


Similar to the results obtained for RBITC@SiO2, RBITC and SiO2 NPs (presented in

the previous section 6.3.2.1) are the results of Eu(PW11O39)2, Eu3+ salt

Eu(PW11O39)2@SiO2 NPs, and SiO2 NPs on cell esterase activity in the three cell lines

used.

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

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E u3 +

P O M s

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S iO 2 N P s

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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,

Eu3+ 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

cells of a POM ([Co4(H2O)2(PW9O34)2]10-) 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.

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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).

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Figure 6.10 - Representative phase contrast microscopy images of SH-SY5Y cells at 24 hours after incubation with SiO2


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Figure 6.11 - Representative phase contrast microscopy images of Hepa RG cells at 24 hours after incubation with SiO2


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.

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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


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Figure 6.14 - Representative phase contrast microscopy images of Hepa RGcells at 24 hours after incubation with SiO2


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

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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.

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III

Concluding Remarks and Perspectives

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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

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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

showed by 31P 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

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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

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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.

development of fluorescent silica ddevelopment of fluorescent silica nanoparticles encapsulating...

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