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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
D
D
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D
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
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“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
3
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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person along this journey, but nonetheless you’ll never gave up on me. You have been
a supporting rock in all the bad moments and in the good ones you have been the most
cheerful person. Thank you for believing in me when I doubted, for being a friend, a
listener, a supporter and most important my companion for life.
To all of those who contributed somehow to this thesis and prevented me to go
insane and aren't mentioned here, I appreciate your effort and caring. I leave you a
sincere thanks and the following quote:
“Those who pass us by do not go alone, do not leave us alone. Leave a bit of
themself, take a little of us.” (Antoine de Saint-Exupéry)
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AUTHOR PUBLICATIONS CONTAINING WORK RELATED WITH THIS THESIS:
Articles and book chapters in peer-reviewed journals
Published/submitted papers:
Cristina S. Neves, Carlos M. Granadeiro, Luis Cunha-Silva, Duarte Ananias,
Sandra Gago, Gabriel Feio, Patricia A. Carvalho, Peter Eaton, Salete S. Balula, Eulália
Pereira, Europium-polyoxometalates encapsulated into sílica nanoparticles:
characterization and photoluminescence studies, European Journal of Inorganic
Chemistry 16, 2877-2886 (2013).
Book chapter:
Pedro V. Baptista, Gonçalo Doria, Pedro Quaresma, Miguel Cavadas, Cristina S.
Neves, Inês Gomes, Peter Eaton, Eulália Pereira, Ricardo Franco, Nanoparticles in
Molecular Diagnosis, Progress in Molecular Biology and Translational Science, Vol.
104, 427-488, (Antonio Villaverde, Ed.) Elsevier (2011).
Awards:
Young Scientist Award on European Materials Research Society (E-MRS) 2013
Spring Meeting, Strasbourg, France, 26th – 31st May 2013.
Best Paper Award on XI International Conference on Nanostructured Materials,
Rhodes, Greece, 23rd – 31st August 2012.
Articles not included in this dissertation, but published in the course of the
Ph.D. work:
Silvia C. Lopes, Cristina Neves, Peter Eaton, Paula Gameiro, Improved model
systems for bacterial membranes from differing species: the importance of varying
composition in PE/PG/cardiolipin ternary mixtures, Molecular Membrane Biology, 29
(2012), 207-217.
Maria J. Medeiros, Cristina S. S. Neves, A. R. Pereira, Elizabeth Duñach,
Electroreductive intramolecular cyclisation of bromoalkoxylated derivatives catalized by
nickel (I) tetrametylcyclam in “green” media, Electrochimica Acta, 56 (2011), 4498-
4503.
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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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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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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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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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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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5.1.2.1. Elemental analysis .................................................................................... 132
5.1.2.2. Vibrational Spectroscopy .......................................................................... 132
5.1.2.3. Solid state NMR ........................................................................................ 132
5.1.2.4. Transmission electron microscopy ............................................................ 133
5.1.2.5. Scanning electron microscopy .................................................................. 133
5.1.2.6. Dynamic light scattering ............................................................................ 133
5.1.2.7. Atomic force microscopy ........................................................................... 133
5.1.2.8. X-ray crystallography ................................................................................ 134
5.1.2.9. Photoluminescence and lifetime measurements ....................................... 134
5.1.2.10. Quantum efficiency ................................................................................... 135
5.1.3. Synthesis of europium polyoxometalates Eu(PW11)x (x = 1 and 2) .......... 136
5.1.4. Encapsulation of Eu(PW11)x (x = 1 and 2) into silica nanoparticles .......... 136
5.1.5. Functionalization of Eu(PW11)2@SiO2 ..................................................... 137
5.2. Results and Discussion ................................................................................. 137
5.2.1. Characterization of Eu(PW11)x compounds ............................................. 138
5.2.1.1. X-ray crystallography ................................................................................ 138
5.2.1.2. Thermogravimetry ..................................................................................... 140
5.2.1.3. 31P NMR spectroscopy .............................................................................. 141
5.2.2. Characterization of Eu(PW11)x@SiO2 nanoparticles ................................ 142
5.2.2.1. Transmission Electron Microscopy ............................................................ 143
5.2.2.1. Scanning Electron Microscopy .................................................................. 146
5.2.2.2. Characterization by vibrational spectroscopy ............................................ 148
5.2.2.3. Photoluminescence properties .................................................................. 151
5.2.3. Conclusions ............................................................................................ 155
5.3. References .................................................................................................... 156
6. Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles .... 161
6.1. Cytotoxicity assays ........................................................................................ 162
6.2. Materials and Methods .................................................................................. 163
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6.2.1. Chemicals ............................................................................................... 163
6.2.2. Cellular culture ........................................................................................ 164
6.2.3. Nanoparticle uptake ................................................................................ 164
6.2.4. Cell viability by Calcein-AM assay .......................................................... 164
6.2.5. Phase contrast microscopy ..................................................................... 165
6.2.6. Fluorescence spectroscopy .................................................................... 165
6.2.7. Transmission electron microscopy .......................................................... 165
6.2.8. Statistical analysis .................................................................................. 165
6.3. Results and Discussion ................................................................................. 165
6.3.1. Cellular uptake of silica nanoparticles ..................................................... 166
6.3.2. Cell esterase activity (Calcein-AM assay) ............................................... 168
6.3.2.1. Effect of RBITC@SiO2 NPs on Caco-2, SH-SY5Y and Hepa RG cells
viability ................................................................................................................. 169
6.3.2.2. Effect of Eu(PW11O39)2@SiO2 NPs on Caco-2, SH-SY5Y and Hepa RG cells
viability ................................................................................................................. 172
6.3.3. Morphological analysis by phase contrast microscopy ............................ 174
6.3.3.1. RBITC-APTES FSNPS ............................................................................. 174
6.3.3.2. Eu(PW11O39)2@ SiO2 NPs ........................................................................ 177
6.4. Conclusions ................................................................................................... 179
6.5. References .................................................................................................... 181
III – Concluding Remarks and Perspectives
Concluding Remarks and Perspectives..................................................................... 187
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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
II – Research work
Table EB 1– Comparison between the three methods followed to prepare fluorescent
silica NPs using the microemulsion technique............................................................. 95
Table 4.1 - Average hydrodynamic diameter of RBITC FSNPs measured by DLS (by
percentage of number of particles, measurements were repeated 5 times for each
sample). .................................................................................................................... 106
Table 4.2 – Amount of RBITC dye molecules per fluorescent silica nanoparticle ...... 110
Table 4.3 - Fluorescence quantum yields of RBITC, RBITC-APTES conjugate and
FSNPs ...................................................................................................................... 113
Table 4.4 - Lifetime data of RBITC and fluorescent silica nanoparticles (FSNPs) in
absolute ethanol ....................................................................................................... 115
Table 4.5 - Anisotropy (r) and rotational diffusion coefficient (Dr) values of RBITC and
fluorescent silica nanoparticles (FSNPs) adsorbed and covalently bound to silica NPs
in absolute ethanol. ................................................................................................... 120
Table 4.6 – Zeta potential ζ of FSNPs-GPTES and FSNPs-GPTES-DNA ................. 123
Table 5.1 - Crystal and structure refinement data for Eu(PW11)2 ............................... 140
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
III – Concluding Remarks and Perspectives
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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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II – Research work
Figure EB 1 - UV-vis spectrum of fluorescent silica nanoparticles synthesized by
Stober’s method through the adapted procedure described by Bringley[1]. .................. 91
Figure EB 2- TEM images of TRICT fluorescent silica nanoparticles prepared by
Stöber’s method following a similar procedure to that described by Larson[3] et al. ..... 93
Figure EB 3 - TEM images of fluorescent silica nanoparticles prepared by the reverse
microemulsion system following the procedures of Gao[4] (A); Zhang[5] (B) and Shi[6] (C).
................................................................................................................................... 96
Figure EB 4 - Fluorescence emission spectra of fluorescent silica nanoparticles
prepared by the reverse microemulsion system following the procedures of Gao[4] (A);
Zhang[5] (B) and Shi[6] (C). ........................................................................................... 96
Figure 4.1 - TEM images of RBITC-APTES FSNPS nanoparticles and corresponding
size distribution histogram......................................................................................... 105
Figure 4.2 - SEM images of RBITC-APTES FSNPs showing the spherical morphology
of the NPs. ................................................................................................................ 105
Figure 4.3 - DLS hydrodynamic diameter distribution statistics graph (by percentage of
number of particles) for RBITC FSNPS. Error bars show standard deviation of five
different measurements. ........................................................................................... 107
Figure 4.4 - UV-vis spectra of RBITC and RBITC fluorescent silica NPs (FSNPs) in
ethanol at 25 ºC. UV-vis spectrum of FSNPs was fitted using a 2nd order exponential
decay to remove silica scattering. Both samples were dissolved to a final concentration
with almost the same absorbance (0.09). .................................................................. 108
Figure 4.5 - UV-vis spectra of RBITC and RBITC-APTES conjugate in ethanol at 25 ºC.
................................................................................................................................. 109
Figure 4.6 - Fluorescence excitation spectra of RBITC, RBITC-APTES conjugate and
FSNPs recorded at 25 ºC in absolute ethanol. .......................................................... 111
Figure 4.7 - Fluorescence emission spectra of RBITC, RBITC-APTES conjugate and
FSNPs recorded at 25 ºC in absolute ethanol. .......................................................... 112
Figure 4.8 - (A) RBITC doped fluorescent silica NPs prepared by hydrolysis and
polymerization of TEOS in a microemulsion method; (B) bare silica NPs with RBITC
dye molecules adsorbed onto the nanoparticle’s surface; (C) fluorescent core-shell NPs
with a silicon core and a shell of RBITC dye molecules and TEOS. .......................... 113
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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
Figure 6.4 - Effect of RBITC-APTES@SiO2 NPs, RBITC and SiO2 NPs on esterase
activity of human neuroblastoma SH-SY5Y cells, as assessed by the calcein-AM
assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage
of control (vehicle-treated cells) and data are presented as mean ± SEM (n=4-28 per
group). ...................................................................................................................... 169
Figure 6.5 - Effect of RBITC-APTES@SiO2 NPs, RBITC and SiO2 NPs on esterase
activity of human hepatoma Hepa RG cells, as assessed by the calcein-AM assay, at
24 h (A) and 48 h (B) after exposure. Results were calculated as percentage of control
(vehicle-treated cells) and data are presented as mean ± SEM (n=8-28 per group). . 170
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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25
assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage
of control (vehicle-treated cells) and data are presented as mean ± SEM (n=8-24 per
group). ...................................................................................................................... 173
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
assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated as percentage
of control (vehicle-treated cells) and data are presented as mean ± SEM (n=8-28 per
group). ...................................................................................................................... 173
Figure 6.9 - Representative phase contrast microscopy images of Caco-2 cells at 24
hours after incubation with SiO2 nanoparticles, RBITC dye and RBITC-APTES@SiO2
nanoparticles (100x magnification). ........................................................................... 175
Figure 6.10 - Representative phase contrast microscopy images of SH-SY5Y cells at
24 hours after incubation with SiO2 nanoparticles, RBITC dye and RBITC-
APTES@SiO2 nanoparticles (100x magnification). ................................................... 176
Figure 6.11 - Representative phase contrast microscopy images of Hepa RG cells at
24 hours after incubation with SiO2 nanoparticles, RBITC dye and RBITC-
APTES@SiO2 nanoparticles (100x magnification). ................................................... 177
Figure 6.12 - Representative phase contrast microscopy images of Caco-2 cells at 24
hours after incubation with SiO2 nanoparticles, Eu salt, Eu(PW11O39)2 and
Eu(PW11O39)2@SiO2 nanoparticles (100x magnification). .......................................... 178
Figure 6.13 - Representative phase contrast microscopy images of SH-SY5Ycells at 24
hours after incubation with SiO2 nanoparticles, Eu salt, Eu(PW11O39)2 and
Eu(PW11O39)2@SiO2 nanoparticles (100x magnification). .......................................... 178
Figure 6.14 - Representative phase contrast microscopy images of Hepa RGcells at 24
hours after incubation with SiO2 nanoparticles, Eu salt, Eu(PW11O39)2 and
Eu(PW11O39)2@SiO2 nanoparticles (100x magnification). .......................................... 179
III – Concluding Remarks and Perspectives
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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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28 FCUP
Abbreviations and Symbols
GFP - green fluorescent protein
GPTMS - 3-glycidoxypropyltrimethoxy silane
HBSS – Hank’s balanced salt solution
HC – high component lifetime
HEPA RG - Human hepatoma cells
HepG2 – Human hepatocellular liver carcinoma cells
ICP-MS – Inductively coupled plasma mass spectroscopy
LC – low component lifetime
LMCT- ligand-to-metal charge transfer
Ln - lanthanide
LnPOMs - lanthanide-substituted polyoxometalates
MAS – Magic-angle spinning
MB - methylene blue
MLCT - metal-to-ligand charge transfer
MOF - metal-organic-framework
MPTS - 3-mercaptopropyltrimethoxysilane
MRI – magnetic resonance imaging
MTT - 3-(4, 5-dimethyl-2-thiazolyl)-2, 5-diphenyl-2H-tetrazolium bromide
NMR – nuclear magnetic resonance
NP - nanoparticle
NPs – nanoparticles
ODS – oxidative desulfurization
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Abbreviations and Symbols
29
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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Abbreviations and Symbols
TRITC - tetramethylrhodamine isothiocyanate
UV-vis – ultraviolet-visible spectroscopy
W/O - water-in-oil microemulsion
W0 - water-to-surfactant molar ratio
ε – absorption coefficient
ζ - zeta potential
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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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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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An overview of nanoscale fluorescent materials
35
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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An overview of nanoscale fluorescent materials
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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An overview of nanoscale fluorescent materials
37
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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39
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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43
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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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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47
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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49
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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55
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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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
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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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24. Ruedas-Rama, M.J., Walters, J.D., Orte, A. and Hall, E.A.H., Fluorescent
nanoparticles for intracellular sensing: A review. Analytica Chimica Acta, 2012.
751: p. 1-23.
25. Santra, S., Zhang, P., Wang, K.M., Tapec, R. and Tan, W.H., Conjugation of
biomolecules with luminophore-doped silica nanoparticles for photostable
biomarkers. Analytical Chemistry, 2001. 73(20): p. 4988-4993.
26. Peng, J., Wang, K., Tan, W., He, X., He, C., Wu, P. and Liu, F., Identification of
live liver cancer cells in a mixed cell system using galactose-conjugated
fluorescent nanoparticles. Talanta, 2007. 71(2): p. 833-40.
27. Ow, H., Larson, D.R., Srivastava, M., Baird, B.A., Webb, W.W. and Wiesner, U.,
Bright and stable core-shell fluorescent silica nanoparticles. Nano Letters, 2005.
5(1): p. 113-7.
28. Shi, H., He, X.X., Wang, K.M., Yuan, Y., Deng, K., Chen, J.Y. and Tan, W.H.,
Rhodamine B isothiocyanate doped silica-coated fluorescent nanoparticles
(RBITC-DSFNPs)-based bioprobes conjugated to Annexin V for apoptosis
detection and imaging. Nanomedicine-Nanotechnology Biology and Medicine,
2007. 3(4): p. 266-272.
29. Xu, H., Aylott, J.W., Kopelman, R., Miller, T.J. and Philbert, M.A., A real-time
ratiometric method for the determination of molecular oxygen inside living cells
using sol-gel-based spherical optical nanosensors with applications to rat C6
glioma. Analytical Chemistry, 2001. 73(17): p. 4124-4133.
30. Burns, A., Sengupta, P., Zedayko, T., Baird, B. and Wiesner, U., Core/Shell
fluorescent silica nanoparticles for chemical sensing: towards single-particle
laboratories. Small, 2006. 2(6): p. 723-6.
31. Peng, J.F., He, X.X., Wang, K.M., Tan, W.H., Wang, Y. and Liu, Y.,
Noninvasive monitoring of intracellular pH change induced by drug stimulation
using silica nanoparticle sensors. Analytical and Bioanalytical Chemistry, 2007.
388(3): p. 645-654.
32. Wang, L., Yang, C. and Tan, W., Dual-luminophore-doped silica nanoparticles
for multiplexed signaling. Nano Letters, 2005. 5(1): p. 37-43.
33. Zhao, X.J., Tapec-Dytioco, R. and Tan, W.H., Ultrasensitive DNA detection
using highly fluorescent bioconjugated nanoparticles. Journal of the American
Chemical Society, 2003. 125(38): p. 11474-11475.
34. Zhou, X.C. and Zhou, J.Z., Improving the signal sensitivity and photostability of
DNA hybridizations on microarrays by using dye-doped core-shell silica
nanoparticles. Analytical Chemistry, 2004. 76(18): p. 5302-5312.
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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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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
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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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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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Lanthanopolyoxometalates encapsulated into silica nanoparticles
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
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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
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Lanthanopolyoxometalates encapsulated into silica nanoparticles
the sum of the angular momenta of the electrons in the initial and final states must
change by an odd integer.[51] That is the case of electronic transitions between f orbitals
in lanthanides ions. As a result lanthanide ions suffer from weak light absorption and
because of the low molar absorption coefficients ε (smaller than 10 L mol-1 cm-1) only a
very limited amount of radiation is absorbed by direct excitation in the 4f levels.[49]
Table 3.1 - Commonly observed emission bands of the lanthanide ions Eu3+
, Tb3+
, Nd3+
, Er3+
and Yb3+
in solution.
(Adapted from Werts[50]
)
Ion Transition λemission (nm) Ion Transition λemission (nm)
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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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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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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93
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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95
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
5.33 mL Triton X-100
5.40 mL n-hexanol
22.50 mL cyclohexane
3 mg RBITC-APTES
(dissolved in deionized water
- 1.5 mL)
300 µL ammonium hydroxide
1.77 mL Triton X-100
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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97
References
1. Bringley, J.F., Penner, T.L., Wang, R., Harder, J.F., Harrison, W.J. and
Buonemani, L., Silica nanoparticles encapsulating near-infrared emissive
cyanine dyes. Journal of Colloid and Interface Science, 2008. 320(1): p. 132-9.
2. Rossi, L.M., Shi, L., Quina, F.H. and Rosenzweig, Z., Stober synthesis of
monodispersed luminescent silica nanoparticles for bioanalytical assays.
Langmuir, 2005. 21(10): p. 4277-80.
3. Larson, D.R., Ow, H., Vishwasrao, H.D., Heikal, A.A., Wiesner, U. and Webb,
W.W., Silica nanoparticle architecture determines radiative properties of
encapsulated fluorophores. Chemistry of Materials, 2008. 20(8): p. 2677-2684.
4. Gao, F., Wang, L., Tang, L. and Zhu, C., A Novel Nano-Sensor Based on
Rhodamine-b-Isothiocyanate –Doped Silica Nanoparticle for pH Measurement.
Microchimica Acta, 2005. 152: p. 131-135.
5. Zhang, R.R., Wu, C.L., Tong, L.L., Tang, B. and Xu, Q.H., Multifunctional Core-
Shell Nanoparticles as Highly Efficient Imaging and Photosensitizing Agents.
Langmuir, 2009. 25(17): p. 10153-10158.
6. Shi, H., He, X.X., Wang, K.M., Yuan, Y., Deng, K., Chen, J.Y. and Tan, W.H.,
Rhodamine B isothiocyanate doped silica-coated fluorescent nanoparticles
(RBITC-DSFNPs)-based bioprobes conjugated to Annexin V for apoptosis
detection and imaging. Nanomedicine-Nanotechnology Biology and Medicine,
2007. 3(4): p. 266-272.
7. Ye, Z.Q., Tan, M.Q., Wang, G.L. and Yuan, J.L., Novel fluorescent europium
chelate-doped silica nanoparticles: preparation, characterization and time-
resolved fluorometric application. Journal of Materials Chemistry, 2004. 14(5):
p. 851-856.
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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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Dye doped fluorescent silica nanoparticles
101
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
triethoxysilane – APTES), and the reaction product was incorporated into silica spheres
by hydrolysis and polymerization of TEOS in alkaline media.
RBITC dye (1.0x10-4 mol) was prepared in aqueous solution and then mixed with an
equimolar quantity of APTES, reacting overnight at room temperature. The monomer
precursor RBITC-APTES was directly used to prepare silica-coated fluorescent
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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 =
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100). The mixture was left for incubation at room temperature for 24h under stirring.
Afterwards nanoparticles were centrifuged and washed with potassium phosphate
buffer for three times redispersed in buffer and storage at - 20ºC for further use.
4.2. Results and Discussion
Fluorescent silica nanoparticles (FSNPs) containing RBITC molecules were
synthesized using the reverse microemulsion technique for the alkaline hydrolysis of
TEOS. The dye was firstly coupled to a silane coupling agent (3-aminopropyl
triethoxysilane – APTES), and the reaction product was incorporated into silica spheres
by hydrolysis and polymerisation of TEOS in alkaline media. The obtained
nanoparticles (RBITC-APTES@SiO2) have a mean diameter of approximately 64 nm
with a spherical morphology and a narrow particle size distribution. The particle surface
was also modified, so that the particles could bind to biologically active molecules, such
as oligonucleotides. Lifetime measurements and steady-state anisotropy studies of the
nanoparticles and the free dye were also performed to evaluate the effect of the
encapsulation on the fluorescence emission properties of RBITC.
4.2.1 Characterization of RBITC@SiO2 nanoparticles
4.2.1.1. Characterization by electron microscopy
The size and shape of RBITC-APTES doped fluorescent silica nanoparticles were
measured from TEM images. Based upon the size (diameter) measurement of more
than 100 particles, the average nanoparticle diameter obtained was 64.4 nm (± 7.5 nm
standard deviation). TEM images (Figure 4.1) show that the dye doped fluorescent
silica nanoparticles have a spherical shape, are fairly monodisperse and size
distribution histogram demonstrates a narrow particle size distribution (Figure 4.1
bottom right).
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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.
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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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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.
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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.
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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
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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
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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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Dye doped fluorescent silica nanoparticles
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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Dye doped fluorescent silica nanoparticles
fluorophore molecule should undergo more restricted movement when bound inside
the NP.
Figure 4.12 shows the emission fluorescence anisotropy spectra as a function of
wavelength for RBITC and the FSNPs (dye covalently bound - RBITC-APTES@SiO2;
and dye adsorbed – Ads:RBITC-APTES@SiO2, to the silica matrix). Based on these
results and in the fluorescence lifetime the rotational diffusion coefficient (Dr) was
calculated for the free RBITC in solution and for the FSNPs. The values obtained are
presented in Table 4.5.
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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123
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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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
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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
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3. Fery-Forgues, S. and Lavabre, D., Are fluorescence quantum yields so tricky to
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4. Shi, H., He, X.X., Wang, K.M., Yuan, Y., Deng, K., Chen, J.Y. and Tan, W.H.,
Rhodamine B isothiocyanate doped silica-coated fluorescent nanoparticles
(RBITC-DSFNPs)-based bioprobes conjugated to Annexin V for apoptosis
detection and imaging. Nanomedicine-Nanotechnology Biology and Medicine,
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5. Santra, S., Zhang, P., Wang, K.M., Tapec, R. and Tan, W.H., Conjugation of
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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
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10. Santra, S., Wang, K., Tapec, R. and Tan, W., Development of novel dye-doped
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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-
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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):
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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.
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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.,
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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
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19. Rampazzo, E., Bonacchi, S., Montalti, M., Prodi, L. and Zaccheroni, N., Self-
organizing core-shell nanostructures: Spontaneous accumulation of dye in the
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20. Roy, S., Woolley, R., MacCraith, B.D. and McDonagh, C., Fluorescence lifetime
analysis and fluorescence correlation spectroscopy elucidate the internal
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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
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22. Yip, P., Karolin, J. and Birch, D.J.S., Fluorescence anisotropy metrology of
electrostatically and covalently labelled silica nanoparticles. Measurement
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23. Larson, D.R., Ow, H., Vishwasrao, H.D., Heikal, A.A., Wiesner, U. and Webb,
W.W., Silica nanoparticle architecture determines radiative properties of
encapsulated fluorophores. Chemistry of Materials, 2008. 20(8): p. 2677-2684.
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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
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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.
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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
TEM images were obtained using a HITACHI H-8100 instrument operating at an
acceleration voltage of 200 kV and energy dispersive X-ray spectroscopy (EDS)
analysis was performed on a ThermoNoran spectrometer. Samples for TEM analysis
were prepared by depositing ethanol suspensions of the nanoparticles on carbon
coated copper grids and allowing them to completely dry. TEM images were analysed
using image J software (version 1.44p), available through http://imagej.nih.gov/ij. TEM
was carried out in the institute of materials and surfaces science and engineering
(ICEMS) from Instituto Superior Técnico (IST).
5.1.2.5. Scanning electron microscopy
SEM analysis and EDS elemental mapping were performed on a Hitachi SU-70
instrument operating at an acceleration voltage of 30 kV. Samples for SEM analysis
were prepared by depositing ethanol suspensions of the nanoparticles on carbon
coated copper grids and allowing them to completely dry. Both techniques were carried
out in CICECO associated laboratory by Dr. Duarte Ananias.
5.1.2.6. Dynamic light scattering
DLS measurements were performed at 25ºC, using a Malvern ZetasizerNanoZS
compact scattering spectrometer with a 4.0 mW He-Ne laser (633 nm wavelength) at a
scattering angle of 173º. The average hydrodynamic diameter and the size distribution
of the samples were determined using Malvern Dispersion Technology Software 5.10.
All measurements were repeated five times to verify the reproducibility of the results.
Stable samples were prepared by dissolving the potassium salts of Eu(PW11)x in
Millipore water.
5.1.2.7. Atomic force microscopy
Samples for atomic force microscopy (AFM) were prepared by drying onto freshly
cleaved mica substrates from dilute aqueous solutions. AFM measurements were
made using an AFM Workshop TT-AFM instrument in vibrating (intermittent contact)
mode. A small (15 µm) scanner and low gains were used to ensure high resolution.
Probes from AppNano (ACT) with resonant frequency of around 300 kHz were used.
Images of around 2 µm x 2µm were acquired, and analysed using Gwyddion software
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Europium polyoxometalates encapsulated into silica nanoparticles
and a custom routine to measure accurately the height of nanoparticles. AFM
measurements were carried out by Dr. Peter Eaton.
5.1.2.8. X-ray crystallography
Crystalline K11[Eu(PW11O39)2.xH2O materials suitable for single-crystal X-ray
diffraction analysis were harvested and mounted in a CryoLoop using viscous oil.[1]
Diffraction data were collected at 150 K on a Bruker X8 Kappa APEX II charge-coupled
device (CCD) area-detector diffractometer (Mo K graphite-monochromated radiation)
using the APEX2 software[2], and equipped with an Oxford Cryosystems Series 700
cryo stream controlled by the Cryopad interface.[3] Images were processed with
SAINT+ software[4], and the absorption corrections were performed by the multi-scan
method implemented in SADABS.[5] The structure was solved by direct methods
implemented in SHELXS-97[6, 7], allowing the immediate identification of most of the
heaviest elements, namely Eu and W atoms, while the remaining atoms were
positioned through successive full-matrix least squares refinement cycles on F2 using
SHELXL-97.[6, 8] All atoms of the Europium-phosphotungstate anions and the K+ cations
were refined using anisotropic displacement parameters, while the oxygen atoms of
crystallization water molecules were refined with isotropic parameters. Although the H-
atoms of the water molecules were not located from difference Fourier maps or
positioned in calculated positions, they were added to the molecular formula of the
compound. X-ray crystallography analysis was carried out by Dr. Luis Cunha Silva.
5.1.2.9. Photoluminescence and lifetime measurements
Emission and excitation spectra were recorded at 298 K and 14 K using a
Fluorolog-2® Horiba Scientific (Model FL3-2T) spectroscope, with a modular double
grating excitation spectrometer (fitted with a 1200 grooves/mm grating blazed at 330
nm) and a TRIAX 320 single emission monochromator (fitted with a 1200 grooves/mm
grating blazed at 500 nm, reciprocal linear density of 2.6 nm∙mm-1), coupled to a R928
Hamamatsu photomultiplier, using the front face acquisition mode. The excitation
source was a 450 W Xe arc lamp. Emission spectra were corrected for detection and
optical spectral response of the spectrofluorimeter and the excitation spectra were
corrected for the spectral distribution of the lamp intensity using a photodiode reference
detector. Lifetime measurements were carried out using a 1934D3 phosphorimeter
coupled to the Fluorolog®-3, and a Xe-Hg flash lamp (6 μs/pulse half width and 20-30
μs tail) was used as the excitation source. The variable pressure measurements were
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Europium polyoxometalates encapsulated into silica nanoparticles
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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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Europium polyoxometalates encapsulated into silica nanoparticles
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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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.
5.2. Results and Discussion
In this work photoluminescent core/shell nanoparticles were prepared using
LnPOMs as the core, and using a reverse microemulsion technique for the alkaline
hydrolysis of TEOS around them. Silica encapsulation provides a protective layer
around the LnPOMs molecules, reducing the interaction with the surrounded media
which can adversely affect the properties of LnPOMs. Furthermore core-shell silica
nanoparticles are easily functionalized and chemically stable. The LnPOMs used were
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
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Europium polyoxometalates encapsulated into silica nanoparticles
linkers, namely (3-glycidyloxypropyl)- and (3-chloropropyl)-trimethoxysilanes, in order
to enable their subsequent binding to biologically active molecules. Photoluminescence
studies of the nanocomposites and the LnPOM salts were also performed to evaluate
the effect of the encapsulation on the luminescence properties of the LnPOMs.
5.2.1. Characterization of Eu(PW11)x compounds
The major structural difference between the two compounds synthesized is the
coordination sphere of europium metal centre. In Eu(PW11)2, the europium is
coordinated to eight oxygen atoms from the two PW11 units (four oxygens from the
lacunary region of each unit), and in the case of EuPW11 the europium is coordinated to
four lacunary oxygen atoms from PW11 and three water ligands, as previously
described in the literature (Figure 5.1).[14]
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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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.
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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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Europium polyoxometalates encapsulated into silica nanoparticles
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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
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Europium polyoxometalates encapsulated into silica nanoparticles
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
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Europium polyoxometalates encapsulated into silica nanoparticles
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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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Europium polyoxometalates encapsulated into silica nanoparticles
Compared to Eu(PW11)2@SiO2 nanoparticles prepared using 8 µmol of Eu(PW11)2
these results suggest that the molar quantity of europium-phosphotungstate does have
a relevant influence on the silica nanoparticle final size, with larger concentrations of
POMs leading to greater numbers of seeds, and thus smaller final particle size.
Figure 5.7 - TEM images of (a,b) Eu(PW11)2@SiO2 NPs prepared using 95 mg (16 µmol) of corresponding europium
polyoxometalate.; (c) Size distribution histogram of the mentioned EuPW11@SiO2 NPs. For direct comparison size
distribution the histogram of EuPW11@SiO2 (d) NPs prepared using 8 µmol of the same europium polyoxometalate is
also presented.
5.2.2.1. Scanning Electron Microscopy
Elemental mapping of NPs by scanning electron microscopy-energy dispersive X-
ray spectrometry (SEM-EDS) was performed to evaluate the distribution of the
encapsulated POMs into the NPs. Figure 5.8 and Figure 5.9 present the SEM-EDS
mapping images of EuPW11@SiO2 and Eu(PW11)2@SiO2 respectively.
The mapping identified the presence of tungsten (W) and silicon (Si) through the
NPs. Although it was not possible to identify the lanthanide ion Eu3+ by EDS (Figure 5.9
b), the results show that the W (one of the major elements that constitute POMs
structure) is well distributed in the samples and that it is surrounded by Si.
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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.
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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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Europium polyoxometalates encapsulated into silica nanoparticles
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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Europium polyoxometalates encapsulated into silica nanoparticles
151
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
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Europium polyoxometalates encapsulated into silica nanoparticles
with and without a high vacuum (ca. 5×10-6 mbar) are shown in Figure 5.14. Lifetime
measurements of the mentioned compounds and corresponding nanoparticles were
also performed and are presented in Figure 5.15.
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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Europium polyoxometalates encapsulated into silica nanoparticles
153
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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Europium polyoxometalates encapsulated into silica nanoparticles
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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Europium polyoxometalates encapsulated into silica nanoparticles
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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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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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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
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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
TEM images were obtained using a HITACHI H-8100 instrument operating at an
acceleration voltage of 200 kV. Samples for TEM analysis were prepared by depositing
suspensions of the nanoparticles in high glucose, phenol red-free DMEM on carbon
coated copper grids and allowing them to completely dry.
6.2.8. Statistical analysis
Data are presented as mean ± SEM (standard error of the mean). Statistical
analysis was performed using the GraphPad Prism 6.02 software (San Diego, USA).
Data were analysed by a non-parametric method the Kruskal–Wallis one-way analysis
of variance by ranks followed by Dunn’s test. Significance was accepted at a p value ≤
0.05.
6.3. Results and Discussion
In the studies of cell viability and cell morphology not only the effect of the
fluorescent silica nanoparticles but also the effect of bare silica nanoparticles (with
approximately 67 nm in diameter) and the fluorophores encapsulated in each type of
particle (RBITC and LnPOMs) were evaluated. In the particular case of silica
nanoparticles encapsulating LnPOMs, only the compound with higher stability was
used. As described previously in chapter 5, the compound that leads to the more stable
particles was the sandwich POM (Eu(PW11O39)2). The europium salt (europium
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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
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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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S iO 2 N P s
A
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R B IT C
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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
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A
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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).
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P O M s
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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 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).
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 assay, at 24 h (A) and 48 h (B) after exposure. Results were calculated
as percentage of control (vehicle-treated cells) and data are presented as mean ± SEM (n=8-28 per group).
Similar to the results obtained for RBITC@SiO2, RBITC and SiO2 NPs (presented in
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.
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1 0 0
1 2 5
1 5 0
1 7 5
2 0 0
C o n c e n tra tio n
( g /m L )
Ca
lce
in f
luo
res
ce
nc
e i
nte
ns
ity
(% o
f c
on
tro
l)
B
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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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
nanoparticles, RBITC dye and RBITC-APTES@SiO2 nanoparticles (100x magnification).
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Figure 6.11 - Representative phase contrast microscopy images of Hepa RG cells at 24 hours after incubation with SiO2
nanoparticles, RBITC dye and RBITC-APTES@SiO2 nanoparticles (100x magnification).
Phase contrast microscopy images presented in figures 6.9 to 6.11 shows normal
cell morphology after incubation with RBITC-APTES@SiO2, RBITC and SiO2 NPs. This
indicates that no morphological changes had occurred in the three cell lines after
incubation with the RBITC dye and the NPs. Results on cell morphology are in
agreement with those obtained on cell viability measurements since no cytotoxic
effects were observed by either technique.
6.3.3.2. Eu(PW11O39)2@ SiO2 NPs
Figures 6.12, 6.13 and 6.14 show the images of phase contrast microscopy of Eu+3,
Eu(PW11O39)2, Eu(PW11O39)2@SiO2 and SiO2 NPs incubated with Caco-2, SH-SY5Y
and HEPA RG cells respectively. The images were collected after 24 h incubation at a
100x magnification.
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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
nanoparticles, Eu salt, Eu(PW11O39)2 and Eu(PW11O39)2@SiO2 nanoparticles (100x magnification).
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Figure 6.14 - Representative phase contrast microscopy images of Hepa RGcells at 24 hours after incubation with SiO2
nanoparticles, Eu salt, Eu(PW11O39)2 and Eu(PW11O39)2@SiO2 nanoparticles (100x magnification).
Regarding the potential changes in cell morphology by phase contrast
microscopy it can be observed from figures 6.12 to 6.14 that for the three cellular lines
used there were no morphological changes after cell treatment with Eu3+,
Eu(PW11O39)2, Eu(PW11O39)2@SiO2 or SiO2 NPs. Similar to the results presented in the
previous section (6.3.3.1), phase contrast microscopy images show normal cell
morphology after cell incubation with the compounds of interest after 24 h. Again
results on cell morphology are in agreement with those obtained on cell viability
measurements.
6.4. Conclusions
Cellular uptake, cellular viability and cellular morphology experiments were
performed to evaluate possible cytotoxicity of the fluorescent silica NPs synthesized in
the present work. For this purpose Caco-2, SH-SY5Y and Hepa RG cellular lines were
incubated with increasing concentrations from 0.1 µg/mL up to 3.2 µg/mL of the
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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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6.5. References
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Viscardi, G. and Martra, G., Highly bright and photostable cyanine dye-doped
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11. Barandeh, F., Nguyen, P.L., Kumar, R., Iacobucci, G.J., Kuznicki, M.L.,
Kosterman, A., Bergey, E.J., Prasad, P.N. and Gunawardena, S., Organically
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Modified Silica Nanoparticles Are Biocompatible and Can Be Targeted to
Neurons In Vivo. Plos One, 2012. 7(1).
12. Lin, W.S., Huang, Y.W., Zhou, X.D. and Ma, Y.F., In vitro toxicity of silica
nanoparticles in human lung cancer cells. Toxicology and Applied
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13. Ahmad, J., Ahamed, M., Akhtar, M.J., Alrokayan, S.A., Siddiqui, M.A., Musarrat,
J. and Al-Khedhairy, A.A., Apoptosis induction by silica nanoparticles mediated
through reactive oxygen species in human liver cell line HepG2. Toxicology and
Applied Pharmacology, 2012. 259(2): p. 160-168.
14. Liu, T., Li, L., Fu, C., Liu, H., Chen, D. and Tang, F., Pathological mechanisms
of liver injury caused by continuous intraperitoneal injection of silica
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15. Nishimori, H., Kondoh, M., Isoda, K., Tsunoda, S., Tsutsumi, Y. and Yagi, K.,
Silica nanoparticles as hepatotoxicants. European Journal of Pharmaceutics
and Biopharmaceutics, 2009. 72(3): p. 496-501.
16. Park, E.J. and Park, K., Oxidative stress and pro-inflammatory responses
induced by silica nanoparticles in vivo and in vitro. Toxicology Letters, 2009.
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17. Riss, T.L. and Moravec, R.A., Use of multiple assay endpoints to investigate the
effects of incubation time, dose of toxin, and plating density in cell-based
cytotoxicity assays. Assay and Drug Development Technologies, 2004. 2(1): p.
51-62.
18. Niles, A.L., Moravec, R.A., Hesselberth, P.E., Scurria, M.A., Daily, W.J. and
Riss, T.L., A homogeneous assay to measure live and dead cells in the same
sample by detecting different protease markers. Analytical Biochemistry, 2007.
366(2): p. 197-206.
19. Mosmann, T., Rapid Colorimetric Assay for Cellular Growth and Survival -
Application to Proliferation and Cyto-Toxicity Assays. Journal of Immunological
Methods, 1983. 65(1-2): p. 55-63.
20. Foldbjerg, R., Wang, J., Beer, C., Thorsen, K., Sutherland, D.S. and Autrup, H.,
Biological effects induced by BSA-stabilized silica nanoparticles in mammalian
cell lines. Chemico-Biological Interactions, 2013. 204(1): p. 28-38.
21. Malvindi, M.A., Brunetti, V., Vecchio, G., Galeone, A., Cingolani, R. and Pompa,
P.P., SiO2 nanoparticles biocompatibility and their potential for gene delivery
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22. Kim, Y.J., Yu, M., Park, H.O. and Yang, S.I., Comparative study of cytotoxicity,
oxidative stress and genotoxicity induced by silica nanomaterials in human
neuronal cell line. Molecular & Cellular Toxicology, 2010. 6(4): p. 337-344.
23. Kim, Y.J. and Yang, S.I., Neurotoxic effects by silica TM nanoparticle is
independent of differentiation of SH-SY5Y cells. Molecular & Cellular
Toxicology, 2011. 7(4): p. 381-388.
24. Hungerford, G., Hussain, F., Patzke, G.R. and Green, M., The photophysics of
europium and terbium polyoxometalates and their interaction with serum
albumin: a time-resolved luminescence study. Physical Chemystry Chemical
Physics, 2010. 12(26): p. 7266-75.
25. Geisberger, G., Gyenge, E.B., Hinger, D., Bosiger, P., Maake, C. and Patzke,
G.R., Synthesis, characterization and bioimaging of fluorescent labeled
polyoxometalates. Dalton Transactions, 2013. 42(27): p. 9914-9920.
26. Geisberger, G., Paulus, S., Carraro, M., Bonchio, M. and Patzke, G.R.,
Synthesis, Characterisation and Cytotoxicity of Polyoxometalate/Carboxymethyl
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Concluding Remarks and Perspectives
Fluorescent silica nanoparticles show unique chemical and optical properties such
as bright fluorescence, higher photostability and biocompatibility compared to classical
fluorophores. Moreover, fluorescent silica nanoparticles are quite easy to prepare,
exhibit good monodispersity and their surface can be easily functionalized for further
bioconjugation. Due to these attractive features, fluorescent silica nanoparticles have
received an increasing interest for biological applications in the past few years.
However, despite the increasing applicability in biological fields the biocompatibility of
these nanoparticles is not completely clear and remains an issue of debate. Therefore,
nanotoxicology research has been intensified in order to obtained detailed information
about biocompatibility of silica based nanomaterials
The purpose of this research work was to prepare and optimize the synthesis of two
different kinds of fluorescent silica nanoparticles incorporating organic (rhodamine b
isothiocyanate – RBITC) and inorganic (lanthanide-based polyoxometalates –
LnPOMs) fluorophores. It was also an aim of this study the characterization of these
nanomaterials in order to evaluate the effects of silica encapsulation on the stability,
fluorescence emission and quantum yield of the fluorophores. In addition, the
evaluation of the potential cytotoxicity of these nanoparticles was also a study purpose.
In this work it was clearly demonstrated that using a reverse microemulsion
methodology for the alkaline hydrolysis of TEOS it was possible to produce fluorescent
silica nanoparticles incorporating the different fluorophores. All the samples of the
fluorescent nanoparticles synthesized show a narrow polydispersity. Furthermore, in
the synthesis of fluorescent silica NPs encapsulating the inorganic fluorophores
(LnPOMs) the ratio of LnPOM/TEOS used has a strong influence on the nanoparticle
final size indicating that LnPOMs act as seeds for the growth of the silica shell. In this
work, it was found that the reverse-microemulsion-based methodology gives superior
results compared to the Stöber method regarding size and shape control and
reproducibility and also in terms of fluorescent yield and stability.
After encapsulation of the different fluorophores within the silica NPs the structural
integrity, stability and quantum yield were evaluated. RBITC silica NPs show similar
spectral properties (absorption, fluorescence excitation and emission) to the free dye,
indicating the successful doping of dye molecules into silica matrix. One of the
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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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189
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.