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

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 Development of fluorescent silica nanoparticles encapsulating organic and inorganic fluorophores: synthesis and characterization Cristina Sofia dos Santos Neves PhD FCUP FCT/UNL 2014 3.º CICLO D D 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 “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) FCUP 1 Acknowledgments “No man is an island, Entire of itself, Every man is a piece of the continent, A part of the main.” (John Donne) During the development of this work many challenges have emerged, and with them several people who helped me in one way or another, to overcome them. This thesis wouldn’t be the same without their support and for that I would like to take a few lines to thank those who believed in me and in this work. First of all I would like to thank my supervisors Dr. Peter Eaton and Dr. Eulália Pereira for the opportunity to perform this work. Thank you for accepting me in the lab and for providing all means for the development of this project. Also for the guidance and words of encouragement that were needed in some curves of this way. Thank you also for the opportunities you gave me. Part of the work present in this thesis wouldn’t be possible without the help of Dr. Salete Balula. Her guidance, support and friendship were of great importance to achieve my goals. Thanks for introducing me to the amazing world of polyoxometalates and giving me the chance to learn and grow up in this field. To Dr. Carlos Granadeiro and Dr. Luis Cunha Silva a special thanks for the help with the synthesis and characterization of some nanoparticles in particular the help with X-ray crystallography and FT-RAMAN spectroscopy. To Dr. Sandra Gago from the Chemistry Department and to Dr. Gabriel Feio from the Department of Materials Science (CENIMAT), Faculty of Sciences and Technology - University Nova de Lisboa, for the solid-state nuclear magnetic resonance experiments and all the patience and help with interpretation of those results. To Dr. Duarte Ananias from Centre for Research in Ceramics and Composite Materials (CICECO) associated laboratory, University of Aveiro for the photoluminescence studies and his help with results interpretation. I also thank CICECO associated laboratory where part of the characterization techniques were carried out. 2 FCUP Acknowledgments To Dr. Patricia Carvalho for all availability in the matters concerned with transmission electron microscopy and for all the suggestions and contributions related to this work. To Dr. César Laia and Dr. João Lima from the group of photochemistry of the Chemistry Department, Faculty of Sciences and Technology - University Nova de Lisboa, for full availability on anisotropy and fluorescence lifetime measurements and for all the patience, help and support. To Dr. Sónia Fraga from the laboratory of toxicology of the Faculty of Pharmacy from University of Porto I would like to thank all the help with the cytotoxic measurements. I also thank all the enthusiasm and the patience during this final stage of my work. To Fundação para a Ciência e a Tecnologia (FCT) I thank the financial support through the PhD grant SFRH/BD/61137/2009. Along these last five years in Porto I had the chance to work and live with several persons. Each one of them on their particular way made me feel at home. To my lab colleagues Pedro Quaresma and Leonor Soares I have to thank the way they integrated me in their world, how they helped me during my first steps in nanochemistry and nanotechnology fields. To them and also to others that were coming and going I thank the excellent environment, the fellowship and friendship, and those times that we laugh to tears. Also to Pedro Quaresma I thank the numerous times we discuss strategies and plans even when they were just guesses. To Ana Claudia for her optimism, and trying to make me see the bright side of everything, I know it was a great effort. I would also like to thank Catarina Loureiro, for her friendship and care. I create many bonds over the years and I couldn’t go without mentioning some of them that were very important to me. First I’d like to thank Sónia Patricio, she was the first person I met when I arrived, she was the face that welcomed me and it was a pleasure to meet her and becoming her friend. To Carla Queirós for her friendship and caring even in my worst moments and a big thank you for helping me whenever I needed. For all the support, help, caring and words of encouragement during these rough times a sincere thanks to Daniela Leite, André Barbosa, Susana Ribeiro and Ana Margarida Silva. FCUP 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 4 FCUP Acknowledgments person along this journey, but nonetheless you’ll never gave up on me. You have been a supporting rock in all the bad moments and in the good ones you have been the most cheerful person. Thank you for believing in me when I doubted, for being a friend, a listener, a supporter and most important my companion for life. To all of those who contributed somehow to this thesis and prevented me to go insane and aren't mentioned here, I appreciate your effort and caring. I leave you a sincere thanks and the following quote: “Those who pass us by do not go alone, do not leave us alone. Leave a bit of themself, take a little of us.” (Antoine de Saint-Exupéry) FCUP 5 AUTHOR PUBLICATIONS CONTAINING WORK RELATED WITH THIS THESIS: Articles and book chapters in peer-reviewed journals Published/submitted papers: Cristina S. Neves, Carlos M. Granadeiro, Luis Cunha-Silva, Duarte Ananias, Sandra Gago, Gabriel Feio, Patricia A. Carvalho, Peter Eaton, Salete S. Balula, Eulália Pereira, Europium-polyoxometalates encapsulated into sílica nanoparticles: characterization and photoluminescence studies, European Journal of Inorganic Chemistry 16, 2877-2886 (2013). Book chapter: Pedro V. Baptista, Gonçalo Doria, Pedro Quaresma, Miguel Cavadas, Cristina S. Neves, Inês Gomes, Peter Eaton, Eulália Pereira, Ricardo Franco, Nanoparticles in Molecular Diagnosis, Progress in Molecular Biology and Translational Science, Vol. 104, 427-488, (Antonio Villaverde, Ed.) Elsevier (2011). Awards: Young Scientist Award on European Materials Research Society (E-MRS) 2013 Spring Meeting, Strasbourg, France, 26th – 31st May 2013. Best Paper Award on XI International Conference on Nanostructured Materials, Rhodes, Greece, 23rd – 31st August 2012. Articles not included in this dissertation, but published in the course of the Ph.D. work: Silvia C. Lopes, Cristina Neves, Peter Eaton, Paula Gameiro, Improved model systems for bacterial membranes from differing species: the importance of varying composition in PE/PG/cardiolipin ternary mixtures, Molecular Membrane Biology, 29 (2012), 207-217. Maria J. Medeiros, Cristina S. S. Neves, A. R. Pereira, Elizabeth Duñach, Electroreductive intramolecular cyclisation of bromoalkoxylated derivatives catalized by nickel (I) tetrametylcyclam in “green” media, Electrochimica Acta, 56 (2011), 4498- 4503. 6 FCUP Author publications Silvia Lopes, Cristina S. Neves, Peter Eaton, Paula Gameiro, Cardiolipin, a key component to mimic the E. Coli bacterial membrane in model systems revealed by dynamic light scattering and steady-state fluorescence anisotropy, Analytical & Bioanalytical Chemistry, 398 (2010) 1357-1366. Cristina S. Neves, Pedro Quaresma, Pedro V. Baptista, Patricia A. Carvalho, João P. Araújo, Eulália Pereira, Peter Eaton, New insights into the use of magnetic force microscopy to discriminate between magnetic and nonmagnetic nanoparticles, Nanotechnology, 21 (2010) 30576. FCUP General Index 13 4.1.1 Chemicals ................................................................................................. 99 4.1.2 Instrumentation and methodologies ........................................................ 100 4.1.1.1. Elemental Analysis .................................................................................... 100 4.1.1.2. UV-visible spectroscopy ............................................................................ 100 4.1.1.3. Fluorescence spectroscopy, quantum yield and lifetime ........................... 100 4.1.1.4. Steady-state anisotropy ............................................................................ 101 4.1.1.5. Transmission electron microscopy ............................................................ 102 4.1.1.6. Dynamic light scattering and zeta potential ............................................... 102 4.1.3 Preparation of core-shell nanoparticles with rhodamine B isothiocyanate (RBITC-APTES@SiO2) ...................................................................................... 102 4.1.4 Surface functionalization of silica nanoparticles ...................................... 103 4.1.5 DNA grafting ........................................................................................... 103 4.2. Results and Discussion ................................................................................. 104 4.2.1 Characterization of RBITC@SiO2 nanoparticles ..................................... 104 4.2.1.1. Characterization by electron microscopy ................................................... 104 4.2.1.1. Dynamic light scattering ............................................................................ 106 4.2.1.2. Characterization by UV-vis spectroscopy .................................................. 107 4.2.1.3. Fluorescence excitation and fluorescence emission spectra ..................... 110 4.2.1.4. Fluorescence quantum yield ..................................................................... 112 4.2.1.5. Lifetime measurements ............................................................................. 115 4.2.1.6. Fluorescence anisotropy ........................................................................... 119 4.2.2 Characterization of RBITC-APTES@SiO2 NPs grafted with DNA ........... 122 4.2.3 Conclusions ............................................................................................ 124 4.3. References .................................................................................................... 127 5. Europium polyoxometalates encapsulated into silica nanoparticles ...................... 131 5.1. Materials and Methods .................................................................................. 131 5.1.1. Chemicals ............................................................................................... 131 5.1.2. Instrumentation and methodologies ........................................................ 132 14 FCUP General Index 5.1.2.1. Elemental analysis .................................................................................... 132 5.1.2.2. Vibrational Spectroscopy .......................................................................... 132 5.1.2.3. Solid state NMR ........................................................................................ 132 5.1.2.4. Transmission electron microscopy ............................................................ 133 5.1.2.5. Scanning electron microscopy .................................................................. 133 5.1.2.6. Dynamic light scattering ............................................................................ 133 5.1.2.7. Atomic force microscopy ........................................................................... 133 5.1.2.8. X-ray crystallography ................................................................................ 134 5.1.2.9. Photoluminescence and lifetime measurements ....................................... 134 5.1.2.10. Quantum efficiency ................................................................................... 135 5.1.3. Synthesis of europium polyoxometalates Eu(PW11)x (x = 1 and 2) .......... 136 5.1.4. Encapsulation of Eu(PW11)x (x = 1 and 2) into silica nanoparticles .......... 136 5.1.5. Functionalization of Eu(PW11)2@SiO2 ..................................................... 137 5.2. Results and Discussion ................................................................................. 137 5.2.1. Characterization of Eu(PW11)x compounds ............................................. 138 5.2.1.1. X-ray crystallography ................................................................................ 138 5.2.1.2. Thermogravimetry ..................................................................................... 140 5.2.1.3. 31P NMR spectroscopy .............................................................................. 141 5.2.2. Characterization of Eu(PW11)x@SiO2 nanoparticles ................................ 142 5.2.2.1. Transmission Electron Microscopy ............................................................ 143 5.2.2.1. Scanning Electron Microscopy .................................................................. 146 5.2.2.2. Characterization by vibrational spectroscopy ............................................ 148 5.2.2.3. Photoluminescence properties .................................................................. 151 5.2.3. Conclusions ............................................................................................ 155 5.3. References .................................................................................................... 156 6. Cytotoxicity evaluation of RBITC and LnPOMs fluorescent silica nanoparticles .... 161 6.1. Cytotoxicity assays ........................................................................................ 162 6.2. Materials and Methods .................................................................................. 163 FCUP General Index 15 6.2.1. Chemicals ............................................................................................... 163 6.2.2. Cellular culture ........................................................................................ 164 6.2.3. Nanoparticle uptake ................................................................................ 164 6.2.4. Cell viability by Calcein-AM assay .......................................................... 164 6.2.5. Phase contrast microscopy ..................................................................... 165 6.2.6. Fluorescence spectroscopy .................................................................... 165 6.2.7. Transmission electron microscopy .......................................................... 165 6.2.8. Statistical analysis .................................................................................. 165 6.3. Results and Discussion ................................................................................. 165 6.3.1. Cellular uptake of silica nanoparticles ..................................................... 166 6.3.2. Cell esterase activity (Calcein-AM assay) ............................................... 168 6.3.2.1. Effect of RBITC@SiO2 NPs on Caco-2, SH-SY5Y and Hepa RG cells viability ................................................................................................................. 169 6.3.2.2. Effect of Eu(PW11O39)2@SiO2 NPs on Caco-2, SH-SY5Y and Hepa RG cells viability ................................................................................................................. 172 6.3.3. Morphological analysis by phase contrast microscopy ............................ 174 6.3.3.1. RBITC-APTES FSNPS ............................................................................. 174 6.3.3.2. Eu(PW11O39)2@ SiO2 NPs ........................................................................ 177 6.4. Conclusions ................................................................................................... 179 6.5. References .................................................................................................... 181 III – Concluding Remarks and Perspectives Concluding Remarks and Perspectives..................................................................... 187 FCUP 16 FCUP 17 List of Tables I - Introduction Table 2.1 - Chemical binding for bioconjugation of silica NPs. (adapted from Yao et al. [2]) ................................................................................................................................ 53 Table 3.1 - Commonly observed emission bands of the lanthanide ions Eu3+, Tb3+, Nd3+, Er3+ and Yb3+ in solution. (Adapted from Werts[50]) ............................................. 74 II – Research work Table EB 1– Comparison between the three methods followed to prepare fluorescent silica NPs using the microemulsion technique............................................................. 95 Table 4.1 - Average hydrodynamic diameter of RBITC FSNPs measured by DLS (by percentage of number of particles, measurements were repeated 5 times for each sample). .................................................................................................................... 106 Table 4.2 – Amount of RBITC dye molecules per fluorescent silica nanoparticle ...... 110 Table 4.3 - Fluorescence quantum yields of RBITC, RBITC-APTES conjugate and FSNPs ...................................................................................................................... 113 Table 4.4 - Lifetime data of RBITC and fluorescent silica nanoparticles (FSNPs) in absolute ethanol ....................................................................................................... 115 Table 4.5 - Anisotropy (r) and rotational diffusion coefficient (Dr) values of RBITC and fluorescent silica nanoparticles (FSNPs) adsorbed and covalently bound to silica NPs in absolute ethanol. ................................................................................................... 120 Table 4.6 – Zeta potential ζ of FSNPs-GPTES and FSNPs-GPTES-DNA ................. 123 Table 5.1 - Crystal and structure refinement data for Eu(PW11)2 ............................... 140 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 18 FCUP FCUP 19 List of Figures I - Introduction Figure 1.1 - Several common fluorescent nanoscale materials including (a) organic dye molecules (tetramethylrhodamine); (b) green fluorescent protein; (c) polymer-coated, water soluble semiconductor quantum dots and (d) fluorophore-doped silica particles. (Adapted from Burns et al.[3]) ...................................................................................... 33 Figure 1.2 - Plain and ball-and-stick structures of fluorescein isothiocyanate (FTIC). .. 34 Figure 1.3 – Plain and ball-and-stick structures of rhodamine 6G (top) and rhodamine b (bottom). ..................................................................................................................... 35 Figure 1.4 - Basic structure of cyanine dyes............................................................... 36 Figure 1.5 – Plain and ball-and-stick structures of Alexa Fluor 350. ............................ 36 Figure 1.6 – Plain and ball-and-stick structures of Alexa Fluor 430. ............................ 37 Figure 1.7 - Structure of the Aequorea victoria green fluorescent protein. (Source: Ormö et al.[15]) ............................................................................................................. 38 Figure 1.8 - Photograph and spectra of CdSe quantum dots. The samples represent different sizes of QDs, which produce different colours upon UV light. An increase in particle size produces a red shift in the emission spectra. (Source: Nauman et al. [17]) ................................................................................................................................... 39 Figure 1.9 - Schematic illustration of the surface functionalization of silica NPs with, for example, peptides, antibodies, aptamers, enzymes, DNA-fragments and different functional moieties. (Source: Schulz et al.[21]) .............................................................. 41 Figure 2.1 - TEM images: (A) silica-based nanoparticles prepared by the Stöber method; and (B) silica nanoparticles prepared by the microemulsion process............. 47 Figure 2.2 - Typical structure of a reverse micelle (source: Malik et al.[11]) .................. 48 Figure 2.3 – Silica nanoparticles growth mechanism in a reverse micellar system composed. (Source: Osseo-Asare et al.[21]) ................................................................. 51 Figure 2.4 - Schematic illustration of the surface functionalization of silica NPs for biological applications. (Source: Smith et al.[4]) ........................................................... 52 Figure 2.5 - Representative bioconjugation schemes for attaching biomolecules to dyedoped silica NPs for bioanalysis. (source: Wang et al. [3]) ........................................... 54 20 FCUP List of Figures Figure 2.6 - Confocal fluorescence microscopy images (overlaid and bright field)of pH sensors in rat basophilic leukemia mast cells showing a) reference dye (RBITC) channel, b) sensor dye (FTIC) channel, c) overlaid images and d) false-colour ratiometric imaging of pH in various intracellular compartments (Source: Burns et al.[30]) ................................................................................................................................... 57 Figure 2.7 - Schematic representation of a sandwich assay based on dye-doped silica NPs. (Source: Zhao et al.[33]) ....................................................................................... 59 Figure 3.1 - Ball-and-stick (left) and polyhedral (right) representations of the 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 FCUP List of Figures 21 II – Research work Figure EB 1 - UV-vis spectrum of fluorescent silica nanoparticles synthesized by Stober’s method through the adapted procedure described by Bringley[1]. .................. 91 Figure EB 2- TEM images of TRICT fluorescent silica nanoparticles prepared by Stöber’s method following a similar procedure to that described by Larson[3] et al. ..... 93 Figure EB 3 - TEM images of fluorescent silica nanoparticles prepared by the reverse microemulsion system following the procedures of Gao[4] (A); Zhang[5] (B) and Shi[6] (C). ................................................................................................................................... 96 Figure EB 4 - Fluorescence emission spectra of fluorescent silica nanoparticles prepared by the reverse microemulsion system following the procedures of Gao[4] (A); Zhang[5] (B) and Shi[6] (C). ........................................................................................... 96 Figure 4.1 - TEM images of RBITC-APTES FSNPS nanoparticles and corresponding size distribution histogram......................................................................................... 105 Figure 4.2 - SEM images of RBITC-APTES FSNPs showing the spherical morphology of the NPs. ................................................................................................................ 105 Figure 4.3 - DLS hydrodynamic diameter distribution statistics graph (by percentage of number of particles) for RBITC FSNPS. Error bars show standard deviation of five different measurements. ........................................................................................... 107 Figure 4.4 - UV-vis spectra of RBITC and RBITC fluorescent silica NPs (FSNPs) in ethanol at 25 ºC. UV-vis spectrum of FSNPs was fitted using a 2nd order exponential decay to remove silica scattering. Both samples were dissolved to a final concentration with almost the same absorbance (0.09). .................................................................. 108 Figure 4.5 - UV-vis spectra of RBITC and RBITC-APTES conjugate in ethanol at 25 ºC. ................................................................................................................................. 109 Figure 4.6 - Fluorescence excitation spectra of RBITC, RBITC-APTES conjugate and FSNPs recorded at 25 ºC in absolute ethanol. .......................................................... 111 Figure 4.7 - Fluorescence emission spectra of RBITC, RBITC-APTES conjugate and FSNPs recorded at 25 ºC in absolute ethanol. .......................................................... 112 Figure 4.8 - (A) RBITC doped fluorescent silica NPs prepared by hydrolysis and polymerization of TEOS in a microemulsion method; (B) bare silica NPs with RBITC dye molecules adsorbed onto the nanoparticle’s surface; (C) fluorescent core-shell NPs with a silicon core and a shell of RBITC dye molecules and TEOS. .......................... 113 22 FCUP List of Figures Figure 4.9 - Fluorescence lifetime decay curves of RBITC (A), RBITC-APTES conjugate (B), RBITC-APTES FSNPs with dye covalently bound to silica matrix (RBITC-APTES@SiO2) (C),and RBITC-APTES FSNPs with dye adsorbed to silica surface (Ads:RBITC-APTES@SiO2) (D), all at ambient temperature (298 K) .The excitation was fixed at 370 nm and the emission was monitored at 550 nm. ............. 116 Figure 4.10 – Structures of rhodamine b isothiocyanate (RBITC) isomers. Left: rhodamine b 5-isothiocyanate and right: rhodamine b 6-isothiocyanate. ................... 117 Figure 4.11 – Structures of RBITC-APTES conjugate for RBITC 5-isomer (top) and RBITC 6-isomer (bottom) .......................................................................................... 118 Figure 4.12 - Steady state emission fluorescence anisotropy of RBITC and RBITC FSNPs with dye adsorbed (Ads:RBITC-APTES@SiO2) and dye covalently bound (RBITC-APTES@SiO2) to silica matrix. The excitation wavelength was 530 nm. ...... 120 Figure 4.13 - Representation of the wobbling-in-cone model, where θc is the angle between the probe (dye) axis (direction of the optical transition moment) and the symmetry axis of the wobbling motion (cone axis). ................................................... 121 Figure 4.14 - Strategy for immobilisation of thiolated oligonucleotides onto dye loaded silica nanoparticle surfaces. ...................................................................................... 122 Figure 4.15 - UV-vis spectra of DNA and functionalized FSNPS before (FSNPs- GPTES) and after (FSNPs-GPTES-DNA) DNA immobilization in potassium phosphate buffer (10 mM, pH = 8). Inset: zoom in the UV-vis spectra of FSNPS before and after DNA immobilization................................................................................................... 123 Figure 5.1 - (a) The structures of the sandwich type europium-phosphotungstate anion, [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 FCUP 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 30 FCUP Abbreviations and Symbols TRITC - tetramethylrhodamine isothiocyanate UV-vis – ultraviolet-visible spectroscopy W/O - water-in-oil microemulsion W0 - water-to-surfactant molar ratio ε – absorption coefficient ζ - zeta potential I Introduction FCUP 33 1. An overview of nanoscale fluorescent materials Nanoscale materials are a broadly defined set of substances that have at least one critical dimension less than 100 nm and possess unique optical, magnetic, electrical or other properties.[1] Thus, when particle size is be nanoscale, properties such as melting point, fluorescence, electrical conductivity, magnetic permeability, and chemical reactivity can change as a function of the size of the particle. Nanoscale fluorescent materials gained particular interest in the fields of chemistry, biology, medical science and biotechnology.[2] Because nanoscale fluorescent particles may not be visible to the naked eye, it is possible to use them as hidden fluorescent substances which only become fluorescent once they have been excited by light of a specific wavelength. This property makes them of particular interest for biological applications where in past decades many progresses have been made. Due to their high signal-to-noise ratio, excellent spatial resolution, and ease of implementation, fluorescent materials are ideal to investigate biology down to nanoscale.[3, 4] Every application has its own particular restrictions, but the most important properties for any fluorescent material are the same: brightness and stability. There are several classes of materials currently employed as fluorescent emitters/probes, which includes organic dye molecules, fluorescent proteins, semiconductor quantum dots, polymer/dye-based nanoparticles and silica/fluorophores hybrid particles (Figure 1.1), and each of them have their own advantages and disadvantages. Figure 1.1 - Several common fluorescent nanoscale materials including (a) organic dye molecules (tetramethylrhodamine); (b) green fluorescent protein; (c) polymer-coated, water soluble semiconductor quantum dots and (d) fluorophore-doped silica particles. (Adapted from Burns et al.[3]) 34 FCUP An overview of nanoscale fluorescent materials 1.1. Organic dye molecules Organic dye molecules are the smallest fluorescent emitters used today. These fluorophores are commercially available with emissions from UV to the near infrared region of the spectrum (~300-900nm). These dye molecules have a small size (~1nm) which makes them an excellent choice for many applications especially in biology where they are the most commonly used fluorophores. Although they present some limitations like short stokes shifts (difference between maximum wavelengths of absorption and emission bands), rapid photobleaching, poor photochemical stability and decomposition under repeated excitation, the organic dyes are still widely used due to their low cost, availability and ease of usage. Examples of commonly used organic dyes include fluorescein, rhodamine, cyanine and Alexa dyes. 1.1.1. Fluoresceins and rhodamines Fluoresceins are amine-reactive organic fluorophores widely used in biolabelling.[5, 6] They belong to the xanthene class of dyes[7] with absorption and fluorescence maxima in the visible region (e.g. fluorescein λabs = 490 nm and λem = 512 nm in water). Fluoresceins have high extinction coefficient and quantum yield and also high solubility in water. However they present some major drawbacks such as photobleaching, pH sensitivity, relatively broad emission spectra and tendency for self-quenching after bioconjugation.[5, 8, 9] As a result the use of fluorescein dyes in ultra-sensitive biological studies is limited. Figure 1.2 - Plain and ball-and-stick structures of fluorescein isothiocyanate (FTIC). FCUP 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). 36 FCUP 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. FCUP 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] 38 FCUP An overview of nanoscale fluorescent materials Figure 1.7 - Structure of the Aequorea victoria green fluorescent protein. (Source: Ormö et al.[15]) 1.3. Semiconductor quantum dots Semiconductor nanocrystals, also known as Quantum Dots (QDs), are a novel inorganic fluorophores class which have become popular in the past two decades due to their exceptional photophysical properties. These semiconductors nanocrystals have sizes in the range of 1-10 nm and their fluorescence is due to quantum confinement effects. Also due to quantum confinement the absorption and emission wavelengths of QDs is size dependent, meaning that they can be synthesized in a wide range of sizes with the wavelength of light emitted related to the size of the QDs (Figure 1.8). QDs are composed of combined elements from periodic groups II–VI (e.g. ZnS, CdSe, CdTe), III–V (e.g. InP, GaAs, InAs), or IV–VI (e.g. PbS, PbSe, PbTe) and they provide a new class of biomarkers that can overcome the usual limitations of organic dyes.[16] QDs exhibit high photostability, broad absorption, narrow and symmetric emission spectra, slow excited decay rates and large absorption cross-sections. Their broad absorption and narrow emission spectra allows a single laser to excite QDs of a wide size-range, with each dot emitting its own specific colour, contrasting to organic-based fluorophores, which are characterized by narrow Stokes shifts.[17] Also when compared with traditional organic fluorophores they present unique fluorescence properties.[18] Unlike organic fluorophores which bleach after only a few minutes on exposure to light, QDs are extremely stable and can undergo repeated cycles of excitation and fluorescence for hours with a level of brightness and photobleaching threshold.[12, 19] QDs are used in biological applications such as cellular labelling, cell tracking and tissue imaging.[20] However, QDs themselves are hydrophobic and non-biocompatible, requiring layers of polymeric or inorganic material to make them compatible for biological applications.[3] Additionally some QDs contain toxic components, such as FCUP 45 2. Fluorescent silica nanoparticles doped with organic dyes Fluorescent dye-doped silica NPs have been extensively used for a wide range of applications in biological detection and diagnosis in the past several years. Dye dopedsilica NPs are extremely bright and photostable because a large number of fluorescent dye molecules are encapsulated in the silica matrix that serves to protect the dye from photodegradation. This also enables NPs to exhibit strong emission signal when properly excited which can dramatically lower the analyte detection limit in biological samples. Using appropriate synthetic conditions, a large variety of either organic or inorganic dye molecules can be incorporated inside a single silica particle. Despite the fact that incorporating a large amount of dye molecules into a single NP would be expected to lead to some fluorescence quenching phenomena due to the particle small volume, the goal of obtaining a particle with brighter luminescence is largely successful.[1] Photostability is an important criterion in observation of fluorescence signal, especially under intense excitation and one of the major concerns when using bare dyes in bioanalysis. However, when dye molecules are encapsulated into a silica matrix which provides an effective barrier from the surrounding environment, the doped dye molecules are protected from oxygen and both photobleaching and photodegradation phenomena that often affect conventional dyes can be minimized. Moreover, the encapsulation enables the fluorescence to be constant providing an accurate measurement making these NPs suitable for applications where high intensity or intense excitations are needed.[1, 2] In addition, when dye doped silica NPs are used in real biological media the dye molecules are protect against degradation by the complex biological environment due to high resistance to chemical and metabolic degradation of the silica matrix.[3] Silica is a good matrix material due to its flexible chemistry which allows surface modification with different types of functional groups. This property makes silica a versatile and biocompatible substrate for biomolecule immobilization. Biochemically modified dye-doped silica NPs can be used to express the activity of a desired process, such as enzyme immobilization, or can be used as an affinity ligand to capture or 46 FCUP Fluorescent silica nanoparticles doped with organic dyes modify target molecules or cells.[3] In addition the silica surface makes these NPs chemically inert and physically stable.[4] Furthermore, studies on the cytotoxicity of these NPs have showed the benign nature of silica based NPs, exhibiting low or no cytotoxicity.[3] Owing to these properties the use of dye-doped silica NPs as labelling reagents for bioanalysis and bioimaging have been widely studied and used.[1, 3] For example these NPs have been used to create assays for oligonucleotides, proteins and antibodies[5], cell targeting[3] and intracellular sensors[6]. 2.1. Nanoparticle formation Commonly, silica NPs are formed as a result of the base-catalysed hydrolysis of an organic precursor like tetraethylortosilicate (TEOS) and subsequent condensation to a silica network as shown in the equations (1) and (2).[6] (1) (2) Silica NPs are generally synthesized by one of two chemical routes. These are solgel synthesis also known as the Stöber method[7] and the reverse microemulsion approach[8]. In the Stöber method, TEOS is hydrolysed in a dilute ethanolic solution and in the microemulsion method, TEOS is hydrolysed inside the water droplets of a water-in-oil microemulsion.[6] Both methods offer the possibility to incorporate fluorophore molecules inside the silica matrix. In the case of dye-doped silica NPs, the dye encapsulation is achieved by either covalent attachment of the dye with silica precursors (e.g. 3-(aminopropyl)triethoxysilane, APTES), before the hydrolysis in Stöber’s method, or by first solubilizing the dye in the microemulsion reaction media and then carrying out the polymerization.[9] 2.1.1. Stöber method The Stöber method is a physical chemistry process for the generation of silica particles that was discovered in 1968 by Werner Stöber and Arthur Fink.[7] This method has been used to prepare spherical silica nanoparticles with different sizes. Silica NPs are formed by the addition of TEOS to an excess of water containing ammonia and a low molar-mass alcohol such as ethanol. The resulting silica particles have diameters FCUP Fluorescent silica nanoparticles doped with organic dyes 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] 48 FCUP Fluorescent silica nanoparticles doped with organic dyes Figure 2.2 - Typical structure of a reverse micelle (source: Malik et al.[11]) In addition, during the later stages of growth, steric stabilization provided by the surfactant layer prevents the nanoparticles from aggregating.[11] Similar to Stöber’s method the particles synthesized by microemulsion technique are formed by the hydrolysis and polymerization of silane precursors in presence of ammonia. The reverse microemulsion method produces fairly uniform and monodisperse nanoparticles (Figure 2.1 B), but takes 24 to 48 hours to complete. The advantage of the reverse microemulsion method apart from the fact that it produces highly spherical and monodisperse nanoparticles of various sizes is the ability to encapsulate a wide variety of organic and inorganic fluorophores as well other materials such as luminescent quantum dots. In case of the last ones previously to encapsulation QDs have to undergo a ligand exchange process in order to make them hydrophilic. 2.1.3. Incorporation of organic fluorophores The Stöber and reverse microemulsion methods offer the possibility to incorporate fluorophore molecules inside the silica matrix either by physical entrapment or by covalent binding. The physical entrapment of dye molecules is usually obtained by adding the fluorophore to the reaction media however this incorporation method leads very often to dye leakage from the NPs. Covalent binding of the dye to the silica matrix is obtained by reaction of the dye to a silane agent such as APTES before the hydrolysis and condensation of TEOS. This approach reduces dye leakage from the silica matrix and also enables the incorporation of a variety of organic dye molecules into the silica NPs.[3] The first report on the covalent incorporation of organic fluorophores into colloidal silica NPs was made in 1992 by Van Blaaderen and co-workers.[12] They report the FCUP Fluorescent silica nanoparticles doped with organic dyes 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] 50 FCUP Fluorescent silica nanoparticles doped with organic dyes On the other hand heterogeneous nucleation occurs more often than homogeneous nucleation. Since heterogeneous nucleation takes place at preferential sites, it requires less energy than homogeneous nucleation because the effective surface energy at those sites is lower, thus diminishing the free energy barrier and this facilitates the nucleation. In this type of nucleation, some energy is released by the partial destruction of the previous surface allowing the new phase to form without the need for supersaturation.[19] 2.2.2. Growth mechanisms In classical growth theory it is assumed that particle growth occurs by molecule-by- molecule attachment to a pre-existing surface.[18] Based on this theory, the molecules diffuse onto the particle surface where it will attach itself to a suitable growth site. Alternatively to the classical growth is the growth model based on Ostwald ripening[20], which consists of a mass transfer process where smaller particles in solution dissolve and deposit on larger particles in order to reach a more thermodynamically stable state. Thus small particles decrease in size until they disappear and large particles grow even larger. This shrinking and growing of particles will result in an increase in mean particle size. Ostwald ripening is often found in water-in-oil emulsions where oil molecules will diffuse through the aqueous phase and join larger oil droplets. 2.2.3. Particle growth in the reverse micellar system Particle growth in the reverse microemulsion system is illustrated in Figure 2.3. This mechanism can be viewed as a fluid phase consisting of reverse micelles that are filled with silica particles or empty (free of particles but containing hydrolysed TEOS molecules). Additionally there is a fraction of TEOS molecules that remain in the oil phase during the reaction.[21] Particle growth can then result from the transfer of the hydrolysed TEOS in the reverse micelles to the micelles filled with silica nanoparticles. Direct interaction of TEOS that remain in the oil phase (non-hydrolysed TEOS) with the particle filled micelles can also occur. Prior to growth, hydrolysis of TEOS has to proceed in the water-shell (or hydration layer) that surrounds the particles.[21] FCUP Fluorescent silica nanoparticles doped with organic dyes 51 Figure 2.3 – Silica nanoparticles growth mechanism in a reverse micellar system composed. (Source: Osseo-Asare et al.[21]) 2.3. Surface functionalization of dye-doped silica NPs Controlling the surface chemical composition of the nanoparticles can confer them with stability, biocompatibility and enables their use in a wide range of bioapplications.[22] Due to the versatility of silica chemistry it is possible to modify the particle’s surface with various functional groups for biological applications (Figure 2.4). A variety of methods are available for particle surface modification, among them are the physical absorption and the chemical binding. Physical absorption relies on the formation of noncovalent interactions and is commonly employed to modify the silica NPs surface with avidin. Avidin is a glycoprotein with an overall positive charge, which can attach to the negatively charged silica surface through electrostatic interactions (see Figure 2.5 bottom).[1, 14] 52 FCUP Fluorescent silica nanoparticles doped with organic dyes Figure 2.4 - Schematic illustration of the surface functionalization of silica NPs for biological applications. (Source: Smith et al.[4]) The chemical binding takes advantage of the condensation of alkoxy groups of organosilanes with silanol groups on the particle surface. The particle surface is first modified with functional groups such as, thiol (-SH), amine (-NH2) and carboxyl (COOH) groups through an additional silica coating (post-coating) that contains the functional groups of interest. Afterwards biomolecules such as proteins, antibodies, oligonucleotides, etc. can be conjugated to silica nanoparticles through interaction with those functional groups following standard conjugation methods. These binding methods are listed in Table 2.1 and schematically represented in Figure 2.5. For example, thiol functionalized NPs can conjugate with dissulfide modified oligonucleotides by a dissulfide coupling chemistry while amine modified NPs can be coupled to a wide variety of haptens (small molecules that react with a specific antibody) via succinimidyl esters and isothiocyanates.[3] The carboxyl modified NPs are suitable for covalent coupling with proteins or other amine containing biomolecules trough carbodiimide chemistry.[3] In case of Stöber nanoparticles, the surface modification is usually achieved after nanoparticle synthesis to avoid potential secondary nucleation. Surface modification of microemulsion NPs in the other hand can be done in the same manner or via direct hydrolysis and co-condensation of TEOS and other organosilanes in the microemulsion reaction media.[3, 23] After the bioconjugation step, the nanoparticles can be separated from unbound biomolecules by centrifugation, dialysis, filtration, or other laboratory techniques. 53 FCUP Fluorescent silica nanoparticles doped with organic dyes Table 2.1 - Chemical binding for bioconjugation of silica NPs. (adapted from Yao et al. [2]) Organosilanes used on surface 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 54 FCUP Fluorescent silica nanoparticles doped with organic dyes Figure 2.5 - Representative bioconjugation schemes for attaching biomolecules to dye-doped silica NPs for bioanalysis. (source: Wang et al. [3]) 2.4. Nanoparticle characterization Characterization of NPs is important to elucidate the structure, characteristics and mechanism of nanoparticle formation. Evaluation of the nanoparticles in relation to their photostability, surface properties, size and morphology provides information that can be used to improve and enhance the synthesis protocol. Typical particle characterization methods include particle size and shape measurements, determination of surface charge and functionality, and determination of the optical and spectral characteristics. Chemical characterization is also important to quantify the amounts of doped dye molecules and surface-immobilized biomolecules.[3] 2.4.1. Measure particle size Several techniques are currently available for measuring particle size including transmission electron microscopy (TEM), scanning electron microscopy (SEM), atomic force microscopy (AFM) and dynamic light scattering (DLS). TEM and SEM are commonly used for size characterization of nanoparticles in vacuum, while AFM is used for both dry and wet samples at normal atmospheric pressure. DLS allows particle size measurements in aqueous media giving information on nanoparticle size distribution and relative dispersion.[3] DLS determines the hydrodynamic diameter of the NPs meaning that it measures the Brownian movement of the various particles FCUP Fluorescent silica nanoparticles doped with organic dyes 61 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., Surfacefunctionalized nanoparticles for biosensing and imaging-guided therapeutics. Nanoscale, 2013. 5(8): p. 3127-3148. 23. Deng, G., Markowitz, M.A., Kust, P.R. and Gaber, B.P., Control of surface expression of functional groups on silica particles. Materials Science & Engineering C-Biomimetic and Supramolecular Systems, 2000. 11(2): p. 165- 172. 62 FCUP Fluorescent silica nanoparticles doped with organic dyes 24. Ruedas-Rama, M.J., Walters, J.D., Orte, A. and Hall, E.A.H., Fluorescent nanoparticles for intracellular sensing: A review. Analytica Chimica Acta, 2012. 751: p. 1-23. 25. Santra, S., Zhang, P., Wang, K.M., Tapec, R. and Tan, W.H., Conjugation of biomolecules with luminophore-doped silica nanoparticles for photostable biomarkers. Analytical Chemistry, 2001. 73(20): p. 4988-4993. 26. Peng, J., Wang, K., Tan, W., He, X., He, C., Wu, P. and Liu, F., Identification of live liver cancer cells in a mixed cell system using galactose-conjugated fluorescent nanoparticles. Talanta, 2007. 71(2): p. 833-40. 27. Ow, H., Larson, D.R., Srivastava, M., Baird, B.A., Webb, W.W. and Wiesner, U., Bright and stable core-shell fluorescent silica nanoparticles. Nano Letters, 2005. 5(1): p. 113-7. 28. Shi, H., He, X.X., Wang, K.M., Yuan, Y., Deng, K., Chen, J.Y. and Tan, W.H., Rhodamine B isothiocyanate doped silica-coated fluorescent nanoparticles (RBITC-DSFNPs)-based bioprobes conjugated to Annexin V for apoptosis detection and imaging. Nanomedicine-Nanotechnology Biology and Medicine, 2007. 3(4): p. 266-272. 29. Xu, H., Aylott, J.W., Kopelman, R., Miller, T.J. and Philbert, M.A., A real-time ratiometric method for the determination of molecular oxygen inside living cells using sol-gel-based spherical optical nanosensors with applications to rat C6 glioma. Analytical Chemistry, 2001. 73(17): p. 4124-4133. 30. Burns, A., Sengupta, P., Zedayko, T., Baird, B. and Wiesner, U., Core/Shell fluorescent silica nanoparticles for chemical sensing: towards single-particle laboratories. Small, 2006. 2(6): p. 723-6. 31. Peng, J.F., He, X.X., Wang, K.M., Tan, W.H., Wang, Y. and Liu, Y., Noninvasive monitoring of intracellular pH change induced by drug stimulation using silica nanoparticle sensors. Analytical and Bioanalytical Chemistry, 2007. 388(3): p. 645-654. 32. Wang, L., Yang, C. and Tan, W., Dual-luminophore-doped silica nanoparticles for multiplexed signaling. Nano Letters, 2005. 5(1): p. 37-43. 33. Zhao, X.J., Tapec-Dytioco, R. and Tan, W.H., Ultrasensitive DNA detection using highly fluorescent bioconjugated nanoparticles. Journal of the American Chemical Society, 2003. 125(38): p. 11474-11475. 34. Zhou, X.C. and Zhou, J.Z., Improving the signal sensitivity and photostability of DNA hybridizations on microarrays by using dye-doped core-shell silica nanoparticles. Analytical Chemistry, 2004. 76(18): p. 5302-5312. FCUP 63 3. Lanthanopolyoxometalates encapsulated into silica nanoparticles Polyoxometalates (POMs) are a highly versatile and easily modified class of inorganic compounds. The diversity of structures as well as the high number of elements that can make up the structure of a polyoxometalate (POM) allows for a wide range of applications in fields such as catalysis, medicine, biology and more recently in nanotechnology.[1, 2] Lanthanide-containing polyoxometalates (LnPOMs) in particular, are readily obtained through the coordination of lanthanide ions to lacunary POMs, and exhibit interesting luminescent properties and other specific characteristics that result from the synergy between the properties of lanthanide ions and POM units.[3-5] LnPOMs have been applied in different areas such as catalysis,[6, 7] magnetism,[8, 9] luminescence[3, 10] and medicine.[11, 12] However, the use of LnPOMs in biological applications is hindered by the possible toxicity of the lanthanides and by their interaction with biological media, which may lead to coordination of biological ligands, hydrolysis, and other reactions that may adversely affect the properties of LnPOMs. One strategy to avoid possible adverse interactions with biological molecules is the encapsulation of LnPOMs into an inert nanomaterial. The high stability, chemical inertness and optical transparency of silica makes it the ideal candidate for encapsulation while preserving the properties of the encapsulated material, in particularly the optical properties.[13] Moreover, the surface of silica nanoparticles can be easily functionalized enabling their application in the preparation of biosensors and cell labeling.[14, 15] 3.1. Polyoxometalates 3.1.1. Definition Polyoxometalates (POMs) are anionic species consisted by polyhedral units of transition metal polyoxoanions (MOx, generally MO6 octahedrons – Figure 3.1), linked together by shared oxygen atoms to form a large and closed 3-dimensional framework. 64 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles Figure 3.1 - Ball-and-stick (left) and polyhedral (right) representations of the fundamental unit MO6. (Source: Fernandez[16]) The linkage of MO6 units can be done by edge and corner-sharing of MO6 octahedrons. Less often this linkage can also be accomplished by face-sharing of the MO6 octahedrons (Figure 3.2). Figure 3.2 - Representation of the three possible unions between two MO6 octahedral units: A) corner-sharing, B) edgesharing 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 metaloxide framework while the heteropolyanions apart from this framework also have an internal heteroatom X. The metal atoms that make up the framework, also called addenda atoms, are typically V, Nb, Ta, Mo and W in the V or VI oxidation states (electronic configuration d0 or d1).[17, 18] When more than one addenda atom is present in the framework the cluster is called a mixed addenda cluster. In general, any element can participate as X in a POM cluster since there are no strict physical requirements for this position, X can be a non-metal (e.g. P), a semi-metal (such as B and Si), a transition metal (e.g. Co and Fe) or a p-block metal (as Al).[19, 20] The heteroatom X, when present, is the central atom and forms a central tetrahedron XO4. The heteroatoms can be classified as primary or secondary (also called peripherals). The primary heteroatoms are indispensable for the heteropolyanions basic structure since they cannot be removed without destroying the anion. In the other hand and since they FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles 65 are not essential for the maintenance of the POM structure, the secondary heteroatoms can be removed from the heteropolyanion giving rise to other anionic stable species.[21] In Figure 3.3 are represented some examples of the two different classes of POMs. Among them, the most studied and well-known structures are the Keggin-type and Wells-Dawson. In this work, were used the Keggin-type heteropolyanions and for this reason special attention will be given in section 3.2 to this type of structure. Figure 3.3 - Polyhedral representation of common polyoxoanions: A) Lindqvist ([M6O19)n-) isopolyanion; B) Anderson- Evans ([XM6O24]n-); C) Keggin ([XM12O40]n-); D) Wells-Dawson ([X2M18O62]n-) and E) Preyssler ([XP5W30O110]n-) heteropolyanions. (Source: Lopez et al. [22]) 3.1.2. Historical context In 1826 Berzelius[23] reported the discovery of the first POM, the phosphomolybdate, of formula [PMo12O40]3-. Years later, in 1862, and after the discovery of the silicotungstic acid and its salts by Marignac[24] the analytical composition of these compounds began to be analysed. In 1929, Pauling[25] gave the first steps trying to understand the structure of POMs, proposing that the anion [PMo12O40]3- discovered by Berzelius had structure formed by a central tetrahedron XO4 surrounded by twelve MO6 octahedra sharing corners. However, in 1933 Keggin[26] solved, by X-ray diffraction, the structure of the phosphotungstic acid (H3PW12O40·5H2O) demonstrating that the anion [XM12O40]nwas 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 66 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles 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 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles 67 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 edgesharing 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] 68 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles Figure 3.5 - Polyhedral representation of the five rotational isomers of the Keggin anion. The rotated M3O13 groups are highlighted (dark blue). (source: Lopez et al.[22]) The metal-oxygen bonds present in a POM structure are arranged in such a framework that can be divided according to position of the oxygen atoms in the structure. Thus an oxygen atom linked to the central atom X (in case of the heteropolyanions) is designated by Oa, atoms that share a corner or an edge are 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. FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles 69 From the Keggin anion it is possible to obtain several lacunar structures by removing one or more MOx octahedrons.[5, 21] The monolacunar anion [XM11O39](n+4)- derives from the removal of a MO4+ unit (a metal with its terminal oxygen) by alkaline hydrolysis, giving origin to a gap with five oxygen atoms potentially coordinating (Figure 3.7). Figure 3.7 - Formation scheme of the monolacunar anion [XM11O39](n+4)- The monolacunar anion [XM11O39](n+4)- can then coordinate with metallic cations and originate complexes of the type 1:1 [XM11M’(L)O39]n- or 1:2 [M’(XM11O39)2]n- (Figure 3.8). Figure 3.8 - Representation of the complexes of the type 1:1 [XM11M’(L)O39]n- (left) and 1:2 [M’(XM11O39)2]n- (right). The complexes of the type 1:1 are formed when the metallic cation (M’) is a transitional metal (such as V3+, Mn2+ or Co2+) or an element from the p group (e.g. Al3+, Ga3+ or Ge4+). In these complexes, to maintain the octahedral coordination of the ion M’ a monodentate ligand (L) is used. In the case of lanthanide ions, since they are larger ions, bind preferentially to two lacunar units forming 1:2 complexes in which the metal coordinates through eight bonds (four with each lacunar unit). 70 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles 3.3. Polyoxometalates containing lanthanide ions Lanthanide (Ln) ions, when combined with POMs, confer additional properties, such as excellent luminescent characteristics. This property makes the lanthanidesubstituted 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). FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles 77 technological areas, such as catalysis, optical/magnetic sensors and medical imaging.[5] LnPOMs can be incorporated in several composites, namely nanostructured thin films, silica NPs, layered double hydroxides and metal-organic frameworks (MOFs). In the particular case of the Keggin type LnPOMs, this type of composite has been applied as a catalyst in the oxidation of alcohols, alkenes and aldehydes, and also as electrocatalysts and photocatalysts.[5] Griffith and coworkers[63] were the first to report the use of lanthanophosphopolyoxotungstates ([Ln(PW11O39)2)]11- with Ln3+ = La3+, Pr3+, Sm3+ and Tb3+) as oxidation catalysts in the presence of hydrogen peroxide (H2O2). These LnPOMs with H2O2 as co-oxidant catalyse the oxidation of primary and secondary alcohols to aldehydes and ketones and the epoxidation of alkenes. More recently Kholdeeva et al.[64] reported the oxidation of formaldehyde mediated by cerium (Ce) containing POM. The [CeSiW11O39]4- complex was shown to be a selective and effective catalyst for the aerobic oxidation of formaldehyde to formic acid under mild conditions, including the ambient ones. A novel application of LnPOMs as catalyst in oxidative desulfurization (ODS) process was report by Ribeiro et al.[6] In this work the authors describe the use of a LnPOM incorporated into a metal-organic-framework (MOF) as catalyst to complete desulfurization of sulphur refractory compounds from model oil. The LnPOMs-MOFs composite revealed to be an effective catalyst for ODS of oils containing refractory sulphur compounds. Furthermore these composites are recyclable which allows their use in several cycles.[6] In electrocatalysis field Cheng and coworkers[65], for example, reported the use of mixed addenda molybdotungstates coordinated with neodymium (Nd) as catalysts. In this report the authors studied the catalytic property of [Nd(SiMo7W4)]13- and this complex proved to be an efficient catalyst in the reduction of bromate to bromide in aqueous solution. POMs can also be applied as photocatalysts in degradation of organic pollutants. In the last two decades they gathered attention exhibiting potential application to degrade and mineralize organic pollutants in wastewater. An example of this is the work of Feng et al.[66] where is reported the study on photodegradation of the organic pollutant Azo dye by polyoxometalates/polyvinyl alcohol complexes. Azo dyes contain some aromatic hydrocarbons such as methyl orange, Congo red or Panceau 2R, hazardous chemicals for the water environment. The photocatalysts were prepared by using LnPOMs as the active sites and polyvinyl alcohol (PVA) as a support. The LnPOMs were first immobilized into PVA support in order to decrease water solubility of POMs and enable their recovery from the reaction system and subsequent recycling. A series of 78 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles 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. FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles 79 3.6. References 1. Liu, S.Q. and Tang, Z.Y., Polyoxometalate-based functional nanostructured films: Current progress and future prospects. Nano Today, 2010. 5(4): p. 267- 281. 2. Long, D.L. and Cronin, L., Towards polyoxometalate-integrated nanosystems. Chemistry - A European Journal, 2006. 12(14): p. 3698-706. 3. 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Faustino, W.M., Malta, O.L. and de Sa, G.F., Intramolecular energy transfer through charge transfer state in lanthanide compounds: A theoretical approach. Journal of Chemical Physics, 2005. 122(5). 84 FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles 54. D'Aleo, A., Picot, A., Beeby, A., Williams, J.A.G., Le Guennic, B., Andraud, C. and Maury, O., Efficient Sensitization of Europlum, Ytterbium, and Neodymium Functionalized Tris-Dipicolinate Lanthanide Complexes through Tunable Charge-Transfer Excited States. Inorganic Chemistry, 2008. 47(22): p. 10258- 10268. 55. Ferreira, R.A.S., Nobre, S.S., Granadeiro, C.M., Nogueira, H.I.S., Carlos, L.D. and Malta, O.L., A theoretical interpretation of the abnormal D-5(0)-> F-7(4) intensity based on the Eu3+ local coordination in the Na-9[EuW10O36] center dot 14H(2)O polyoxometalate. Journal of Luminescence, 2006. 121(2): p. 561- 567. 56. Ballardini, R., Chiorboli, E. and Balzani, V., Photophysical properties of Eu(SiW11O39)2 13- and Eu(BW11O39)2 15-. Inorganica Chimica Acta, 1984. 95(6): p. 323-327. 57. Green, M., Harries, J., Wakefield, G. and Taylor, R., The synthesis of silica nanospheres doped with polyoxometalates. Journal of the American Chemical Society, 2005. 127(37): p. 12812-12813. 58. Balula, M.S.S., Nogueira, H.I.S. and Cavaleiro, A.M.V., New Polyoxotungstates with Ln(III) and Co(II) and their Immobilization in Silica Particles. Materials Science Forum, 2006. Vols. 514-516: p. 1206-1210. 59. Granadeiro, C.M., Ferreira, R.A.S., Soares-Santos, P.C.R., Carlos, L.D., Trindade, T. and Nogueira, H.I.S., Lanthanopolyoxotungstates in silica nanoparticles: multi-wavelength photoluminescent core/shell materials. Journal of Materials Chemistry, 2010. 20(16): p. 3313-3318. 60. Sousa, J.L.C., Santos, I.C.M.S., Simoes, M.M.Q., Cavaleiro, J.A.S., Nogueira, H.I.S. and Cavaleiro, A.M.V., Iron(III)-substituted polyoxotungstates immobilized on silica nanoparticles: Novel oxidative heterogeneous catalysts. Catalysis Communications, 2011. 12(6): p. 459-463. 61. Zhao, Y.Y., Li, Y., Li, W., Wu, Y.Q. and Wu, L.X., Preparation, Structure, and Imaging of Luminescent SiO2 Nanoparticles by Covalently Grafting Surfactant- Encapsulated Europium-Substituted Polyoxometalates. Langmuir, 2010. 26(23): p. 18430-18436. 62. Zhao, Y.Y., Qi, W., Li, W. and Wu, L.X., Covalent Dispersion of Surfactant- Encapsulated Polyoxometalates and In Situ Incorporation of Metal Nanoparticles in Silica Spheres. Langmuir, 2010. 26(6): p. 4437-4442. FCUP Lanthanopolyoxometalates encapsulated into silica nanoparticles 85 63. Griffith, W.P., Moreea, R.G.H. and Nogueira, H.I.S., Lanthanide complexes as oxidation catalysts for alcohols and alkenes. Polyhedron, 1996. 15: p. 3493- 3500. 64. Kholdeeva, O.A., Timofeeva, M.N., Maksimov, G.M., Maksimovskaya, R.I., Neiwert, W.A. and Hill, C.L., Aerobic oxidation of formaldehyde mediated by a Ce-containing polyoxometalate under mild conditions. Inorganic Chemistry, 2005. 44(3): p. 666-672. 65. Cheng, L., Zhang, X.M., Xi, X.D., Liu, B.F. and Dong, S.J., Electrochemical behavior of the molybdotungstate heteropoly complex with neodymium, K10H3[Nd(SiMo7W4O39)(2)]center dot xH(2)O in aqueous solution. Journal of Electroanalytical Chemistry, 1996. 407(1-2): p. 97-103. 66. Feng, C., Zhuo, X. and Liu, X.J., Study on photodegradation of Azo dye by polyoxometalates/polyvinyl alcohol Journal of Rare Earths, 2009. 27: p. 717- 722. 67. Feng, J.H., Li, X.J., Pei, F.K., Sun, G.Y., Zhang, X. and Liu, M.L., An evaluation of gadolinium polyoxornetalates as possible MRI contrast agent. Magnetic Resonance Imaging, 2002. 20(5): p. 407-412. 68. Sun, G.Y., Feng, J.H., Wu, H.F., Pei, F.K., Fang, K. and Lei, H., Investigation of sandwiched gadolinium(III) complexes with tungstosilicates as potential MRI contrast agents. Magnetic Resonance Imaging, 2004. 22(3): p. 421-426. 69. Lehmann, J., Gaita-Arino, A., Coronado, E. and Loss, D., Spin qubits with electrically gated polyoxometalate molecules. Nature Nanotechnology, 2007. 2(5): p. 312-7. 86 FCUP FCUP Experimental Background 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 94 FCUP Experimental Background binding of RBITC to the silica matrix. For that purpose 0.1 mL of APTS was added to 2.5 mL of an anhydrous ethanol solution containing 27.5 mg of RBITC (20.5 mM) and left to react for 24 h in the dark. After that period the solution was centrifuged and the obtained powder was dried in a desiccator for further use. In a second step the reverse microemulsion was prepared by mixing 5.3 mL of Triton X-100, 5.4 mL of n-hexanol, 22.5 mL of cyclohexane, 3.0 mg of RBITC-APTES conjugate dissolved in 1.5 mL of deionized water, and 300 µL of ammonium hydroxide. The microemulsion was stirred for 30 min before 300 µL of TEOS was added. The solution was stirred for another 24 h after which 20 mL of pure acetone were added to precipitate the nanoparticles from the microemulsion. Nanoparticles were then washed by repeated cycles of centrifugation/resuspension in ethanol to remove any surfactant or unreacted molecules. The third approach is similar to the previous method described and is based in the work of Shi[6] et al. An aqueous solution of RBITC (0.1 M) was prepared and then mixed with an equimolar quantity of APTES, and left to react overnight at room temperature. The RBITC-APTES conjugate was directly used to prepare the fluorescent silica nanoparticles trough the reverse microemulsion technique as follows: 1.77 mL of Triton X-100 was mixed with 7.5 mL of cyclohexane, 1.8 mL of n-hexanol, 400 µL of water and 100 µL of the as prepared RBITC-APTES conjugate. After stirring for 1 hour, 200 µL of TEOS and 100 µL of ammonium hydroxide were then added to the previous mixture and the reaction was allowed to continue for 24 hours at room temperature. When the reaction was completed the nanoparticles were isolated by addition of 20 mL of acetone followed by a washing step through repeated cycles of centrifugation/resuspension in ethanol. FCUP Experimental Background 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. 96 FCUP Experimental Background Figure EB 3 - TEM images of fluorescent silica nanoparticles prepared by the reverse microemulsion system following the procedures of Gao[4] (A); Zhang[5] (B) and Shi[6] (C). 550 575 600 625 650 675 700 0 200 400 600 800 1000 Intensity (a.u) Wavelength (nm) A CB Figure EB 4 - Fluorescence emission spectra of fluorescent silica nanoparticles prepared by the reverse microemulsion system following the procedures of Gao[4] (A); Zhang[5] (B) and Shi[6] (C). Based on the results shown above, the experimental work presented in the next section, regarding the development of fluorescent silica nanoparticles encapsulating the organic fluorophore RBITC, was done following the adapted procedure from Shi and coworkers. In the case of fluorescent silica nanoparticles encapsulating the inorganic fluorophores (lanthanopolyoxometalates), these were also synthesized through the reverse microemulsion technique but following a procedure described by Ye[7] et al. for similar fluorescent silica nanoparticles. FCUP Experimental Background 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 timeresolved fluorometric application. Journal of Materials Chemistry, 2004. 14(5): p. 851-856. 98 FCUP FCUP 99 4. Dye doped fluorescent silica nanoparticles Incorporation of fluorescent organic molecules inside silica matrixes shields the molecules from several environmental factors that can interfere with their fluorescence emission, photostability or quantum yield. For these reasons fluorescent dye molecules are widely used nowadays doped into silica matrixes to produce fluorescent nanomaterials. Dye doped fluorescent silica nanoparticles containing RBITC dye molecules were synthesized using the reverse microemulsion technique for the alkaline hydrolysis of TEOS. In this particularly case of silica nanoparticles containing RBITC dye molecules, the dye was firstly coupled to a silane coupling agent (3-aminopropyl triethoxysilane – APTES), and the reaction product was incorporated into silica spheres by hydrolysis and polymerisation of TEOS in alkaline media. The obtained nanoparticles were then characterized by fluorescence and ultraviolet-visible (UV-Vis) spectroscopy, transmission electron microscopy (TEM) and dynamic light scattering (DLS). To investigate the influence of silica encapsulation in the fluorescence emission properties of the dye, lifetime measurements and steady state anisotropy studies were perform for both nanoparticles and the free dye in solution. The produced RBITC encapsulated silica nanoparticles had a mean diameter of around 64 nm. Fluorescence and UV-Vis spectra of the same nanoparticles show the typical fluorescence and absorption band of RBITC indicating that silica encapsulation does not interfere with the emission properties of the fluorophore. It was also noticed that silica encapsulation improved RBITC quantum yield and fluorescence lifetime when compared to free RBITC in solution. Particle’s surface have also been modified, so that the particles could bind to biologically active molecules, such as oligonucleotides. 4.1. Material and Methods 4.1.1 Chemicals Rhodamine b isothiocyanate (mixed isomers Aldrich), rhodamine b (dye content ~95% Sigma), (3-aminopropyl)triethoxysilane (≥98% Sigma-Aldrich), water (molecular 100 FCUP Dye doped fluorescent silica nanoparticles biology reagent Sigma), tetraethoxysilane (99% Aldrich), Triton X-100 (Aldrich), 1- hexanol (98% Merck), cyclohexane (99% Aldrich), ammonia (25% Merck), ethanol (99.5% Panreac), acetone (99.9% Fluka), acetonitrile (99.9% ROMIL), (3- glycidyloxypropyl)trimethoxysilane (99% Aldrich), potassium phosphate monobasic (≥99% Sigma-Aldrich), potassium phosphate dibasic (≥99% Sigma-Aldrich), were used as received. Oligonucleotide sequences (5’-gat cgc ctc cac gtc c-3’) were acquired from STAB vida (Lisbon, Portugal) and were purified through a NAP-5 column from GE Healthcare (UK) before use. 4.1.2 Instrumentation and methodologies 4.1.1.1. Elemental Analysis Elemental analysis for carbon and hydrogen were performed on a Leco CHNS-932, the technique was carried out in the University of Santiago de Compostela. 4.1.1.2. UV-visible spectroscopy Absorption spectra were observed on a Varian Cary bio50 spectrophotometer, using quartz cells with 1 cm path length. Absorption spectra of RBITC doped silica nanoparticles were fitted using a second order exponential decay in order to remove the background scattering from silica. Based on the absorbance, the amount of RBITC molecules was calculated with an assumption that the molar absorption coefficient (ε) of RBITC molecules are equal in a RBITC solution and a RBITC doped nanoparticle solution if they have the same absorbance. Using the known values including the density of silica matrix (2.2 g mL-1), the concentration and the size of the nanoparticles, the amount of RBITC molecules in each nanoparticle was calculated.[1, 2] 4.1.1.3. Fluorescence spectroscopy, quantum yield and lifetime Fluorescence measurements were performed in a Varian Cary Eclipse spectrofluorometer, equipped with a constant-temperature cell holder (PeltierMulticell Holder). Fluorescence quantum yield determination was done as described by Fery-Forgues and Lavabre.[3] Absorption spectra were recorded with a Varian Cary bio50 spectrophotometer, equipped with a Varian Cary single cell Peltier accessory, using FCUP 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 102 FCUP Dye doped fluorescent silica nanoparticles were carried out in the chemistry department of the Faculty of Sciences and Technologies from the University Nova de Lisboa. 4.1.1.5. Transmission electron microscopy TEM images were obtained using a HITACHI H-8100 instrument operating at an acceleration voltage of 200 kV. Samples for TEM analysis were prepared by depositing ethanol suspensions of the nanoparticles on carbon coated copper grids and allowing them to completely dry. TEM images were analysed using image J software (version 1.44p), available through http://imagej.nih.gov/ij. TEM was carried out in the institute of materials and surfaces science and engineering (ICEMS) from Instituto Superior Técnico (IST). 4.1.1.6. Dynamic light scattering and zeta potential DLS measurements were performed at 25ºC, using a Malvern ZetasizerNanoZS compact scattering spectrometer with a 4.0 mW He-Ne laser (633 nm wavelength) at a scattering angle of 173º. The average hydrodynamic diameter and the size distribution of the samples were determined using Malvern Dispersion Technology Software 5.10. For zeta potential measurements NPs were redispersed in potassium phosphate buffer (10 mM, pH=8) and analysed in disposable polystyrene zeta potential cuvettes with gold-coated electrodes (Malvern) at 25.0 ºC. All measurements were repeated five times to verify the reproducibility of the results. 4.1.3 Preparation of core-shell nanoparticles with rhodamine B isothiocyanate (RBITC-APTES@SiO2) Nanoparticles were synthesized using the reverse microemulsion technique for the alkaline hydrolysis of tetraethoxysilane (TEOS) following a procedure described in literature.[4] For the silica nanoparticles containing rhodamine b isothiocyanate (RBITC) molecules, the dye was firstly coupled to a silane coupling agent (3-aminopropyl triethoxysilane – APTES), and the reaction product was incorporated into silica spheres by hydrolysis and polymerization of TEOS in alkaline media. RBITC dye (1.0x10-4 mol) was prepared in aqueous solution and then mixed with an equimolar quantity of APTES, reacting overnight at room temperature. The monomer precursor RBITC-APTES was directly used to prepare silica-coated fluorescent FCUP Dye doped fluorescent silica nanoparticles 109 Figure 4.4 shows RBITC and RBITC FSNPs absorption spectra in ethanol at 25 ºC with a final concentration with almost the same absorbance value of approximately 0.09. Free RBITC molecules in solution and RBITC FSNPs show a similar broad absorption band. The absorption spectrum of RBITC presents a maximum at 542 nm wavelength while the one for RBITC FSNPs is at 555 nm wavelength. This small shift to a higher wavelength could be due to the silica matrix being a less polar medium than ethanol, causing a red shift of the spectrum maximum of RBITC dye molecules.[13, 14] Similar findings were reported for silica nanoparticles encapsulating tetramethylrhodamine-dextran (TMR-Dex); tetramethylrhodamine-5-isothhiocyanate (TRICT) modified with APTES and Rubpy when compared to the free dyes in water.[13, 14] Regarding the possible effect that conjugation of APTES to RBITC dye could have in the emission properties of the dye, the UV-vis spectrum of a RBITC-APTES conjugate solution was recorded and compared to that of the RBITC dye in solution (Figure 4.5). Both solutions were prepared in absolute ethanol with a concentration of 6.7x10-7 M and the spectra were recorded at 25 ºC. As shown in Figure 4.5 the UV-vis spectrum of RBITC-APTES conjugate presents a broad absorption band with a maximum absorbance at 542 nm similar to UV-vis spectrum of RBITC. This indicates that the RBITC emission properties remain unchanged after conjugation with APTES 350 400 450 500 550 600 650 700 0.00 0.01 0.02 0.03 0.04 0.05 Abs Wavelength (nm) RBITC RBITC-APTES Figure 4.5 - UV-vis spectra of RBITC and RBITC-APTES conjugate in ethanol at 25 ºC. 110 FCUP Dye doped fluorescent silica nanoparticles The amount of RBITC dye in FSNPs was determined quantitatively by UV-Vis spectroscopy by comparison with the free dye in solution. The quantity of RBITC dye molecules per NP is equal to the RBITC dye weight per unit volume (derived through the absorption), divided by the number of NPs per unit volume (determined by TEM). Table 4.2 shows the results obtained for the determination of the amount of RBITC dye molecules per FSNP. These determinations were made for FSNP samples with similar size (a and b from the same batch; c and d from different batches relatively to the former one), and with different mass concentrations of NPs (0.8 mg/mL and 1.6 mg/mL). Table 4.2 – Amount of RBITC dye molecules per fluorescent silica nanoparticle 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 FCUP Dye doped fluorescent silica nanoparticles 111 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 Intensity (a.u.) Wavelength (nm) Figure 4.6 - Fluorescence excitation spectra of RBITC, RBITC-APTES conjugate and FSNPs recorded at 25 ºC in absolute ethanol. 112 FCUP Dye doped fluorescent silica nanoparticles 540 560 580 600 620 640 660 680 700 0 200 400 600 800 1000 RBITC RBITC-APTES FSNPs Intensity (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 FCUP Dye doped fluorescent silica nanoparticles 113 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 RBITCAPTES@SiO2 FSNPs. Figure 4.8 - (A) RBITC doped fluorescent silica NPs prepared by hydrolysis and polymerization of TEOS in a microemulsion method; (B) bare silica NPs with RBITC dye molecules adsorbed onto the nanoparticle’s surface; (C) fluorescent core-shell NPs with a silicon core and a shell of RBITC dye molecules and TEOS. Table 4.3 - Fluorescence quantum yields of RBITC, RBITC-APTES conjugate and FSNPs 114 FCUP Dye doped fluorescent silica nanoparticles Sample Quantum yield (Φ)a SD RBITC 0.24 0.03 RBITC-APTES 0.23 0.04 RBITC-APTES@SiO2 FSNPs 0.34 0.04 Ads:RBITC-APTES FSNPs 0.09 0.02 Shell:RBITC-APTES@SiO2 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] FCUP Dye doped fluorescent silica nanoparticles 115 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 RBITCAPTES@SiO2 FSNPs (both dye adsorbed and covalently bound to the silica matrix) the fluorescence decay could only be fitted by a 2nd exponential decay, indicating two different microenvironments around RBITC molecules (two lifetimes values see Table 4.4). The lifetime of the free dye was measured to be 2.44 ns in absolute ethanol while RBITC-APTES conjugate and the NPs exhibit two-component lifetime behaviour with a low (LC) and a high (HC) lifetime components of 1.04 ns (LC) and 2.99 (HC) for RBITC-APTES conjugate; 0.87 ns (LC) and 2.66 ns (HC) for NPs with dye adsorbed (Ads:RBITC-APTES@SiO2) and 1.27 ns (LC) and 3.44 ns (HC) for NPs with dye covalently bound (RBITC-APTES@SiO2). Table 4.4 - Lifetime data of RBITC and fluorescent silica nanoparticles (FSNPs) in absolute ethanol Sample FIT Τ1 (ns) Τ2 (ns) Χ2 RBITC 2.44 --- 1.16 RBITC-APTES 2.99 1.04 1.20 RBITC-APTES@SiO2 FSNPs 3.44 1.27 1.12 Ads:RBITC-APTES@SiO2 FSNPs 2.66 0.87 1.35 116 FCUP Dye doped fluorescent silica nanoparticles 200 400 600 800 1000 10 100 1000 Log10 I Time (ns) A T(RBITC) = 2.44 200 400 600 800 1000 10 100 1000 T2(RBITC-APTES) = 1.04 Log10 I time (ns) B T1(RBITC-APTES) = 2.99 200 400 600 800 1000 10 100 1000 T2(RBITC-APTES@SiO2) = 1.27 Log10 I Time (ns) C T1(RBITC-APTES@SiO2) = 3.44 200 400 600 800 1000 10 100 1000 T2(Ads:RBITC-APTES@SiO2) = 0.87 Log10 I Time (ns) D T1(Ads:RBITC-APTES@SiO2) = 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 FCUP Dye doped fluorescent silica nanoparticles 117 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. 118 FCUP Dye doped fluorescent silica nanoparticles O N+CH3 CH3 NH3C CH3 NH COOH CNH Si O O O S Si O O O NH CNH 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:RBITCAPTES@SiO2) the two-component lifetime (0.87 ns LC and 2.66 ns HC) are smaller compared to free RBITC-APTES conjugate in solution. In this case the smaller and higher lifetime components are also related to the RBITC-APTES conjugation. For the longer lifetime decay (2.66 ns) suggested to be related with APTES conjugate with RBITC 6-isomer, the value is similar to the free dye in solution (2.99 ns), indicating that the adsorbed RBITC 6-isomer presents a free RBITC-APTES conjugate behaviour. Regarding the smaller lifetime decay (0.87 ns) this is suggested to be related with APTES conjugate with RBITC 5-isomer and therefore with the effects of high electronic density near dye molecules and effects of charge transfer mentioned previously. The fact that these values are smaller compared to the free RBITC-APTES conjugate could be related to the fact that the dye is adsorbed to the surface and wobbling with the solvent around the NP.[22] This means that the adsorbed RBITC-APTES conjugate have a movement restriction and when associated with a high electronic density the dye FCUP Dye doped fluorescent silica nanoparticles 125 adsorbed) and a layer containing dye and silica (FSNPs with dye covalently bound). Results obtained showed that when the dye is adsorbed to NPs surface (Ads:RBITCAPTES@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:RBITCAPTES@SiO2) were shown to be similar to that of RBITC-APTES FSNPs suggesting that the dye is embedded in the silica matrix and it is protected from the outer environment. Furthermore these two types of FSNPS with dye covalently bound to the silica matrix (RBITC-APTES@SiO2 and Shell:RBITC-APTES@SiO2) were optically very similar. Fluorescence lifetime decay and steady state fluorescence anisotropy of RBITC also increase with silica encapsulation. The results obtained suggest that the dye is encapsulated within the silica matrix corroborating the previous results on the quantum yield. Furthermore dye distribution inside the NP can be classified according to the interaction of dye isomers with the adjacent APTES molecules presenting two different environments each corresponding to a small and a high lifetime component. The reaction between RBITC and APTES to produce RBITC-APTES conjugate is accomplished through the isothiocyanate functional group (NCS) in the dye with the amine group (NH2) from the silane. Depending on the position of NCS group in the isomers, the coupling with APTES can occur in different positions of the aromatic ring and thus influence dye lifetime due to the interaction of the dye with the silane adjacent molecules that can quench the dye fluorescence. The smaller and largest lifetimes are associated with APTES conjugate with RBITC 5-isomer and RBITC 6-isomer respectively. In the case of APTES conjugated with RBITC 5-isomer there is a high electronic density around the dye molecules which can quench dye fluorescence by effects of charge transfer. Regarding the conjugation of APTES with RBITC 6-isomer the electronic density around dye molecules is lower compared to the former one and therefore the effects of charge transfer are minor. Moreover increase of dye lifetime after silica encapsulation again suggests dye shielding due to the silica matrix. With respect to steady state fluorescence anisotropy the large increase in emission anisotropy from RBITC in solution (0.025) to RBITC encapsulated in silica NPS (0.187), indicates that the motion of RBITC molecules in silica is strongly restricted due to the confinement inside the NP. The motion of the encapsulated dye molecules is described by a wobbling-in-cone model. A similar behaviour is observed for the NPs with the dye 126 FCUP Dye doped fluorescent silica nanoparticles adsorbed to the silica surface. For these NPs the dye rotational motion is similar to that of the free dye in solution, suggesting that the dye is physically bound but wobbling with the solvent around the NPs. Lifetime increase value denotes the rigid environment of RBITC dye molecules within silica NPs in accordance with the obtain steady state fluorescence anisotropy values. The particle surfaces were also modified with an organosilane (GPTES) in order to allow the biological binding of the NPs to oligonucleotides. C and H elemental analysis revealed that the resulting functionalized NPs (RBITC-APTES@GPTEsSiO2) contain 0.20 mmol of GPTEs per 1 g of material. The oligonucleotide-modified silica nanoparticles were prepared by covalent immobilization of thiolated oligonucleotides onto the silica nanoparticles surface. UV-vis spectroscopy was used to check if immobilization occurred but the technique seems not to be sensitive enough to prove that the binding occurred. Zeta potential measurements shown a difference in particle’s surface after reaction with DNA that could indicate immobilization, however to prove that reaction actually took place further experiments will be needed. FCUP Dye doped fluorescent silica nanoparticles 127 4.3. References 1. Chen, G.W., Song, F.L., Wang, X., Sun, S.G., Fan, J.L. and Peng, X.J., Bright and stable Cy3-encapsulated fluorescent silica nanoparticles with a large Stokes shift. Dyes and Pigments, 2012. 93(1-3): p. 1532-1537. 2. He, X., Chen, J., Wang, K., Qin, D. and Tan, W., Preparation of luminescent Cy5 doped core-shell SFNPs and its application as a near-infrared fluorescent marker. Talanta, 2007. 72(4): p. 1519-26. 3. Fery-Forgues, S. and Lavabre, D., Are fluorescence quantum yields so tricky to measure? A demonstration using familiar stationery products. Journal of Chemical Education, 1999. 76(9): p. 1260-1264. 4. Shi, H., He, X.X., Wang, K.M., Yuan, Y., Deng, K., Chen, J.Y. and Tan, W.H., Rhodamine B isothiocyanate doped silica-coated fluorescent nanoparticles (RBITC-DSFNPs)-based bioprobes conjugated to Annexin V for apoptosis detection and imaging. Nanomedicine-Nanotechnology Biology and Medicine, 2007. 3(4): p. 266-272. 5. Santra, S., Zhang, P., Wang, K.M., Tapec, R. and Tan, W.H., Conjugation of biomolecules with luminophore-doped silica nanoparticles for photostable biomarkers. Analytical Chemistry, 2001. 73(20): p. 4988-4993. 6. He, X., Duan, J., Wang, K., Tan, W., Lin, X. and He, C., A novel fluorescent label based on organic dye-doped silica nanoparticles for HepG liver cancer cell recognition. Journal of Nanoscience and Nanotechnology, 2004. 4(6): p. 585-9. 7. He, X.X., Wang, K.M., Tan, W.H., Li, J., Yang, X.H., Huang, S.S., Li, D. and Xiao, D., Photostable luminescent nanoparticles as biological label for cell recognition of system lupus erythematosus patients. Journal of Nanoscience and Nanotechnology, 2002. 2(3-4): p. 317-320. 8. Rosi, N.L. and Mirkin, C.A., Nanostructures in biodiagnostics. Chemical Reviews, 2005. 105(4): p. 1547-62. 9. Knopp, D., Tang, D.P. and Niessner, R., Bioanalytical applications of biomolecule-functionalized nanometer-sized doped silica particles. Analytica Chimica Acta, 2009. 647(1): p. 14-30. 10. Santra, S., Wang, K., Tapec, R. and Tan, W., Development of novel dye-doped silica nanoparticles for biomarker application. Journal of Biomedical Optics, 2001. 6(2): p. 160-6. 11. Pereira, C., Biernacki, K., Rebelo, S.L.H., Magalhaes, A.L., Carvalho, A.P., Pires, J. and Freire, C., Designing heterogeneous oxovanadium and copper 128 FCUP Dye doped fluorescent silica nanoparticles acetylacetonate catalysts: Effect of covalent immobilisation in epoxidation and aziridination reactions. Journal of Molecular Catalysis a-Chemical, 2009. 312(1- 2): p. 53-64. 12. Mahajan, S., Sethi, D., Seth, S., Kumar, A., Kumar, P. and Gupta, K.C., Construction of Oligonucleotide Microarrays (Biochips) via Thioether Linkage for the Detection of Bacterial Meningitis. Bioconjugate Chemistry, 2009. 20(9): p. 1703-1710. 13. Liang, S., Shepard, K., Pierce, D.T. and Zhao, J.X., Effects of a nanoscale silica matrix on the fluorescence quantum yield of encapsulated dye molecules. Nanoscale, 2013. 5: p. 9365-9373. 14. Ma, D.L., Kell, A.J., Tan, S., Jakubek, Z.J. and Simard, B., Photophysical Properties of Dye-Doped Silica Nanoparticles Bearing Different Types of Dye- Silica Interactions. Journal of Physical Chemistry C, 2009. 113(36): p. 15974- 15981. 15. Ferrie, M., Pinna, N., Ravaine, S. and Vallee, R.A.L., Wavelength-dependent emission enhancement through the design of active plasmonic nanoantennas. Optics Express, 2011. 19(18): p. 17697-17712. 16. Tsou, C.J., Chu, C.Y., Hung, Y. and Mou, C.Y., A broad range fluorescent pH sensor based on hollow mesoporous silica nanoparticles, utilising the surface curvature effect. Journal of Materials Chemistry B, 2013. 1: p. 5557-5563. 17. Casey, K.G. and Quitevis, E.L., Effect of solvent polarity on nonradiative processes in xanthene dyes: Rhodamine B in normal alcohols. J. Phys. Chem., 1988. 92(23): p. 6590–6594 18. Cohen, B., Martin, C., Iyer, S.K., Wiesner, U. and Douhal, A., Single Dye Molecule Behavior in Fluorescent Core-Shell Silica Nanoparticles. Chemistry of Materials, 2012. 24(2): p. 361-372. 19. Rampazzo, E., Bonacchi, S., Montalti, M., Prodi, L. and Zaccheroni, N., Selforganizing core-shell nanostructures: Spontaneous accumulation of dye in the core of doped silica nanoparticles. Journal of the American Chemical Society, 2007. 129(46): p. 14251-14256. 20. Roy, S., Woolley, R., MacCraith, B.D. and McDonagh, C., Fluorescence lifetime analysis and fluorescence correlation spectroscopy elucidate the internal architecture of fluorescent silica nanoparticles. Langmuir, 2010. 26(17): p. 13741-6. 21. Santra, S., Liesenfeld, B., Bertolino, C., Dutta, D., Cao, Z.H., Tan, W.H., Moudgil, B.M. and Mericle, R.A., Fluorescence lifetime measurements to FCUP Dye doped fluorescent silica nanoparticles 129 determine the core-shell nanostructure of FITC-doped silica nanoparticles: An optical approach to evaluate nanoparticle photostability. Journal of Luminescence, 2006. 117(1): p. 75-82. 22. Yip, P., Karolin, J. and Birch, D.J.S., Fluorescence anisotropy metrology of electrostatically and covalently labelled silica nanoparticles. Measurement Science & Technology, 2012. 23(8). 23. Larson, D.R., Ow, H., Vishwasrao, H.D., Heikal, A.A., Wiesner, U. and Webb, W.W., Silica nanoparticle architecture determines radiative properties of encapsulated fluorophores. Chemistry of Materials, 2008. 20(8): p. 2677-2684. 24. Kinosita, K., Jr., Ikegami, A. and Kawato, S., On the wobbling-in-cone analysis of fluorescence anisotropy decay. Biophysical Journal, 1982. 37(2): p. 461-4. 25. Escorihuela, J., Banuls, M.J., Puchades, R. and Maquieira, A., Development of oligonucleotide microarrays onto Si-based surfaces via thioether linkage mediated by UV irradiation. Bioconjugated Chemistry, 2012. 23(10): p. 2121-8. FCUP 130 FCUP 131 5. Europium polyoxometalates encapsulated into silica nanoparticles The incorporation of europium-polyoxometalates into silica nanoparticles can lead to a biocompatible nanomaterial with suitable luminescent properties for applications in biosensors, biological probes and imaging. Europium Keggin-type polyoxometalates Eu(PW11)x (x = 1 and 2) with different europium coordination environments were prepared using simple methodologies and no expensive reactants. These luminescent compounds were then encapsulated into silica nanoparticles for the first time through the water-in-oil microemulsion methodology with a non-ionic surfactant. The europium-polyoxometalates and the nanoparticles were characterized using several techniques (FT-IR, FT-Raman, 31P MAS NMR, TEM-EDS, AFM, DLS and ICP analysis). The stability of the material and the integrity of the europium compounds incorporated were also examined. Furthermore, the photo-luminescence properties of nanomaterials Eu(PW11)x@SiO2 were evaluated and compared with the free europium-polyoxometalates. The silica surface of the most stable nanoparticles was successfully functionalized with appropriate organosilanes to enable covalent binding of oligonucleotides. 5.1. Materials and Methods 5.1.1. Chemicals Sodium tungstate ( 99 % Sigma), sodium hydrogen phosphate ( 97 % Sigma), europium chloride (99.9% Sigma), hydrochloric acid (37% Panreac), potassium chloride (> 99.5 % Merck), tetraethoxysilane (99% Aldrich), Triton X-100 (Aldrich), 1- hexanol (98% Merck), cyclohexane (99% Aldrich), ammonia (25% Merck), ethanol (99.5% Panreac), acetone (99.9% Fluka), acetonitrile (99.9% ROMIL), (3- glycidyloxypropyl)trimethoxysilane (99% Aldrich) and (3-chloropropyl)trimethoxysilane (99% Aldrich), were used as received. 132 FCUP Europium polyoxometalates encapsulated into silica nanoparticles 5.1.2. Instrumentation and methodologies 5.1.2.1. Elemental analysis Elemental analysis for K, W, Eu and P was performed by ICP-MS on a Varian 820- MS and C and H analysis was executed on a Leco CHNS-932, both techniques were carried out in the University of Santiago de Compostela. Hydration water contents were determined by thermogravimetric analysis performed in air between 30 °C and 700 °C, with a heating speed of 5 ºC/min, using a TGA-50 Shimadzu thermobalance. Thermogravimetry technique was carried out in the center for research in ceramics and composite materials (CICECO) associated laboratory from the University of Aveiro by Dr. Duarte Ananias. 5.1.2.2. Vibrational Spectroscopy Fourier transform Infrared (FT-IR) absorption spectra were obtained on a Mattson 7000 FT-IR spectrometer. Spectra were collected in the 400–4000 cm-1 range, using a resolution of 4 cm-1 and 64 scans. Fourier transform Raman (FT-Raman) spectra were recorded on a RFS-100 Bruker FT spectrometer, equipped with a Nd:YAG laser with excitation wavelength at 1064 nm, with laser power set to 350 mW. The FT-Raman studies were carried out in the CICECO associated laboratory by Dr. Carlos Granadeiro 5.1.2.3. Solid state NMR 31P MAS NMR spectra were recorded for liquid solutions using a Bruker Avance III 400 spectrometer and chemical shift are given with respect to external 85% H3PO4. The 31P NMR MAS solid-state measurements were performed in a 7 T (300 MHz) AVANCE III Bruker spectrometer under a magic angle spinning of 15 KHz at room temperature. The spectra were obtained by a solid echo sequence with an echo delay of 15 microseconds, a 90 degree pulse of 10.5 microseconds at a power of 20 W and a relaxation delay of 30 seconds. Potassium phosphate (K3PO4) was used as reference. This technique was carried out in the department of science materials (CENIMAT/I3N) of the Faculty of Sciences from University Nova de Lisboa by Dr. Gabriel Feio. FCUP Europium polyoxometalates encapsulated into silica nanoparticles 133 5.1.2.4. Transmission electron microscopy TEM images were obtained using a HITACHI H-8100 instrument operating at an acceleration voltage of 200 kV and energy dispersive X-ray spectroscopy (EDS) analysis was performed on a ThermoNoran spectrometer. Samples for TEM analysis were prepared by depositing ethanol suspensions of the nanoparticles on carbon coated copper grids and allowing them to completely dry. TEM images were analysed using image J software (version 1.44p), available through http://imagej.nih.gov/ij. TEM was carried out in the institute of materials and surfaces science and engineering (ICEMS) from Instituto Superior Técnico (IST). 5.1.2.5. Scanning electron microscopy SEM analysis and EDS elemental mapping were performed on a Hitachi SU-70 instrument operating at an acceleration voltage of 30 kV. Samples for SEM analysis were prepared by depositing ethanol suspensions of the nanoparticles on carbon coated copper grids and allowing them to completely dry. Both techniques were carried out in CICECO associated laboratory by Dr. Duarte Ananias. 5.1.2.6. Dynamic light scattering DLS measurements were performed at 25ºC, using a Malvern ZetasizerNanoZS compact scattering spectrometer with a 4.0 mW He-Ne laser (633 nm wavelength) at a scattering angle of 173º. The average hydrodynamic diameter and the size distribution of the samples were determined using Malvern Dispersion Technology Software 5.10. All measurements were repeated five times to verify the reproducibility of the results. Stable samples were prepared by dissolving the potassium salts of Eu(PW11)x in Millipore water. 5.1.2.7. Atomic force microscopy Samples for atomic force microscopy (AFM) were prepared by drying onto freshly cleaved mica substrates from dilute aqueous solutions. AFM measurements were made using an AFM Workshop TT-AFM instrument in vibrating (intermittent contact) mode. A small (15 µm) scanner and low gains were used to ensure high resolution. Probes from AppNano (ACT) with resonant frequency of around 300 kHz were used. Images of around 2 µm x 2µm were acquired, and analysed using Gwyddion software 134 FCUP Europium polyoxometalates encapsulated into silica nanoparticles and a custom routine to measure accurately the height of nanoparticles. AFM measurements were carried out by Dr. Peter Eaton. 5.1.2.8. X-ray crystallography Crystalline K11[Eu(PW11O39)2.xH2O materials suitable for single-crystal X-ray diffraction analysis were harvested and mounted in a CryoLoop using viscous oil.[1] Diffraction data were collected at 150 K on a Bruker X8 Kappa APEX II charge-coupled device (CCD) area-detector diffractometer (Mo K graphite-monochromated radiation) using the APEX2 software[2], and equipped with an Oxford Cryosystems Series 700 cryo stream controlled by the Cryopad interface.[3] Images were processed with SAINT+ software[4], and the absorption corrections were performed by the multi-scan method implemented in SADABS.[5] The structure was solved by direct methods implemented in SHELXS-97[6, 7], allowing the immediate identification of most of the heaviest elements, namely Eu and W atoms, while the remaining atoms were positioned through successive full-matrix least squares refinement cycles on F2 using SHELXL-97.[6, 8] All atoms of the Europium-phosphotungstate anions and the K+ cations were refined using anisotropic displacement parameters, while the oxygen atoms of crystallization water molecules were refined with isotropic parameters. Although the H- atoms of the water molecules were not located from difference Fourier maps or positioned in calculated positions, they were added to the molecular formula of the compound. X-ray crystallography analysis was carried out by Dr. Luis Cunha Silva. 5.1.2.9. Photoluminescence and lifetime measurements Emission and excitation spectra were recorded at 298 K and 14 K using a Fluorolog-2® Horiba Scientific (Model FL3-2T) spectroscope, with a modular double grating excitation spectrometer (fitted with a 1200 grooves/mm grating blazed at 330 nm) and a TRIAX 320 single emission monochromator (fitted with a 1200 grooves/mm grating blazed at 500 nm, reciprocal linear density of 2.6 nm∙mm-1), coupled to a R928 Hamamatsu photomultiplier, using the front face acquisition mode. The excitation source was a 450 W Xe arc lamp. Emission spectra were corrected for detection and optical spectral response of the spectrofluorimeter and the excitation spectra were corrected for the spectral distribution of the lamp intensity using a photodiode reference detector. Lifetime measurements were carried out using a 1934D3 phosphorimeter coupled to the Fluorolog®-3, and a Xe-Hg flash lamp (6 μs/pulse half width and 20-30 μs tail) was used as the excitation source. The variable pressure measurements were FCUP Europium polyoxometalates encapsulated into silica nanoparticles 141 with the formulas K4[PW11Eu(H2O)3O39)].4H2O and K11[Eu(PW11O39)2].5H2O for EuPW11 and Eu(PW11)2 respectively. Figure 5.2 - Thermogravimetric curves of EuPW11 (in blue) and Eu(PW11)2 (in red). Both compounds exhibit one step of weight loss, but over different temperature ranges. The TG of Eu(PW11)2 shows weight loss in the range of 50 – 150 ºC attributed to the release of crystal water: 1.4% (the calculated value for 5 water molecules is 1.5%). A more extensive weight loss was found for EuPW11, 4.3% (the calculated value for 7 water molecules is 4.0%), in the range 50 - 230 ºC. The weight loss in the range 150 – 230 ºC (experimental result: 1.2%; calculated for 3 water molecules: 1.5%) is typical of coordinated water molecules whereas the remaining weight loss is due to crystallization water. 5.2.1.3. 31P NMR spectroscopy 31P NMR spectroscopy was also used to identify and to characterize the monosubstituted 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 0200 400 600 800 Weight loss (%) Temperature ºC 142 FCUP Europium polyoxometalates encapsulated into silica nanoparticles Figure 5.3 - 31P 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 europiumpolyoxometalates 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 europiumpolyoxometalates. 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 0.36 -10.10 PW11 EuPW11 Eu(PW11)2 (ppm) 5.58 FCUP Europium polyoxometalates encapsulated into silica nanoparticles 143 encapsulated nanoparticles Eu(PW11)2@SiO2 were functionalized with two organosilanes: (3-glycidyloxypropyl)-trimethoxysilanes (GPTEs) and (3-chloropropyl)- trimethoxysilanes (CPTEs), by reaction of the hydroxyl groups of their surface, in a post-synthesis step. 5.2.2.1. Transmission Electron Microscopy TEM images of the nanocomposites of silica doped with Eu(PW11)x (x = 1 or 2) show uniform nanosized spheres with a core-shell structure (Figure 5.4). As expected, the EDS analysis revealed that the imaged nanoparticles have europium tungsten and silica in their constitution (Figure 5.5). The EuPW11@SiO2 and Eu(PW11)2@SiO2 nanoparticles prepared using equal weight of europium-polyoxometalates have a mean diameter of around 16 ± 1.9 nm and 51 ± 5.8 nm, respectively (calculated from more than 100 randomly selected nanoparticles on the TEM grid). Figure 5.4 - TEM images of (a,b) EuPW11@SiO2 and (d,e) Eu(PW11)2@SiO2 nanoparticles showing the core/shell structure (both materials prepared using 50 mg of corresponding europium compounds); (c,f) Size distribution histograms of EuPW11@SiO2 and Eu(PW11)2@SiO2 nanoparticles respectively. 144 FCUP Europium polyoxometalates encapsulated into silica nanoparticles 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 Cu W EuEu W Counts KeV CO CuWPEu K Si Cu EuPW11@SiO2 0.0 2.5 5.0 7.5 10.0 12.5 0 100 200 300 400 500 CuWW Eu W Cu W Counts KeV C O Si CuW PKEu Eu(PW11)2@SiO2 FCUP Europium polyoxometalates encapsulated into silica nanoparticles 145 Figure 5.6 - AFM topography and amplitude images respectively of EuPW11 (a,b) and Eu(PW11)2 (c,d). Topography images show the presence of features with dimensions larger than expected for single POMs (i.e. larger than 2 nm). The formation of monodisperse colloidal nanoparticles involves two sequential steps: nucleation and growth. According to Vanblaaderen and co-workers,[35, 36] particle growth occurs through monomer addition, with the growth rate being controlled by the rate of alkoxide hydrolysis. Polydispersity and final particle size can be determined by the balance between monomer addition and nucleation.[37] Increasing the ratio of europium-polyoxometalate (nuclei) / TEOS (monomer), increases the concentration of seeds competing for the monomer in the growing process and could thus lead to a reduction of the final particle size. To clarify the relation between the nanoparticle size and the molar quantity of Eu(PW11)x present in the reaction medium, another preparation of europium-polyoxometalate nanoparticles was performed with the sandwich-type Eu(PW11)2. Silica nanoparticles were synthesized following the same reverse microemulsion methodology but using higher amount (95 mg) of Eu(PW11)2, which corresponds to the same molar amount (16 µmol) used before for the preparation of EuPW11@SiO2. In this case, the Eu(PW11)2@SiO2 nanocomposites obtained had a mean diameter of 28 nm, as determined by TEM (Figure 5.7). 146 FCUP Europium polyoxometalates encapsulated into silica nanoparticles Compared to Eu(PW11)2@SiO2 nanoparticles prepared using 8 µmol of Eu(PW11)2 these results suggest that the molar quantity of europium-phosphotungstate does have a relevant influence on the silica nanoparticle final size, with larger concentrations of POMs leading to greater numbers of seeds, and thus smaller final particle size. Figure 5.7 - TEM images of (a,b) Eu(PW11)2@SiO2 NPs prepared using 95 mg (16 µmol) of corresponding europium polyoxometalate.; (c) Size distribution histogram of the mentioned EuPW11@SiO2 NPs. For direct comparison size distribution the histogram of EuPW11@SiO2 (d) NPs prepared using 8 µmol of the same europium polyoxometalate is also presented. 5.2.2.1. Scanning Electron Microscopy Elemental mapping of NPs by scanning electron microscopy-energy dispersive X- ray spectrometry (SEM-EDS) was performed to evaluate the distribution of the encapsulated POMs into the NPs. Figure 5.8 and Figure 5.9 present the SEM-EDS mapping images of EuPW11@SiO2 and Eu(PW11)2@SiO2 respectively. The mapping identified the presence of tungsten (W) and silicon (Si) through the NPs. Although it was not possible to identify the lanthanide ion Eu3+ by EDS (Figure 5.9 b), the results show that the W (one of the major elements that constitute POMs structure) is well distributed in the samples and that it is surrounded by Si. FCUP Europium polyoxometalates encapsulated into silica nanoparticles 147 Figure 5.8 - (a) STEM image of EuPW11@SiO2 NPs; (b) overlapping of EDS mapping for Si (red) and W (green), (c, d) separated EDX mapping for Si and W respectively Figure 5.9 - (a) STEM image of Eu(PW11)2@SiO2 NPs; (b) EDS spectra of Eu(PW11)2@SiO2, (c, d) separated EDS mapping for Si and W respectively. The Copper (Cu), aluminium (Al) and tin (Sn) peaks come from the support grid. 148 FCUP Europium polyoxometalates encapsulated into silica nanoparticles 5.2.2.2. Characterization by vibrational spectroscopy Spectroscopic methods including FT-IR, FT-Raman and solid state 31P NMR were used to characterize the nanoparticles and to analyse the integrity of the incorporated Eu(PW11)x. The FT-IR spectra of these nanomaterials as well as those from the corresponding europiumpolyoxometalates 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. FCUP Europium polyoxometalates encapsulated into silica nanoparticles 149 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@GPTEsSiO2 Wavenumber (cm-1) Eu(PW11)2 Eu(PW11)2@SiO2 Eu(PW11)2@CPTEsSiO2 B A 150 FCUP 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 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 silicacoated 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@SiO2 Eu(PW11)2@GPTEsSiO2 Eu(PW11)2@CPTEsSiO2 A B