Research Collection
Doctoral Thesis
Nucleic acid/inorganic particle hybrid systems from design to application
Author(s): Puddu, Michela
Publication Date: 2015
Permanent Link: https://doi.org/10.3929/ethz-a-010609177
Rights / License: In Copyright - Non-Commercial Use Permitted
This page was generated automatically upon download from the ETH Zurich Research Collection. For more information please consult the Terms of use.
ETH Library DISS. ETH No. 23177
NUCLEIC ACID/INORGANIC PARTICLE HYBRID SYSTEMS:
FROM DESIGN TO APPLICATION
A thesis submitted to attain the degree of
DOCTOR OF SCIENCES of ETH ZURICH
(Dr. sc. ETH Zurich)
presented by
MICHELA PUDDU
MSc Materials Science
born on 02.03.1987
citizen of Italy
accepted on the recommendation of
Prof. Dr. Wendelin J. Stark, examiner Prof. Dr. János Vörös, co-examiner Dr. Robert N. Grass, co-examiner
2015 2
3
To my parents
“Considerate la vostra semenza: fatti non foste a viver come bruti, ma per seguir virtute e canoscenza."
Dante Alighieri, Divina Commedia, Inferno, canto XXVI, vv. 18-20. 4
5
Acknowledgments The PhD has been a wonderful and exciting experience, and I want to take a moment to thank the many people who took part in this journey.
Foremost, I would like to express my gratitude to Prof. Wendelin Stark for giving me the opportunity to do research and develop a mature scientific approach in a highly stimulating and innovative environment. I am indebted to him for providing me with entrepreneurial spirit and vision that influenced my thinking and career choice. I also thank him for the precious lesson that a balance between research interests and personal pursuits does not keep away success, but it actually brings one step closer to it.
My most sincere thanks go to my advisor, Dr. Robert Grass, who oriented and supported me with enthusiasms and promptness through the wonders and frustrations of scientific research. He has been a great inspiration for me, a dedicated teacher and mentor, taking care of both my academic, professional and personal development. It has been a real pleasure to work under his supervision, with heaps of fun and excitement, and the imparted knowledge will be a great help throughout my career and life.
I kindly acknowledge Prof. János Vörös for accepting to co-examine my dissertation, concluding together the path started few years ago from his lab, where he firstly welcomed me at ETH for my master thesis project, opening up a range of new opportunities for me.
I want to express my gratitude to Prof. Marcy Zenobi-Wong for hosting me in her lab whenever needed, and for the fruitful scientific discussions. Furthermore, I would like to thank Frank Krumeich, for the high quality transmission electron microscopy and energy- dispersive X-ray spectroscopy analysis, and Hanspeter Hächler for the magnetic hysteresis measurements.
Many thanks go to Vladimir Zlateski for the great support and the enjoyable time we had together within and outside the lab, in Zurich and around the world. I am grateful to Daniela Paunescu for her contribution both as co-worker and friend. I thank Carlos Mora for the stimulating discussions and exchange of ideas (scientific and non). My most recent work and achievements have benefited from the suggestions and aid of Gediminas Mikutis, who is sincerely acknowledged. I thank Dirk, my office mate, neighbour, eventually trainer, for helping and entertaining me in the lab and outside. I would also like to extend my gratitude to all the other current and former member of this research group for their help and for the cheerful atmosphere: Mirjam, Mario, Elia, Samuel Hess, Philipp, Corinne, Antoine, Lukas, 6
Michael Loepfe, Tino, Christoph Kellenberger, Renzo, Jonas, and Nora. It was fun and pleasant to work and spend time with them all.
I want to thank Dr. Neil D. Telling for his support and instructive interaction during my secondment at Keele University. I am grateful to all the “Keele gang” members for their support and for providing a pleasant working atmosphere: Dalibor Soukup, Kaarjel Narayanasamy, Antonella Lisella, and Eva Luther. I thank Rebmann Balder from Freiburg University, with whom I had a productive collaboration and fun time.
I would also like to acknowledge my friend Christopher Millan, always available to share ideas and help. My sincere appreciation is extended to my German teacher and friend Vital Lutz, who made the learning of this complicated language a pleasant and entertaining activity.
I am most grateful to my beloved parents for their unconditional faith in me and for letting me to be as ambitious as I want (under their watchful eye). The love, help and opportunities they have given me over the years have been essential to me, and their ability to tackle challenges has always been exemplary and inspiring. A special thank goes to my partner Vincent Dabir for his constant patience and support in my work as well in life.
This work has been financially supported from the EU-ITN network Mag(net)icFun (PITN- GA-2012-290248), which is kindly acknowledged.
7
Table of contents
Acknowledgments 5
Zusammenfassung 11
Summary 13
1 Nucleic acid nanotechnology: the past, the present and the future 15 1.1 The nucleic acid era 16 1.2 The arise of nucleic acid nanotechnology 17 1.3 Nucleic acid interaction with inorganic particles 20 1.4 Nucleic acid as structural elements 22 1.4.1 Hybridization reaction-directed assembly of inorganic particles 22 1.4.2 Nucleic acid-templated fabrication of inorganic particles 23 1.5 Nucleic acid as functional elements 25 1.5.1 Information storage 25 1.5.2 Nucleic acid-delivery systems 27 1.5.3 Sensing 30
2 Magnetically recoverable, thermostable, hydrophobic DNA/silica encapsulates and their application as invisible oil tags 33 2.1 Introduction 34 2.2 Experimental section 36 2.2.1 Particle synthesis 36 2.2.2 Particle characterization 37 2.2.3 DNA recovery 37 2.2.4 qPCR standard curves 38 2.2.5 DNA absolute quantification 38 2.2.6 Thermal stability 39 2.2.7 Sanger sequencing 39 2.3 Results and discussion 39 2.4 Conclusion 49
3 Silica microcapsules for long-term, robust and reliable room temperature RNA preservation 51 3.1 Introduction 52 3.2 Experimental section 54 3.2.1 Handling RNA 54 3.2.2 RNA sources 54
3.2.3 SiO2/RNA microcapsule synthesis 54 3.2.4 Microcapsule characterization 55 8
3.2.5 RNA recovery 55 3.2.6 Gel electrophoresis 55 3.2.7 One-step RT-qPCR 55 3.2.8 RNA absolute quantification 56 3.2.9 Reactive oxygen species treatment 56 3.2.10 RNase treatment 56 3.2.11 Long term RNA stability 56 3.2.12 Capillary electrophoresis 57 3.2.13 Sanger sequencing 57 3.3 Results and discussion 57 3.4 Conclusion 65
4 Magnetically deliverable calcium phosphate nanoparticles for localized gene expression 67 4.1 Introduction 68 4.2 Experimental section 69 4.2.1 Nanoparticle production 69 4.2.2 Nanoparticle characterization 70 4.2.3 Plasmid preparation 70 4.2.4 DNA binding assay 71 4.2.5 Cell culture 71 4.2.6 Transfection of mammalian cells 71 4.2.7 Localization of gene expression 73 4.2.8 Live/dead assay 73 4.2.9 Microscopy 73 4.2.10 Cell counting 73 4.2.11 Statistical analysis 74 4.3 Results and discussion 74 4.4 Conclusion 82
5 Submicrometer-sized thermometer particles exploiting selective nucleic acid stability 83 5.1 Introduction 84 5.2 Experimental section 85 5.2.1 Nucleic acids 85 5.2.2 Iron oxide particle synthesis 86 5.2.3 Encapsulate synthesis 86 5.2.4 Nucleic acid recovery 86 5.2.5 Encapsulate characterization 87 5.2.6 Reactive oxygen species treatment 87 9
5.2.7 qPCR and RT-qPCR analysis 87 5.2.8 Nucleic acid absolute quantification 88 5.2.9 Temperature tests 88 5.2.10 Kinetic studies 88 5.2.11 Calibration curves 89 5.3 Results and discussion 89 5.4 Conclusion 94
6 Conclusion and outlook 95
Appendix 99 A.1 Supporting information to chapter 2 100 A.1.1 Particle density 100 A.1.2 Particle sedimentation velocity 100 A.1.3 Particle aggregation time 101 A.1.4 Independent two-sample t-test 101 A.2 Supporting information to chapter 3 106 A.2.1 Kinetics 106 A.2.2 Simplified procedure for RT-qPCR 107 A.3 Supporting information to chapter 4 110 A.3.1 Particle density 110 A.3.2 Primary particle size 110 A.3.3 Particle size dispersity 110 A.3.4 Number of particles per aggregate 110 A.3.5 Particle long-term storage 111 A.4 Supporting information to chapter 5 115
References 119
10
11
Zusammenfassung Hybridsysteme aus Nukleinsäuren und anorganischen Partikel kombinieren die einzigartigen chemischen und physikalischen Eigenschaften der anorganischen Partikel mit der Hybridisierungs- und Codierungsfähigkeit von Nukleinsäuren. Diese Ausgangsmaterialien sind vielversprechend für die Entwicklung von neuartigen und intelligenten Materialien für biologische und nichtbiologische Anwendungen. Die vorliegende Doktorarbeit setzt sich mit den Fortschritten in der Entwicklung und Anwendung von Nukleinsäure/anorganischen Partikel-Hybriden in verschiedenen Disziplinen auseinander.
Kapitel 1 fasst die Auswirkungen der Nukleinsäure-Forschung auf unsere Gesellschaft unter der Betrachtung verschiedener Aspekte zusammen. Es wird ein Überblick über den Wissensstand im Bereich der Manipulation von Nukleinsäuren zur Herstellung von supramolekularer Strukturen aufgezeigt. Zudem wird erklärt wie die Nukleinsäure- Technologie zusammen mit der Partikel-Technologie verwendet werden kann, um die Möglichkeiten der Nukleinsäuren in verschiedenen Bereichen, wie beispielsweise dem "Bottom-up" Aufbau von anorganischen Partikeln, in der Informationsüberlieferung und – speicherung und in der Sensorik, aufzuzeigen.
In Kapitel 2 wird die Verwendung von Nukleinsäure/Partikel-Hybriden als Fälschungsschutz- Marker zur Kennzeichnung von Flüssigkeiten und Produkten beschrieben. Diese magnetisch wiedergewinnbaren und hydrophoben Desoxyribonukleinsäure (DNA)/Siliziumdioxid- Partikel-Hybride wurden für die Entwicklung einer kostengünstigen Plattform zur Markierung von Ölen und Ölprodukten synthetisiert. Die Marker wurden diesen Produkten, welche bewiesenermassen oft gefälscht werden, bei sehr tiefen Konzentration (ppb = 1 μg Marker pro Liter Flüssigkeit) hinzugegeben und anschliessend durch magnetische Separation zurückgewonnen. Im Anschluss wurden die Partikel aufgelöst und mittels real-time Polymerase-Kettenreaktion analysiert. Das Verfahren wurde für die Kennzeichnung eines Kraftstoffes (Benzin), eines kosmetischen Öles (Bergamotte-Öl) und eines Lebensmittel-Öles (Olivenöl) getestet. Die Echtheit der gekennzeichneten Artikel wurde nachgewiesen und verdünnte Produkte sind erkannt worden.
In Kapitel 3 werden Siliziumdioxid- Partikel als praktische und zuverlässige Lagerlösung für Ribonukleinsäure (RNA) Proben bei Raumtemperatur vorgestellt, die somit den Gebrauch von Gefrierschränken für die Lagerung dieser Moleküle erübrigen. RNA konnte innerhalb der Siliziumdioxid-Teilchen über einen längeren Zeitraum bei Raum- und erhöhter Temperatur sicher aufbewahrt werden. Das Verfahren erwies sich als kompatibel mit gängigen RNA 12
Analysemethoden. Es vereinfacht den Umgang mit RNA Proben sowie deren Lagerung und Transport, und verhindert die kostenaufwändigen Lieferbedingungen und Raumprobleme von Gefriergeräten.
Kapitel 4 beschreibt die Verwendung von Eisenoxid markierten Trikalziumphosphat Partikeln als Transfektionvektoren. In diesem Materialverbund erleichtert das Kalziumphosphat den Eintritt der DNA in die Zellen ohne die Anwendung von toxischen kationischen Mediatoren, während das Eisenoxid eine magnetische Partikel-Dirigierung zu einem Zielort ermöglicht. Das Material wurde mittels Flammenspraysynthese hergestellt, da dieses Verfahren eine einfache, kostengünstige und reproduzierbare Synthese von Nanopartikeln ermöglicht. Die produzierten Nanopartikel, welche in Anwesenheit von Kalziumchlorid DNA binden, ermöglichten die effiziente Transfektion von humanen embryonalen Nierenzellen 293. Durch die Anwendung eines magnetischen Feldgradienten konnten die Partikel an bestimmten Orten in einer Zellkulturplatte konzentriert werden, um eine lokalisierte Transfektion zu erzielen.
In Kapitel 5 wird ein neuer Ansatz zur Thermometrie mit einem Hybridsensor aus Nukleinsäure und Siliziumdioxid vorgestellt. Der Sensor besteht aus Nukleinsäuremolekülen unterschiedlicher thermischer Stabilität, einer DNA- und einer RNA-Sequenz, die innerhalb von Siliziumdioxidpartikeln oder magnetischen Siliziumdioxidpartikeln eingeschlossen sind. Der Sensormechanismus basiert auf dem selektiven und kontrollierten Zerfall einer der beiden eingeschlossen Nukleinsäure-Sequenzen. Mit Hilfe der Quantifizierung der Nukleinsäure- Konzentrationen durch der real-time Polymerase-Kettenreaktion, und durch das Wissen der differenziellen Zerfallskinetik der zwei Nukleinsäure-Sequenzen, kann mit diesen Partikeln eine akkumulierte Temperaturhistorie ermittelt werden. Die hohe thermische und chemische Stabilität und die Fähigkeit die Temperaturinformationen lange Zeit zu speichern, ist ein entscheidender Vorteil im Vergleich zu aktuellen Mikrothermometern.
Kapitel 6 beinhaltet allgemeine Schlussfolgerungen der untersuchten Hybridsysteme und gibt einen Ausblick auf mögliche Anwendungen und die Entwicklung der hier vorgestellten Materialien.
13
Summary Nucleic acid/inorganic particle hybrid systems combine the unique chemical and physical properties of inorganic particles with the information encoding and recognition properties of nucleic acids. These materials hold great promise for the development of novel advanced, smart materials for both biological and non-biological use. In the present thesis recent advances in the design and application of nucleic acid/inorganic particle hybrids in different disciplines are reported.
Chapter 1 summarises the impact of nucleic acid research on various aspects of our society. It reviews the state of the art of nucleic acid manipulation to create supramolecular structures, and examines how nucleic acid technology is merged with particle technology, allowing the properties of nucleic acids to be exploited in various fields, including but not limited to the “bottom-up” assembly of inorganic particles, information delivery and storage, and sensing.
Chapter 2 introduces the use of nucleic acid/particle hybrids as anti-counterfeit tags to label bulk fluids and products. Magnetically recoverable, hydrophobic deoxyribonucleic acid (DNA)/silica particle hybrids were engineered and utilized to develop a low-cost platform for the tracing/tagging of oils and oil-derived products, which are often adulterated. To act as tags, the particles are dispersed in the product at parts per billion levels (1 ppb = 1 μg taggant per liter of fluid) and further retrieved by magnetic separation, followed by particles dissolution and DNA analysis by real-time polymerase chain reaction. The procedure was tested for the labeling of a fuel (gasoline), a cosmetic oil (bergamot oil), and a food grade oil (extra virgin olive oil), being able to verify the authenticity of tagged products and discriminate diluted items.
In Chapter 3, silica particles are presented as practical and reliable room temperature storage media for ribonucleic acid (RNA) samples as opposed to freezing strategies. RNA could be safely preserved within the silica particles for prolonged periods of time at ambient and elevated temperatures, and showed compatibility with standard downstream analysis. The method greatly facilitates the handling, storage and transport of RNA samples, avoiding the expensive shipments and the problems of space presented by freezers.
Chapter 4 shows the use of iron oxide/tricalcium phosphate composite particles produced by flame spray technology as transfection vehicles. While calcium phosphate facilitates DNA entry into cells without the need for toxic cationic mediators, magnetic iron oxide allows for particle localization at a target site. Additionally, flame spray synthesis ensures easy and low- 14 cost nanoparticle production in a reproducible way as opposed to standard calcium phosphate precipitation. The nanoparticles, exhibiting a DNA-binding capacity in the presence of calcium chloride, allowed to efficiently transfect human embryonic kidney 293 cells, and could be concentrated at specific sites in a cell culture plate through the application of a magnetic field gradient to achieve localized transfection.
In Chapter 5 a novel approach to thermometry employing a nucleic acid/silica hybrid sensor is introduced. The sensor is composed of nucleic acid molecules of different thermal stability (i.e. a DNA sequence and a RNA sequence), which are encapsulated within sub-micrometer sized silica particles or, alternatively, magnetic silica particles to achieve easier sensor recovery. The sensing mechanism relies on the selective and controlled encapsulated nucleic acid sequence degradation. Temperature read-out is obtained by quantifying the differential nucleic acid damage by real-time polymerase chain reaction. Thanks to the high thermal and chemical stability, and the capability of storing the read temperature over time, the sensor overcomes limitations of current small scale thermometers.
To conclude, Chapter 6 includes general considerations on the investigated hybrid systems and gives an outlook on other potential applications and further development of the presented materials.
15
1 Nucleic acid nanotechnology: the past, the present and the future
16
1.1 The nucleic acid era
Based on the current biological knowledge, it is difficult to imagine how poor the understanding of the molecular basis of life was 60 years ago. Although the contribution of many scientists brought to the present stage, a paper which appeared in Nature in 1953 can be legitimately considered as a turning point. Authored by James Watson and Francis Crick, the article reported the discovery of the structure of deoxyribonucleic acid (DNA), the molecule storing the genetic information required to generate and maintain life.1
“The double helix is indeed a remarkable molecule. Modern man is perhaps 50,000 years old, civilization has existed for scarcely 10,000 years and the United States for only just over 200 years; but DNA and RNA have been around for at least several billion years. All that time the double helix has been there, and active, and yet we are the first creatures on Earth to become aware of its existence.”
Francis Crick (1916-2004)
Their model, explaining how DNA replicates and how genetic information is encoded on it, marks the beginning of a new era of rapid progresses in biology, which continues until today. Within few decades, a detailed understanding of cellular processes has been reached and many technical advances have been achieved. Between these, it is worth mentioning the development of DNA sequencing, the process determining the nucleotide order within a DNA molecule, and the invention of the polymerase chain reaction (PCR) technique, which enables to produce millions of copies of a DNA sequence in approximately one hour, starting from one or a few DNA molecules. From the first rudimental techniques, sequencing methods have rapidly evolved, bringing to the development of automated processes and, more recently, to next-generation technologies, which provide unprecedented speed and throughput. Similarly, the PCR has established as the standard technique for gene analysis, routinely used not only in research, but also in medical diagnostics and forensics.
Since 1953, the DNA helical structure has been largely exploited and manipulated in various fields of science and technology, medicine, food, agriculture, data storage and even art (Figure 1.1). Biomedicine has evolved into a global industry, and DNA has become a cultural icon used from commercial and governmental enterprises to talk about identity, spirituality, social relations, progress and stability.2 Historically, this is an exceptional phenomenon since no other molecule ever reached this status.3 17
Figure 1.1 Overview of how DNA revolutionized medicine,4 evolution and environmental biology,5 agriculture (golden rice photo source: www.flickr.com, author: IRRI, date: February 2015),6 forensics, art,7 and many other important fields of our society.
Similar was the fate of ribonucleic acid (RNA), the other nucleic acid molecule carrying genetic information, even though advances in the field were delayed with respect to the DNA case. For ages RNA has been in the shadow of DNA and proteins: although scientists knew that RNA was involved in several cellular processes, it was thought to have a mere supportive role. It was only when previously unknown forms of RNA with surprising functions were discovered in the late 20th century, that the “RNA revolution” started. Since then, the research and the financial interest in the field has grown enormously, the various RNAs being used to develop new experimental medications against cancer and infectious or chronic diseases, which are soon expected to perform better than many available drugs.8
1.2 The arise of nucleic acid nanotechnology
Apart from the undiscussed centrality of nucleic acids in molecular biology, the field which mostly benefits from nucleic acids is surely nanotechnology,9-12 a busy research area. The same properties that make nucleic acids successful as genetic material, make them appealing in nanotechnology. Because of the nanoscale size and the special molecular structure, the 18 nucleic acids are in fact convenient and logical building blocks for constructing new nanomaterials.
Nucleic acids consist of a linear sequence of nucleotides, linked together by phosphodiester bonds. Each nucleotide is composed of a nitrogen-containing base - either cytosine (C), guanine (G), adenine (A), thymine (T, present only in DNA), and uracil (U, present only in RNA) -, a 5-carbon sugar (deoxyribose in DNA and ribose in RNA), and a phosphate group. The order of bases along a nucleic acid strand encodes the genetic information. While DNA is composed of two strands, hold together by hydrogen bonds between base pairs (A with T, and C with G), RNA is more often found in nature as a single-stranded molecule, and intramolecular base pairing (G with C and A with U) can occur. Such structure brings unique features, such as the ability to hybridize (pairing of complementary strands) and to carry information, and the possibility to combine nucleic acids with other functional molecules or nanostructures. Additionally, nucleic acid use in nanotechnology is now facilitated by the availability of simple and well established nucleic acid synthesis and detection techniques.
Nucleic acids have been used to build a number of nanoscale structures and devices, either alone or in combination with other materials. Within such structures the nucleic acid molecules can have different roles, which can be basically divided in two classes: structural and functional. In most of the fully nucleic acid-based constructions, the molecules are used as structural units to produce specific geometrical shapes and objects. This field is better known as structural nucleic acid technology,9 and it is very well established, starting 30 years ago with the pioneering work of Seeman,10, 13-15 who firstly described the idea of using DNA as construction material. For this purpose, DNA is indeed particularly appropriate due to its structural and chemical stability. The prominent construction strategy is the use of “sticky ends”, short single-stranded extensions protruding from a double-stranded DNA molecule.10 Two complementary sticky ends spontaneously combine through complementary base pairing to form a duplex. This predictable and highly specific self-assembly has been used to construct a variety of two- and three-dimensional structures, such as lattices and crystals,14-19 reaching is some cases macroscopic dimensions.20 A particularly successful implementation has been DNA origami, in which a long scaffold strand is folded by hundreds of short auxiliary strands into a complex shape.21
Compared to DNA nanotechnology, RNA nanotechnology is an emerging field which has only recently started expanding, following the recognition of RNA in medicine.19, 22 Similarly to DNA, RNA can be manipulated to produce a variety of 2D and 3D structures.23-24 Although 19 more chemically labile than DNA, RNA displays a richer assortment of rigid loops and motifs, allowing for the construction of complex architectures.22-23
Although relevant applications have begun to emerge, the main aim of structural nucleic acid technology has so far been of purely controlling and reshaping the matter at a molecular level. More promising applications can be developed if a number of issues are addressed, the most significant being the high cost of nucleic acids and the error rate of self-assembly. These problems are partially circumvented when nucleic acids are combined with other micro- or nanoscale materials, such as inorganic particles, to form hybrid systems. In such systems, the nucleic acids can still serve as structural elements, to direct the “bottom-up” assembly of inorganic bricks, or alternatively, the molecules can take an active role (Figure 1.2).
Figure 1.2 Nucleic acid-inorganic particle hybrid units for the bottom-up construction of complex architectures, for sensing purposes, or for the delivery/storage of biological and non-biological information. While the structural complexity decreases from left to the right, the amount and the complexity of relevant information encoded in the nucleic acids increases.
Beside the powerful base-pair recognition property utilized to control the structure of materials, the nucleic acids possess in fact other special functionalities which, combined to the properties offered by particles, lead to the construction of more intelligent nanostructures and nanodevices. If the double helix is the core of structural nucleic acid nanotechnology, here the nucleic acid-nanoparticle hybrids are functional elements for storing information, curing (gene delivery), and sensing (Figure 1.2). 20
In the following paragraphs, some of the mentioned structural or functional nucleic acids- inorganic particle hybrids are described.
1.3 Nucleic acid interaction with inorganic particles
Because of their size, micro- and nanoparticles (NPs) possess per se unique properties, and therefore find application in a variety of areas. Coupling nucleic acids to particles yields remarkable results, allowing the design of new structures (Figure 1.3) for both biological and non-biological uses.
Figure 1.3 Varieties of nucleic acid-inorganic particle structures which can be obtained.
The conjugation of particles with nucleic acids can be achieved by direct covalent linkage or non-covalent interactions. Since synthetic oligomers with a large variety of functional end groups are commercially available, the coupling is often easily achieved.
Alkyne-modified DNA has been covalently coupled to azide-terminated gold25 and superparamagnetic nanoparticles26 through copper-catalyzed click chemistry. Oligonucleotides have also been covalently grafted onto silica and titania particle surface employing copper free alkyne-azide cycloaddition reaction.27 Alternatively, covalent coupling can be obtained by reacting amino-functionalized nucleic acids to nanoparticles with 21 carboxylic groups.28-29 Disulfide-coupling chemistry has also proven to be a simple and efficient method for the immobilization of oilgonocleotides onto the surface of inorganic beads.30-31 Furthermore, thiolated DNA oligonucleotides have been covalently attached to monomaleimido gold nanoparticles.32-33
A well-established route to couple DNA molecules to gold NPs uses strong Au-S bonds. To this aim, DNA molecules are synthesized with alkylthiol groups at the 3'- or 5'-ends34-36 or, alternatively, with several thiophosphate residues at their ends.37 The thiols spontaneously bind to a gold surface, allowing for DNA-Au particle coupling. As an alternative, the use of a cyclic disulfide linker38 or a multiple thiol-anchor39 for binding oligonucleotides to gold nanoparticles results in a more stable anchoring through two sulfur atoms.
An extensively used approach, based on a combination of various noncovalent interactions, employs highly specific receptor-ligand binding for the coupling of particles and DNA. For instance, biotinylated DNA can be easily immobilized onto streptavidin coated particles,28, 40- 42 or alternatively, DNA-streptavidin conjugates can be coupled to biotinylated nanoparticles.43
Nucleic acids are often attached to particles through noncovalent electrostatic interactions. DNA and RNA, being negatively charged, adsorb quickly onto positively charged nanoparticle surface. To this aim, nanoparticles are functionalized with positively charged groups or polymers.44-53 Electrostatic interactions also drive the adsorption of DNA on calcium phosphate (CaP) particles. There are in fact evidences of interaction between the Ca2+ ions in calcium phosphate and the negatively charged phosphate backbone of DNA. 54-56 It is considered that such electrostatic interactions are involved in the co-precipitation of calcium phosphate and DNA in a supersaturated solution, producing insoluble calcium phosphate- DNA composites.56 Similarly, it has been hypothesized that Ca2+ ions mediate the binding of DNA to other inorganic nanoparticles (aluminum, cerium, and zirconium oxide) in the 57 presence of CaCl2.
The adsorption of DNA on silica particles in the presence of chaotropic salts is based on electrostatic interactions, dehydration of DNA and silica surface, and hydrogen bond formation.58-59 The specifics of these interactions depend on the pH and on the salt concentration. The adsorption of nucleic acids on silica surfaces in the presence of chaotropic agents is regularly employed in the solid phase purification of nucleic acids using silica beads or gel. 22
Finally, an increasing number of scientists are investigating the adsorption of nucleic acids on clay particle surfaces, since it could have played a role in the creation of primitive life. The interaction of nucleic acids with clay minerals involves electrostatic forces, cation bridging, ligand exchange and hydrogen bonding,60-61 and is affected by the pH and the ionic strength.61 DNA has been also intercalated within synthetic layered double hydroxide clays (LDH), where the negatively charged backbone of the DNA interacts electrostatically with the positively charged LDH layers.62-63
1.4 Nucleic acid as structural elements
1.4.1 Hybridization reaction-directed assembly of inorganic particles
There is a growing interest in controlling the formation of organized particle patterns, driven by the need to further miniaturize device components and to exploit the properties of small scale materials, which are different from the same materials at the macro scale. Therefore, many methods have been developed for assembling particles into useful structures, including the use of nucleic acids acting as “smart glue” rationally organizing and holding components in place. Such hybrid systems exhibit different properties than those of both particles and nucleic acids alone.64
Mirkin and Alivisatos are the scientists who started to use DNA to organize particles32, 34 and who mostly contributed to the research field. Mirkin developed a method for assembling gold NPs, involving the use of two non-complementary DNA oligonucleotides which are immobilized onto the gold surface. The assembly of the DNA-functionalized gold particles proceeds by adding to the colloid solution an oligonucleotide with “sticky ends”, which are complementary to the immobilized sequences, and can be reversed by thermal denaturation.34 Concurrently, Alivisatos and co-workers developed a similar technique where the DNA is used to control the relative spatial arrangement of Au nanocrystals. There gold particles individually functionalized with a single DNA oligonucleotide strand are assembled into dimers and trimers by adding a complementary DNA template sticking particles together.32, 65
The use of a complementary DNA linker has also allowed for the generation of networks made of Au particles of different size.66 In a similar approach, oligonucleotide-functionalized CdSe/ZnS quantum dots (QDs) were assembled to form QD nanoparticle structures.67 23
A different self-assembly strategy relies on the use of DNA capped gold clusters, which assemble in the presence of a complementary carrier RNA, used as a template to bind and 43 organize the particles in one dimension.
The potential of the bottom-up assembly of nucleic acid-inorganic particle components has also been exploited for the formation of complex structures, such as 2D arrays of Au particles,68-69 tubular structures carrying Au nanoparticles,70 pyramids of DNA with gold nanocrystals at each vortex,71 and different crystallographic arrangements diffracting X- rays.72-73
New, interesting and useful hybrid systems have been generated through such studies. In comparison to nucleic acid architectures, these hybrid structures show greater potential in many practical applications, and the smaller amount of required nucleic acid material facilitates their implementation.
1.4.2 Nucleic acid-templated fabrication of inorganic particles
Nucleic acids-templated synthesis of nanoparticles holds enormous potential for the rational design of nanoparticles. It is recognized that nanoparticle size, morphology and physico- chemical properties can be finely tuned by modulating template nucleic acid conformation and sequence.74
Silver nanoparticles have been produced and positioned in order to form a nanowire structure through DNA metallization. The process is based on the localization of silver ions along the DNA molecule through electrostatic interactions with the negatively charged backbone of the molecule or formation of complexes between the silver and the DNA bases.75-76 The complexes are then reduced to form separated silver nanoparticles bound to the DNA skeleton. The formed nanoparticles serve as a catalytic surface for further reduction of silver, allowing a transition from separated nanoclusters into a continuous nanowire. 75 The same approach has been used to produce metallic nanowires of Pd, Pt, Au, and Cu.76 In a different study, oligomers adsorbed on the Au nanoparticles were utilized as template to mediate the formation of flower-like gold nanoparticles. 77 RNA molecules have also been used to mediate the synthesis of metallic nanoparticles and control their shape.78-79
The size and the photoluminescent properties of quantum dots have been controlled by modulating the structure of the templating nucleic acid molecules. Coffer and co-workers were the first ones to utilize DNA as a stabilizer/template for the aqueous synthesis of CdS 24 nanocrystallites from Cd2+ and S2-.80 Furthermore, they have used a circular plasmid DNA molecule anchored to a substrate to form an array of semiconductor nanoparticles matching the shape of the molecule.81 In a similar approach, transfer RNA has been used as a template to direct the synthesis in solution of highly stable CdS nanocrystals, whose structure could be modulated by altering the RNA conformation.82
Calcium phosphate nanoparticles with embedded DNA have been produced and are regularly used in cell transfection. The more popular method to obtain them is the in situ precipitation of calcium phosphate in the presence of DNA.83-84 Gerdon and co-workers demonstrated the ability of DNA to template calcium phosphate mineralization.85 According to them, DNA would act by sequestering calcium and phosphate ions, thereby supersaturating the microenvironment and acting as a nucleation site where the mineral can form.
DNA directed silica growth has been also reported, allowing the formation of 1D, 2D and 3D structures.86 Because of the electrostatic repulsion between negatively charged DNA and silicate, the synthesis requires the “conversion” of DNA into a positively charged molecule. This has been achieved for the first time in 2004 using cationic molecules carrying one guanidinium and one ammonium group as bridge between negatively charged DNA and tetraethyl orthosilicate (TEOS). While the guanidinium group interacts preferentially with DNA phosphate groups, the ammonium group gives a net positive charged to the template molecules, enabling the interaction with anionic TEOS, which condense along the DNA backbone forming DNA-silica hybrid nanotubes.87 Other cationic molecules have been used for the same purpose, such as N-trimethoxysilylpropyl-N,N,N-trimethylammonium chloride (TMAPS), bringing a positively charged quaternary ammonium group able to interact with DNA, and a silane site able to co-condense with TEOS. Using TMAPS as cooperative species, 2D DNA-silica structures88 have been obtained, while 3D impeller-like helical DNA- silica complexes have been synthesized with the same procedure in the presence of magnesium ions.89
Despite tremendous progress, a comprehensive understanding of some underlying mechanisms of nucleic acid templated particle synthesis is still lacking and therefore predicting the resulting structures is not always possible.74 25
1.5 Nucleic acid as functional elements
1.5.1 Information storage
The protection and storage of nucleic acids is of importance in a wide range of fields and disciplines. However, it must be noticed that the perspectives vary considerably with the purposes of the preservation or the storage.
When nucleic acids are delivered into living cells for research or therapeutic purposes, it is fundamental to preserve their integrity so that they can accomplish their function. This is also often the case when nucleic acids are used as sensing elements. Under these circumstances, nucleic acids preservation is required for an amount of time that covers at least the delivery or the sensing event (generally minutes up to days).
Due to the continuous improvement of genomic analysis methods, it is becoming more and more important to sample and store nucleic acids in a long-term stable manner for further investigations using the newly developed techniques. This activity has strategic importance for clarifying unsolved biological and medical problems. While DNA required for medical testing has to be stable for few years, samples used in evolutionary biology have been preserved for thousands of years. Also in the field of forensics a high quality long-term storage of nucleic acids is of extreme importance.
Besides the storage of genetic information, nucleic can be used as carriers of non-genetic information. The use of DNA as an instrument to convey information was initiated by Davis with the a bio-artwork Microvenus (Figure 1.1),7 the first non-biological message encoded in DNA. Short specifications such as name, dates, batch numbers, have been encoded within DNA sequences and used as tracers to label substances. Also vast quantities of data have been encoded in the DNA structure, such as entire books, images and documents.90-92 In the modern internet era, huge amounts of data are being generated and there is a pressing need for more compact and long-lasting storage media. In this context, DNA digital data storage is becoming a hot topic. DNA is in fact recognized as an optimal "apocalypse-proof" storage system, due to its high data density, longevity (if properly stored), and resistance to obsolescence, as it is Nature’s information storage mechanism.
Besides the duration of the preservation, other parameters to consider are the environment and the conditions to which the nucleic acids are exposed to. When nucleic acids have to be transported into cells, they have to be protected from the enzymes (deoxyribonucleases and 26 ribonucleases) present in the extra- and intracellular environment, which are the main reason for their degradation both in vitro and in vivo. Differently, the long term storage requires protection from humidity, oxygen and light causing nucleic acid decay. Specific applications in sensing or tracing can require additional protection against heat or more energetic radiations.
Protection against enzymatic degradation can be easily achieved by DNA complexation with positively charged particles.44-49, 93-96 A few possible explanations have been proposed for the DNA protection based on positively charged nanoparticles. One is that the positive charge of the nanoparticles keeps Mg2+ away by electrostatic repulsion. Since Mg2+ ions are needed for the enzymatic cleavage, the reaction is inhibited. A second one is that DNA binding onto the NP surface results in a variation of the DNA conformation, protecting DNA from cleavage.47, 96 Since amino-modified glass slides do not provide the same protection as the NPs, it has been proposed that the size of the NPs may force the DNA to become bound in such a way that cleavage is either impossible or at least slower on the NP surface.47 Moreover, it has been shown that DNA bound to cationic gold particles is not only stable toward enzymatic digestion, but also physical sonication, while no protection is reported against chemically induced radicals.95
Au NPs functionalized with a dense monolayer of oligonucleotides have also showed resistance to enzymatic degradation. The high DNA density results in high local salt concentration, which could lead to inhibition of nuclease degradation.97 Furthermore, the dense packing of the oligonucleotides on the NP surface likely causes steric enzyme inhibition.98
If DNA-CaP co-precipitates provide only partial protection against enzymatic digestion, improved encapsulation techniques have led to complete protection from external DNase.99 Moreover, outer silica100 or CaP101-102 have been often applied to nucleic acid/nanoparticle systems in order to achieve enhanced resistance against enzymatic degradation.
While numerous approaches have been developed to protect naked nucleic acids from enzymes, only few strategies are available to circumvent exposure to chemicals, radiations and high temperatures, which represent a major threat to nucleic acid preservation.
It has been showed that both DNA103 and RNA104 adsorbed on clay particles are protected from enzymatic degradation,104 heavy metals induced lesions,61 and UV radiation damage.105 Additionally, the binding of nucleic acid molecules on clay does not prevent their ability to 27 hybridize with complementary molecules or to function as templates for reverse transcription and replication.104 These findings could explain how nucleic acids survived in the hostile environment of the early Earth, characterized by challenging conditions for the molecules (high temperature, water, and UV radiation). It has been suggested that clay minerals could have acted as support capable of immobilizing and concentrating the organic material present in a primordial ocean, and protecting it from damage so that its biological potential could evolve and be expressed.104-106
Studies on clay minerals have led to the conclusions that in order to improve their protective effect, the intercalation of DNA into the clay layers could be performed.61 This goal has been achieved by Choy et al., who produced a DNA-inorganic hybrid by intercalating DNA into a synthetic layered double hydroxide structure. The hybrid was able to withstand severe biological, chemical, and thermal conditions. Additionally, the intercalation could be reversed and DNA recovered by acidifying the samples.63
Although the intercalation into layered materials improves nucleic acid protection as compared to surface adsorption, even better results could be achieved by creating a hermetic sealing from the external environment. We proved this concept by developing a method to reversibly encapsulate nucleic acids within silica particles, where both DNA and RNA are protected from heat, biological and chemical attack (chapter 2 and 3).50-53 By adding an additional titania layer, DNA protection against ultraviolet irradiation was ensured.107
1.5.2 Nucleic acid-delivery systems
Gene therapy holds the promise of treating a disorder without using drugs or surgery. This can be achieved by introducing nucleic acids able to induce gene expression (mostly plasmid DNA, pDNA, but also messenger RNA, mRNA) or knockdown (antisense oligonucleotides, microRNA, miRNA, or small interfering RNA, siRNA) into cells. The nucleic acids have to travel through the cell membrane and reach the appropriate cellular compartments in order to accomplish their function.
Naked nucleic acids do not successfully reach their destination themselves, but require the assistance of a suitable vector, which should be non-toxic and ensure that the nucleic acids survive the extra- and intracellular environment. Transduction is the process whereby foreign DNA is introduced into a cell via a viral vector. Transfection is instead the process of introducing nucleic acids into cells by non-viral methods. 28
Although antisense oligonucleotides have demonstrated their ability to silence genes while remaining bound to the particle surface,98 generally gene regulation involves the intracellular release of the delivered nucleic acids from the carrier particles. Therefore, an optimal carrier has to satisfy the requirements of adequate nucleic acids protection and efficient release.
Numerous inorganic particle-based systems, able to pack, transport, and deliver nucleic acids to various targets in the body have been developed.96, 101, 108 Due to their small size, particles are able to penetrate deep tissues and to interact with biomolecules on the cell surface or inside cells to achieve targeted delivery. Among a broad number of particle types, magnetic particles are particularly attractive because a magnetic field can be used to guide them to target cells and tissues, and to enhance the nucleic acid-particle complex cellular uptake,44, 49, 109 thereby decreasing required vector doses. Although magnetic particles have been combined also to viruses, they are mainly emerging as a safer alternative to viral vectors, which showed to be immunogenic.
In the nucleic acid-particle delivery pathway, particles are first adsorbed on the cell membrane and then are taken up by cells by endocytosis (Figure 1.4). Despite extensive studies on particle-based transfection, it is still not clear how nucleic acids are released after the uptake and if DNA reaches the nucleus alone or still bound to particles.101 It is possible that some particles are dissolved by the acidic environment in the endosomes, inducing endosome swelling and burst, and nucleic acid release into the cytoplasm.101, 110-111 For cationic particles it has been instead speculated that the pH-buffering capacity of protonable amino groups promotes endosome swelling and disruption (“proton sponge” effect), with a concomitant intracellular release.110, 112
Nucleic acid delivery with particles can be obtained in many ways. A common method to load nucleic acid onto nanoparticles is to functionalize them with positively charged polymers (usually, polyethylenimine, PEI) or groups able to form complexes with nucleic acids through electrostatic interactions. The positively charged species do not only protect the nucleic acids from enzymatic degradation, but additionally facilitate the interaction of the complexes with the negatively charged cell surface and their internalization. This approach has been widely used with gold colloids,45-46, 48 iron oxide,44, 49, 109 and silica nanoparticles (even in mesoporous form),47, 93-94, 113 as well as with QDs.114-116 In a similar approach, unfunctionalized iron oxide particles have been associated with nucleic acids in the presence of a cationic polymer mediator acting as a transfection enhancer.117-118 Despite their broad use, cationic species are recognized to be cytotoxic and therefore alternative strategies have been developed. 29
Figure 1.4 Schematic illustration of the transfer of particle-nucleic acid conjugates into cells: 1) particle adsorption onto the cell membrane, 2) uptake by endocytosis and 3a) escape from endosome via particle dissolution or 3b) other mechanism (e.g. “proton sponge” effect) with 4) nucleic acid release in the cytoplasm or transport to the nucleus (in the case of pDNA).
Oligonucleotides conjugated to Au NPs demonstrated to efficiently scavenge intracellular DNA or RNA by selective binding, inducing gene knockdown.98, 119 Interestingly, although negatively charged, the nucleic acid-gold NP systems showed high cellular uptake. It has been proposed that the nucleic acid-Au NP systems interact with scavenger receptors which mediate membrane transport.120
Nanostructured calcium phosphate has been shown to be better than other gene delivery methods in terms of immunogenicity and toxicity, although displaying relatively low transfection efficiency. Nucleic acid-CaP co-precipitates are by far the most popular calcium phosphate-based transfection method.83-84, 121 However, the co-precipitation technique does not allow to control the growth and size of the precipitates, which is of great importance.122 In order to produce monodisperse and stable CaP nanoparticles, more reproducible wet synthesis protocols have been developed by exerting a greater control over the mixing of the precursors solutions,123 or the precipitation conditions.124 Alternative synthetic routes have been also explored to obtain multi-shell calcium phosphate-DNA structures with a higher DNA loading.122 30
Flame-spray technology has been used as reproducible synthesis technique to form calcium phosphate, aluminum, cerium, and zirconium oxide nanoparticle, able to bind and deliver 57 DNA in the presence of CaCl2. A novel approach developed by us allows to achieve magnetically targeted gene delivery by using a cation-free, flame-sprayed magnetic calcium phosphate formulation, presented in chapter 4.
Lastly, nucleic acid intercalating layered double hydroxides have been evaluated as DNA delivery systems in various cell lines, obtaining good transfection efficiency.125-126
1.5.3 Sensing
DNA biosensors represent a new research field with interesting possibilities in various fields such as medical and environmental screening. Nowadays, DNA sensing is mostly related to genetic diagnostic or focused on the investigation of toxic effect on DNA, while few cases are reported on the use of DNA as a sensing molecule to detect other physical/chemical entities.
Most of the current DNA-nanoparticle hybrid sensors rely on hybridization. Hybridization tests for the detection of target nucleic acid sequences are routinely used for clinical and forensic analyses. These technologies rely on the immobilization of single-stranded DNA probes onto a substrate and the detection of the hybridization with the targets. The hybridized DNA is detected mainly using fluorescent labels, but presents some drawbacks in terms of cost and stability. As an alternative, nanoparticle-based labels offer several advantages, especially in terms of stability and availability of multiple transducing modes (optical, mechanical and electrochemical) and read-out options.
Au NPs have often been used as DNA hybridization tags. In early works of Mirkin, a colorimetric polynucleotide detection method based on using gold nanoparticle probes has been reported. There, the hybridization of the target with the complementary probes causes particle aggregation and a concomitant red to purple colour change.127-128 Such approaches have later been adapted to solid surfaces, with the advantage of allowing washing steps to remove excess reactants.129-130 Detection of the DNA hybridized on a solid substrate has been achieved optically by using oligonucleotide-modified gold nanoparticles probes.129 The use of colloidal gold nanoparticles has also allowed DNA hybridization detection by means of quartz crystal microbalance131-132 or by electrochemical means.133-134 Apart from gold particles, QDs are also strong competitors to fluorescent dyes in DNA hybridization due to their multicolour optical labelling capabilities, which have allowed for multiplexed assays.135 31
Besides nucleic acid sequence detection, aptamer-conjugated nanoparticles (dye-labeled particles or QDs) have been used for the detection of other biological entities, such as proteins42, 136 and cancer cells,28 by measuring an increase in fluorescence after target binding.
Au nanoparticles have been used for colorimetric metal ion detection. Mercury detection has been accomplished with high selectivity in the presence of other metal ions using oligonucleotide-gold nanoparticle probes and a linker oligonucleotide with a number of thymine-thymine mismatches. The oligonucleotides form DNA duplexes only in the presence of Hg2+ through thymine-Hg2+-thymine coordination chemistry, resulting in the formation of particle aggregates and concomitant colour change.137-138 Cu2+ detection and quantification has been carried out using a colorimetric method based on DNA functionalized Au NPs and copper catalysed click chemistry to induce aggregation.139
DNA hybridization on nanoparticle surfaces has also been used to measure temperature. In this process, polymer-coated magnetic NPs with DNA were let to hybridize with DNA carrying a fluorophore, and local temperature on the NP surface was determined measuring the fluorescence of the denatured DNA present in the supernatant.140
Besides DNA hybridization approaches, several sensors based on DNA damage have been developed. Most of them aim at studying the effect of drugs/agents on DNA and make use of solid supports and electrochemical methods. A few DNA damage-based sensors using nanoparticles have been reported. As an example, the sequence-length-dependent adsorption of single-stranded DNA on gold nanoparticles has been exploited for the colorimetric detection of DNA enzymatic cleavage and oxidative damage.141 A new approach to sensing with nanobiohybrids developed by us, based on nucleic acid damage quantification, has allowed the measurement of temperature and further storage of the measured data, introducing the “lab-on-a-particle” concept (chapter 5).
32
33
2 Magnetically recoverable, thermostable, hydrophobic DNA/silica encapsulates and their application as invisible oil tags
Published in parts as:
M. Puddu, D. Paunescu, W. J. Stark and R. N. Grass
ACS Nano 2014, 8, 2677-2685. 34
2.1 Introduction
Today adulteration extends to almost all types of products. Fuels, high value cosmetics oils and foodstuff (such us extra virgin olive oil), medicines and other chemicals are routinely counterfeited to produce illegitimate profit. Fuel stealing or illegal treading is a real problem in many countries, being responsible for huge money losses for governments.142-143 For this reason, several oil authentication programs have been promoted. Extra virgin olive oil adulteration with cheaper oil has become more frequent in recent years.144-145 Authenticity determination of olive oil product is fundamental for both trader interests and consumer health. Essential oils are expensive oils with a variety of cosmetic, therapeutic and culinary uses which are also often adulterated to increase profit.146 However, only pure products contain full range of components in the right ratios, imparting unique aroma, cosmetic and therapeutic properties that fake imitations cannot replicate.
Several tagging approaches have been proposed against adulteration, making use of imperceptible labels attached or integrated within products, and identifiable by specific instruments and procedures. Besides being invisible, a product tag should be inert, resistant, and harmless. Additionally, it should not affect product properties, should be cheap and easily detected. Tagging technologies include magnetic inks,147 fluorescent labels,148 photocromic and thermocromic inks,149 isotopic tracers,150 and Raman active components.151 polypeptides152 and nucleic acids153-155 have also been used as taggants. Above all DNA offers unique opportunities in this field, supported by mature DNA synthesis and analysis procedures.
Nucleic acids are Nature’s method to store and transmit instructions. Double helix conformation and stability allows DNA to accomplish this vital function. Since DNA related technologies were established, several artificial DNA structures for a broad range of applications have been designed.9, 11-12, 18, 156-160 The use of DNA codes has been also developed: DNA barcoding is now a well known strategy to identify biological species, as DNA fingerprinting is used to prove the identity of people (paternity test, forensics, victim recognition) and the biological origin of foodstuff.144 Instead of using natural occurring genomic sequences, artificial DNA with a unique sequence can also be introduced into goods to be marked.153-155 In this way identification of any branded product becomes possible.
The idea of writing, storing/carrying, and retrieving non-genetic information on DNA is powerful.51, 63, 90-91, 155, 161 However, incredible capabilities of DNA as Nature’s storage system 35 also arise from the ability of organisms to repair DNA when lesions occur. DNA stability per se is limited,162 since it can be damaged by metabolic processes as well as by several environmental factors. When DNA is used as information storage/carrier outside living organisms (an idea dating back about 20 years), there is lack of repairing processes and DNA can be severely damaged. To avoid its degradation and preserve its integrity DNA has to be preventively protected. While good protection against enzymatic degradation can be achieved by DNA complexation with positively charged molecules/particles,47, 113 or encapsulation in polymeric capsules,163 prevention of damage produced by radiation, temperature fluctuations, chemical (e.g. redox) exposure is more challenging. Several cold and dry storage strategies, eventually combined and improved by storage additives,164-165 are used to store/transport DNA. Alternative solutions, such as DNA encapsulation into porous silica matrix,166-167 or within layered metal hydroxide 63, 161 have also been proposed.
In previous studies51, 155 we have encapsulated single-stranded DNA (ssDNA) and double- stranded DNA (dsDNA) of various lengths in silica to produce radical-resistant and heat- resistant “synthetic fossils”. These silica materials could be dissolved with fluoride comprising buffers (i.e. buffered oxide etch, BOE) to recover intact DNA. This approach allows for a protection of DNA against chemical attack since it provides a physical barrier, which completely isolates DNA from the external environment, a situation very similar to that in natural fossils.
For the recovery of analytes from solution, magnetic separation methods are currently applied in different areas of analytical chemistry because of the advantages offered in comparison to similar non-magnetic techniques.168-170 Therein, magnetic separation of analytes allows for better sample handling and up-concentration and is suitable for use in a variety of automated analytical procedures. Fast and easy recovery of magnetic particles, as well as absence of sample volume limitations, results in their broad utilization in separation, identification and quantification of several chemical and biological species. For these reasons the use of magnetic biotechnology159-160 for tracing/tagging is very attractive. 154, 171
In the present article we present how the above described concepts of DNA analytics, magnetic separation and silica encapsulation can be combined to generate potent (read out at 1 ppb) and low-cost (0.02 ¢/L of product) tracers for the marking of oil based items.
We tested our tagging/tracing technology with three model oils: a fuel oil (gasoline), a cosmetic/therapeutic oil (bergamot oil, used in perfumery and cosmetics), and a food grade oil 36
(extra virgin olive oil), to scan a range of different oil types and present the applicability of the newly designed materials.
2.2 Experimental section
2.2.1 Particle synthesis
2+ 3+ 2+ 3+ Fe2O3 particles were synthetized by conventional co-precipitation of Fe / ions (Fe : Fe =
1:2) under alkaline conditions. 4 g FeCl24H2O (Aldrich, 99%) and 10.8 g FeCl36H2O (Aldrich, 98%) was dissolved in 50 mL distilled water. The solution was added dropwise to
500 mL of a 1 M NH4OH (prepared from Sigma Aldrich, 25%) solution under vigorous stirring at room temperature. The obtained particles were washed five times and stored in water at a concentration of ~50 mg/mL.
The prepared magnetic particles were functionalized with ammonium groups using N- trimethoxysilylpropyl-N,N,N-trimethylammonium chloride (TMAPS, 50% in MeOH, ABCR GmbH). 51, 155, 172 10 L of TMAPS were added to 1 mL of particle suspension in isopropanol (~50 mg/mL) and the mixture was stirred overnight (900 rpm) at room temperature. Particles were washed 3 times and stored in isopropanol at ~50 mg/mL.
The dsDNA (5’-ATT CAT GCG ACA GGG GTA AGA CCA TCA GTA GTA GGG ATA GTG CCA AAC CTC ACT CAC CAC TGC CAA TAA GGG GTC CTT ACC TGA AGA ATA AGT GTC AGC CAG TGT AAC CCG AT-3’, purchased by Microsynth AG) was adsorbed on the particles by mixing 1 mL of a 16 µg/mL DNA solution in water with 15 µL of particle suspension, followed by three washing cycles and redispersion in water (0.5 mL).
An additional layer of TMAPS was adsorbed onto the particle surface before growing SiO2 by using tetraethoxysilane (TEOS, ≥ 99%, Aldrich) as Si source. 51, 155 1 L of TMAPS was added to the particle dispersion, which was then vortexed before adding 1 L of TEOS. The mixture was stirred (900 rpm) for 4 h at room temperature and then additional 8 L of TEOS was added. The reaction was run for 4 days, stirred at 900 rpm.
C6 functionalization was achieved by conducting hydrolysis of n-hexyltrimethoxysilane according to the following procedure. Fe2O3/TMAPS/DNA/SiO2 particles were re-dispersed in ethanol (260 µL, ~4 mg/mL). 40 µL of NH4OH (Sigma Aldrich, 25%) was added and the dispersion was stirred (900 rpm) for 30 min at 40°C. A solution consisting of 100 µL of n- hexyltrimethoxysilane (TCI, > 96%) and 50 µL of ethanol was then added. The mixture was 37 stirred overnight at 40°C. The particles were washed three times and stored in toluene (100 µL, 10 mg/mL).
2.2.2 Particle characterization
The Fe2O3 nanoparticles were analyzed using a X'Pert Pro-MPD diffractometer (PANalytical). Diffraction data were acquired by exposing powder samples to Cu-Kα radiation (λ = 1.54060 Å), over a range of 10 - 70° 2θ with a step size of 0.05°. Iron oxide identification was carried out by means of the X'Pert software PANalytical High Score Plus.
Morphology of Fe2O3 and Fe2O3/TMAPS/DNA/SiO2 particles was observed using a transmission electron microscope (TEM, Philips CM12 FEI, W cathode, operated at 100 kV). Samples were prepared by placing dilute ethanol particle dispersion on a carbon foil supported by a copper grid (purchased from Plano GmbH).
Magnetic properties of Fe2O3 and Fe2O3/TMAPS/DNA/SiO2 were measured by vibrating sample magnetometer (Princeton Measurements Corporation, model 3900). Surface modification at each step was checked by Fourier transform infrared spectrometry (FT-IR spectrometer Tensor 27, Bruker Optics, equipped with a diffuse reflectance accessory, DiffuseIR™, Pike Technologies) on samples milled with KBr (2% w/w). Z-potential measurements were performed using a Zetasizer Nano (Malvern, Worcestershire, UK) with a laser beam wavelength of 633 nm. The nanoparticles were also analyzed with the elemental analyzer vario MICRO Cube (Elementar, Hanau, Germany) simultaneously for C, H, N.
Particle size distributions of Fe2O3 and Fe2O3/TMAPS/DNA/SiO2/C6 particles were obtained by an analytical photocentrifuge (dispersion analyser LUMiSizer, LUM GmbH, light source
470 nm). Fe2O3 particle dispersions in water were prepared for the analysis, as well as
Fe2O3/TMAPS/DNA/SiO2/C6 particle dispersions in toluene, gasoline, and limonene (R-(+)- Limonene, ≥ 96%, Fluka). Transmission profiles were recorded with time intervals of 20 seconds up to 8 hours, at a centrifuging speed of 3000 - 4000 rpm. Statistical data elaboration was performed by SEPView® software (LUM GmbH).
2.2.3 DNA recovery
Encapsulated dsDNA was recovered using a buffered oxide etch solution (NH4FHF/NH4F,
0.23 g NH4FHF + 0.19 g NH4F in 10 mL water) prepared and handled as described in 51 Paunescu et al. Since C6 functionalized particles are highly hydrophobic and the etching 38 solution is aqueous, more time was required for their complete dissolution than the time needed to dissolve the non-functionalized ones (few minutes).
2.2.4 qPCR standard curves
Standard curves were obtained from 10-fold dilution series (in water for unfunctionalized particles, in toluene, bergamot oil, gasoline, and extra virgin olive oil for particles bringing C6 modification) of the original particle dispersions. For comparison, both curves without and via magnetic separation were produced. Without magnetic separation: 300 µL of diluted buffered oxide etch (1:100) was added to 10 µL of particle dispersion at each dilution, to dissolve the particles. When toluene was the dispersant, the sample was dried at 45°C (Concentrator plus, Eppendorf AG) for 15 min before adding the etching solution. The solution was directly analyzed by real-time polymerase chain reaction (qPCR). Via magnetic separation: 1 mL of particle dispersion was placed in a magnetic separator and left to allow for complete particle separation. Olive oil suspension was diluted with toluene (1 mL oil + 1 mL toluene) to facilitate separation. Particles collected from oil suspensions were washed once with toluene. Again drying (3 min) was necessary to remove residual toluene. Then 300 µL of diluted buffered oxide etch was added as before and qPCR was performed. A standard qPCR protocol (Roche LightCycler® 96) was used, with the following primers sequences: 5’-ATT CAT GCG ACA GGG GTA AG-3’ (forward primer) and 5’-ATC GGG TTA CAC TGG CTG-3’ (reverse primer), purchased from Microsynth AG. Experiments with water and toluene particle suspensions were performed in triplicates (n = 3) and repeated twice, with similar results, to determine both qPCR analysis reproducibility and reproducibility of the experiments. In the case of the oils 5 samples for each taggant concentration considered were analyzed, each assayed by qPCR in hexaplicates (n = 30). Each qPCR reaction was performed with a Roche LightCycler 96, using the following protocol: 10 L of master mix (Roche, LightCycler 480 SYBR Green I Master Mix), 3 L of PCR grade water (Roche), 2 L primer mix (forward and reverse primers, 1 M each) and 5 L sample. Each reaction was assessed against a negative control, which contains water instead of sample DNA. Data are provided as mean cycle threshold CT value ± standard deviation.
2.2.5 DNA absolute quantification
Sample absolute quantification was accomplished using a standard curve (see Appendix, Figure A1.2), generated from DNA samples of known concentration (determined by Qubit 39 fluorometer using Qubit dsDNA HS assay, Invitrogen). Any unknown sample concentration was then determined by interpolation from this curve.
2.2.6 Thermal stability
For comparison, both encapsulated and free dsDNA were heat treated. Free DNA: 100 μl of dsDNA in water (0.08 μg/mL) was treated at 95°C for duration up to 24 h (2 samples per time point, each analyzed in triplicates by qPCR) and at various temperatures (100°C, 120°C) for 30 minutes (2 samples per temperature, each analyzed in duplicates). Protected DNA: 100 μl 6 of a 10 µg/L Fe2O3/TMAPS/DNA/SiO2 particle dispersion in water was heat treated at 95°C for periods of time up to 24 h (2 samples per time point, each analyzed in triplicates). Then 300 µL of diluted buffered oxide etch (1:100) was added to 10 µL of treated particle dispersion, and the obtained DNA solution was analyzed and quantified by qPCR. 100 μl of a 5 10 µg/L Fe2O3/TMAPS/DNA/SiO2-C6 particle dispersion in decalin (mixture of cis + trans, 98%, Sigma Aldrich) was heat treated at temperatures between 100°C and 160°C for 30 minutes (2 samples per temperature, each analyzed in duplicates). After treatment, the dispersion (100 µl) was diluted with toluene (900 µl) to reduce the viscosity, before separating the particles by magnetic means. The separated particles were dried for 3 min at 45°C and dissolved using 300 μl 1:100 BOE. The DNA solution was directly analyzed and quantified by qPCR. Data and error bars represent concentration mean values and standard deviations obtained.
2.2.7 Sanger sequencing
Particles were separated from 100 µL of 106 µg/L dispersion in decalin by magnetic means. After a wash with toluene, particles were dissolved in 100 µL buffered oxide etch. DNA solution was then purified (QIAquick PCR purification kit) and sequenced. DNA was sequenced in the direction 5’-3’ with the primer: 5’-CAG GGG TAA GAC CAT CAG-3’ (Microsynth AG).
2.3 Results and discussion
Iron oxide nanoparticles (see Appendix, Figure A1.1) were produced by a conventional co- precipitation method under ambient conditions. Since maghemite and magnetite XRD patterns are nearly identical, the iron oxide form obtained could not be identified by this method. 173 However, the aerobic synthesis conditions predicted formation of gamma-Fe2O3. The 40 nanocrystallite size, calculated from the XRD pattern (Figure 2.1a) by means of the Scherrer formula, was estimated to be 12 nm.
Figure 2.1 a) XRD pattern of Fe2O3 nanoparticles prepared by co-precipitation; b) hysteresis loops of Fe2O3 nanoparticles (solid line) and of Fe2O3/TMAPS/DNA/SiO2-C6 particles
(dashed line); c) TEM micrograph of Fe2O3/TMAPS/DNA/SiO2 particles; d) particle size distributions of Fe2O3 nanoparticles in water (solid line) and of Fe2O3/TMAPS/DNA/SiO2-C6 particles in toluene (dashed line).
Magnetization data (Figure 2.1b and Table 2.1) revealed soft magnetic behavior of the nanoparticles, with a saturation magnetization of 50 emu/g, and nearly zero hysteresis as shown by the inset of Figure 2.1b, indicating superparamagnetic properties.
Hydrodynamic size distribution of the produced Fe2O3 nanoparticles was obtained using an analytical photocentrifuge (LUMiSizer). The mean hydrodynamic particle size according to Stokes’ law (calculated using a density of 5.6 g/cm3) was 43 nm.
41
Table 2.1 Saturation magnetization and corresponding coercivity and remanence data for pristine Fe2O3 particles and for Fe2O3/TMAPS/DNA/SiO2-C6.
Sample Magnetization Coercivity Remanence (emu/g) (Oe) (emu/g)
Fe2O3 50 2 0.2
Fe2O3/TMAPS/DNA/SiO2-C6 19 2 0.1
The Fe2O3 nanoparticles were reacted with n-trimethoxysilylpropyl-n,n,n- trimethylammonium chloride (TMAPS),51, 155, 172 to have surface ammonium groups able to interact with negatively charged DNA (Figure 2.2a). The surface functionalization of the iron oxide with TMAPS was studied by IR-spectroscopy (Figure 2.3), by zeta potential measurements and by element microanalysis (Table 2.2).
Figure 2.2 a) Synthesis of Fe2O3/TMAPS/DNA/SiO2-C6 particles and b) analytic route of DNA recovery from oil particle suspensions and subsequent quantification by qPCR.
DNA was readily adsorbed on the surface of the positive charged magnetic nanoparticles, as evidenced by the IR spectrum (Figure 2.3), the zeta potential value (Table 2.2), and by Qubit fluorometric quantitation (1.0 ± 0.5 µg DNA per mg of particles). In the following step the particles were encapsulated in silica by using TMAPS as co-interacting species and tetraethoxysilane (TEOS) as a silica precursor,51, 155 as illustrated in Figure 2.2a. 42
Figure 2.3 IR spectra of a) Fe2O3, (b) Fe2O3/TMAPS, c) Fe2O3/TMAPS/DNA, d)
Fe2O3/TMAPS/DNA/SiO2, and e) Fe2O3/TMAPS/DNA/SiO2-C6 particles.
Table 2.2 Zeta potential data and element microanalysis for carbon, hydrogen and nitrogen.
Sample C% H% N% Zeta Potential (mV)
Fe2O3 0.22 0.75 0.09 6
Fe2O3/TMAPS 2.81 1.15 0.49 34
Fe2O3/TMAPS/DNA 3.39 1.15 0.93 -24
Fe2O3/TMAPS/DNA/SiO2 3.12 1.33 0.64 -24
Fe2O3/TMAPS/DNA/SiO2-C6 5.13 1.60 0.66
A dense silica coating was grown to ensure complete sealing of dsDNA adsorbed onto the iron oxide nanoparticles. Electron microscopy (Figure 2.1c) of the obtained nanostructured particles illustrates the maghemite cores embedded in the silica matrix. The presence of the nanometer thick silica coating was also verified by IR-spectroscopy (Figure 2.3, band around 1090 cm-1) and by zeta potential measurements (Table 2.2). As expected, the presence of the
SiO2 shell reduced the magnetic performance of the material. The observed decrease of the saturation magnetization from 50 to 19 emu/g suggested the presence of ~60 wt% silica in the final material.
For read out of the presence and concentration of the tag, DNA has to be released from the particles. It is well known, that silica dissolves rapidly in fluoride comprising solutions 2- (forming SiF6 ), but with the aid of some acidity iron oxide is also attacked and dissolved in buffered fluoride solutions. Indeed, upon mixing aqueous suspensions of the particles with a 43 buffered fluoride comprising solution (BOE, F- = 0.025 wt% in water, pH ~4), the silica top layer as well as the iron oxide cores were dissolved, and the DNA molecules were freed. The compatibility of dilute BOE solutions and DNA (DNA is not affected by F- ions) has 51, 155 previously been shown. After dissolving Fe2O3 and SiO2 the released DNA was amplified directly using a standard quantitative PCR protocol, without the need for purification prior analysis. Even at the highest concentration of particles in water (106 µg/L = 1 g/L) the concentrations of the ionic species in the final PCR mix did not affect the 2- 2/3+ + - polymerase efficiency (SiF6 , Fe , NH4 and F at calculated concentrations of ≤ 12 µg/mL, 2 µg/mL, 41 µg/mL and 63 µg/mL, respectively). Figure 2.4 shows standard curves (left, triangles), and amplification curves (right) obtained from dilution series of DNA particle dispersions in water.
Figure 2.4 Standard curves (left) and amplification curves (right) of 10-fold serial dilution of particle dispersions obtained both by direct sampling (triangles) and by magnetic separation (squares) from water and by magnetic separation from toluene (spheres). At each particle dilution the use of magnetic separation allowed for sample pre-concentration lowering the cycle threshold CT to significantly lower values (~8 cycles, i.e. 100x more concentrated).
In order to show the advantages of magnetic separation, the particles present in 1 mL of solution were separated from an Eppendorf tube with the use of an external magnet prior to dissolution in BOE. With this procedure 100 times more particles (and DNA) could be sampled, resulting in a ~8 cycles lower threshold CT and a consequently ~100 times lower detection limit. 1 ppb (1 µg/L) of particles could be detected (CT ~24).
To obtain tags dispersible in oils, the silica surface was further reacted with n- hexyltrimethoxysilane to obtain hydrophobic particles. Besides observing substantially 44
increased compatibility with hydrophobic organic solvents, C6 surface functionalization was confirmed by IR spectroscopy (Figure 2.3; bands at 2957, 2926, 2858 cm-1 indicating C-H bonds) and element microanalysis (Table 2.2).
The suspension stability in toluene and the corresponding particle size were assessed by photocentrifugation (LUMiSizer). The hydrodynamic particle size distribution in toluene, with a mean value of 135 nm, is displayed in Figure 2.1d. It was obtained from Stokes’ law, 3 assuming a particle density of 3.3 g/cm (60 wt% SiO2, 40 wt% Fe2O3, see Appendix, subsection A.1.1 for density calculation). This corresponds to a calculated particle sedimentation velocity (in absence of additional centrifugal or magnetic forces, see Appendix, subsection A.1.2 for sedimentation velocity calculation) of 410-8 m/s (i.e. sedimenting at a rate of 4 mm per day).
The surface functionalized particles were dispersed in toluene at various concentrations (ranging from 1 to 106 µg/L) and the release of the DNA from the particles was attempted after evaporation of the toluene. However, DNA recovery was far from quantitative and differed in about 10 cycles compared to the aqueous sample. It seems that residues from the evaporated toluene were interfering with the polymerase reaction. In this scenario, magnetic separation becomes essential, since it permits to collect the encapsulates and discard the supernatant instead of drying it. Additionally bigger volumes can easily be used, increasing the amount of collectable encapsulates and recoverable DNA. As expected, via magnetic separation we observed DNA amplification for all evaluated particle dilutions (from 104 g/L down to 1 g/L = 10 ppm to 1 ppb), both in water and toluene. In all cases the cycle threshold difference from the positive sample to the negative control was ≥ 7 (Figure 2.4). No statistical difference between the aqueous sample and the toluene sample was evident.
A proper tag must not only be dispersible in the final fluid, it must also be invisible, still being detectable. Therefore, it was important to assess the optical properties of particle dispersions. Visible absorption spectra and photographs of particles in toluene suspensions of different concentrations are shown in Figure 2.5 in comparison to pure toluene. We observed that the tags did not affect the transparency of the dispersant at concentrations below 103 g/L: particle suspensions appeared like pure toluene to the naked eye, and possessed identical visible absorption spectra (max visible absorbance < 0.02). 45
Figure 2.5 Visible absorption spectra of several Fe2O3/TMAPS/DNA/SiO2-C6 suspensions in toluene, at different particle concentrations. A photograph insert of toluene (center), of 1 µg/L 4 (left) and 10 µg/L (right) Fe2O3/TMAPS/DNA/SiO2-C6 particles in toluene is included.
Via magnetic separation, we were also able to detect and quantify DNA from particle dispersions in decalin and oils. Decalin was chosen as a non-polar high-boiling model compound. Where necessary (decalin and olive oil, see Experimental section) the oil was first diluted with toluene to decrease its viscosity, the particles were separated by the use of a magnet and were washed with toluene prior to DNA release in BOE buffers. The taggant information (base sequence) was maintained during the procedure, as evidenced in the sequencing chromatograms (Sanger sequencing) in Figure 2.6.51
Figure 2.6 Sequencing chromatograms of bases 95-109 of unprocessed dsDNA sequence (upper panel) and encapsulated/recovered from decalin (lower panel). 46
We tested temperature stability of the taggants in comparison to unprotected DNA. We performed the temperature treatment in water (before the hydrophobization step), since water represents the only good solvent for free DNA and encapsulated DNA. The two systems were heat treated at 95°C for durations of up to 24 h (Figure 2.7a). After 4 h treatment at 95°C, almost all the free DNA was degraded ( 0.002%), while about 15% protected DNA survived. We also performed stability test for shorter durations and higher temperatures (Figure 2.7b), to show the increased stability of protected DNA to high level of stress. For this purpose we used dispersions of the particles in decalin (bp = 189-191°C), which were heat treated at temperatures between 100°C and 160°C for 30 minutes. For comparison, unprotected DNA solutions in water with similar DNA concentrations were also treated. About 80% DNA in
Fe2O3/TMAPS/DNA/SiO2-C6 particles resisted the treatment at 100°C for 30 min, conditions at which less than 0.05% unprotected DNA survived (Figure 2.7b). Although considerable amounts of DNA were degraded at treatments at higher temperatures, the taggants were still detectable after heating to 160°C for 30 minutes (unprotected DNA was no longer detectable after heating in water to 120°C).
Figure 2.7 a) qPCR analysis of encapsulated DNA heat stability in water at 95°C for durations of up to 24 h compared to that of unprotected DNA in water. Gray bars represent protected DNA, patterned bars free DNA. b) qPCR analysis of encapsulated DNA heat stability in decalin at temperature between 100°C and 160°C (30 min treatment) compared to that of unprotected DNA in water (pressurized at 120°C). Rhombus patterned bars represent protected DNA in decalin, line patterned bars free DNA in water. (*) indicates data below detection limit (<10-7 µg/mL DNA). (**) Free DNA samples were not evaluated for temperatures exceeding 120°C due to water evaporation.
47
We additionally tested the long-term storage capability of the particles in decalin by an accelerated aging test at 65°C for 35 days (equivalent to storage at RT for 2 years), without substantial losses (Figure A1.3).
To show the applicability of the magnetic taggants we utilized gasoline as an example for a fuel, bergamot oil as an example of a cosmetic oil and extra virgin olive oil as an example of a food grade oil. The taggants formed dispersions in all three systems. Particle sizes were assessed in gasoline and limonene (the major constituent of bergamot oil) yielding 160 and 162 nm average diameters (Figure A1.4; particle analysis using analytical photocentrifugation was impeded by the high viscosity and color of the olive oil). The particle size and the low concentration used suggest that sedimentation/aggregation phenomena are not significant (see Appendix, subsections A.1.2 and A.1.3 and Table A1.1). A dilution series over the particle concentration range 1-100 µg/L was obtained for each oil by qPCR. Mean CT value for negative control (qPCR reaction set up with water instead of sample) was also recorded. All samples showed a clear dependence of the CT (and hence of DNA concentration) on the amount of taggant present (Figure 2.8).
Figure 2.8 qPCR analysis of DNA in oil dispersions (100 µg/L, 10 µg/L and 1 µg/L
Fe2O3/TMAPS/DNA/SiO2-C6 particles in bergamot oil, gasoline and olive oil), showing significant CT values differences (two-samples t-test) between oil samples at different concentrations; *) p < 0.05; **) p < 0.001; ***) p < 0.1.
To statistically analyze qPCR results we carried out two-samples t-test (Appendix, subsection
A.1.4), showing for each oil that mean Cts of any pair of standard curve points recorded were significantly different if averages were taken from 5 independent samples (Figure 2.8 and Table A1.2 for complete data set). Oil samples with merely 1 µg particles per liter were also 48 differentiated from negative control group to a certainty of > 99.9% (p < 0.001 for each oil), which ensures discrimination of adulterated oil base products. This is close to the limit of detection of the method: an additional 10-fold dilution of the oil sample would only allow differentiation from blanks to a certainty of < 80% (p > 0.2 for CT ≥ 30.5), too low for a commercial use of the product. Closer differentiation of dilution would be possible if more individual samples were averaged, but is limited by the intrinsic logarithmic scale of the qPCR method.
All three fluids are known objects of adulteration and chromatographic and spectroscopic techniques are currently being used to gain insight into the modifications involved.142-143, 145- 146 Results are often unsatisfactory, since it is not always possible to discriminate components added/substituted. DNA fingerprinting has been applied to identify fraudulent olive oil and/or determine its origin. However, it involves DNA extraction procedures which not always ensure enough DNA for analysis, and have to be optimized basing on the amount/type of PCR inhibitors present in the oil sample.144 In contrast thereto, the technology presented here is a universal method which could be applied to each oil type and oil sample without the need for specific optimization.
At a taggant concentration of 1 µg/L (= 1 ppb) and a calculated material cost of the taggant of ~200 USD/g the financial burden of marking a given oil with this technology is 0.02 ¢/L. (calculated from chemical costs on laboratory scale, worst case, see Appendix, Table A1.3 for details). This is well below 0.02 % of the final product value for a bulk commodity and even lower if used to mark a branded product. Furthermore, the cost is in the range of current expenses for the most accepted methods of product labeling (printing barcode on cardboard).
The taggants were designed with biocompatbility in hindsight, as both iron oxides and silica particles are considered biocompatible and a recent study showed that iron may enhance the biodegradability of silica.174 Also both materials are approved as food additives (amorphous silica = E551, iron oxide pigment = E172) and the Experts group on Vitamins and Minerals set a safe upper level for silicon intake at 700 mg per day.175 Still the use of the magnetic taggants to regulated applications in pharmaceutics, food and cosmetics will require further toxicity testing and approval. 49
2.4 Conclusion
We developed a method to produce inert taggants for oil items. They consist of hydrophobic nanoengineered particles embedding artificial dsDNA sequences. The particles have a core/shell structure, made of iron oxide and silica. The iron oxide is responsible for the magnetic properties, while the surrounding silica matrix acts as protective barrier and confers heat stability and surface functionality. DNA could be recovered from the particles upon dissolution in fluoride comprising solution and analyzed by qPCR and Sanger sequencing. The magnetic core of the particles facilitated handling and allowed for sample concentration. The magnetic tags were easily retrieved from oils, identified and quantified on a logarithmic scale: we could successfully detect them in bergamot oil, olive oil, and gasoline suspensions and statistically discriminate 10 fold dilution steps of the products. Incredibly small amounts of particles (down to 1 µg/L) and minute volumes (1 mL) were sufficient to perform authenticity tests of the oil products. Furthermore, the method is universal, since does not require procedure optimization based on oil type.
50
51
3 Silica microcapsules for long-term, robust and reliable room temperature RNA preservation
Published in parts as:
M. Puddu, W. J. Stark and R. N. Grass
Adv. Healthc. Mater. 2015, 4, 1332-1338.
52
3.1 Introduction
RNA research has grown enormously in the last 30 years and in vitro synthesis of RNA molecules is a routine laboratory procedure. While previously RNA was merely known as an intermediate between genetic information and proteins, it is now considered to be a versatile and useful molecule with a variety of functions, and has been protagonist of milestone discoveries in 21st century.176 Transcriptome analysis (expression profiling), which catalogue and quantify RNA in organisms by real-time reverse transcription-polymerase chain reaction (RT-qPCR),177-178 microarrays,179-180 or next-generation sequencing (RNA-Seq),179, 181-182 is well established and has an essential role in understanding biological mechanisms and in clinical diagnostics. Since the discovery of naturally occurring RNA enzymes (ribozymes),183- 186 there has been growing interest in the design of new ribozymes to manipulate gene expression, especially for the development of therapeutics.187 Aptamers, RNAs binding to specific proteins or ligands and inhibiting their functions,188-189 have demonstrated their therapeutic ability.190-191 RNA-based technologies to regulate gene expression, particularly antisense RNAs187, 192-193 and RNA interference,176 are being pursued to develop treatments for diseases in humans. A new promising genome engineering technology, the CRISPR/Cas system, uses short RNAs to direct Cas9 nucleases to a target DNA sequence and induce site- specific cleavage, offering the chance to tackle a range of diseases.194-195
Despite the utility and the opportunities that RNA provides in many fields, it is a very labile molecule. It is rapidly oxidized due to the action of reactive oxygen species (ROS).196 Furthermore, the additional hydroxyl group on its ribose ring makes RNA more prone to hydrolysis than DNA, since it is capable of self-hydrolizing through transesterification.197-198 A phosphodiester bond in RNA has an estimated half-life more than 104 times shorter than in DNA.199 The cleavage of the phosphodiester bond can be catalyzed by several agents (such as acids or bases).197 Ribonucleases (RNases) are though the main RNA degradation promoters. Such enzymes are found everywhere (common sources are skin, dust etc.), are very stable and difficult to inactivate since they do not require cofactors for their activity.200-201 The ubiquitous presence of RNases and the intrinsic chemical instability make RNA handling and storage problematic. This aspect slowed down the application of RNA in drug discovery as well as in nanotechnology,202 as building block for the fabrication of nanostructures, in comparison to the more mature DNA-based techniques.
RNA integrity is essential for successful use in downstream application and for obtaining meaningful gene expression information. If RNA is not properly stored, the resulting damage 53 can affect and invalidate the outcome of expensive, time and work-consuming analysis, such as RT-qPCR and micro-arrays. 203-205 Hence, RNA storage needs to be robust and reliable to ensure an adequate RNA quality after storage and shipment.
For very short periods RNA can be stored in RNase-free water (typically treated with diethylpyrocarbonate).200 However, long-term storage requires dry strategies or low temperature (-80°C or under liquid nitrogen), often combined with the use of stabilizers, to avoid or at least reduce degradation.198, 205 Commercial reagents, such as RNAlater (AMBION), have been developed to stabilize and protect RNA, achieving a better preservation than in water. Still the product requires freezing temperature (-20°C) for long- term storage, while at room temperature can preserve the sample only for one week. Also solubilization in formamide allows long-term RNA storage at -20°C.200 The use of a cold chain though does not completely exclude degradation, since even at low temperatures RNase activity is significant.198 Moreover, the use of freezers implies the need for space, energy input and maintenance costs. Cold strategies are especially problematic when a RNA sample has to be shipped for analysis, since quality of the sample is often compromised due to thawing as a consequence of delays or temperature fluctuations. Additionally, shipment requires the use of bulky and heavy boxes filled with dry ice, which are generally expensive to ship.198, 205-206
For these reasons, several attempts have been made to develop room temperature RNA storage methods. RNAstable (Biomatrica) is a commercially available product preserving RNA dry at room temperature.205-206 Whatman FTA cards are chemically treated filter papers which can stabilize RNA at room temperature for limited amount of time.207 A technology that consists in storing dry RNA samples in the presence of a stabilizer within stainless steel capsules has also been developed.198 The process ensures efficient RNA protection from water and air oxygen, but involves a complicated encapsulation procedure, requiring controlled atmosphere and laser-welding of the minicapsules.
DNA is a much more resilient molecule, being conceived from Nature to safely store genetic information. Its preservation can be easily guaranteed, and various methods have been established to protect DNA against physical or chemical degrading agents, such as complexation with positively charged species,47, 208 encapsulation in polymer,163 or layered metal hydroxide.63 In previous studies51-52, 92, 155 we have shown that the encapsulation of DNA in silica provides protection against the action of reactive oxygen species, heat and 54 moisture. After silica dissolution with diluted fluoride comprising solutions, intact DNA, encoding the original information, is recovered.
Maintaining RNA stable and protecting it from degradation is instead a significant challenge and requires additional care and effort. Applying the earlier developed encapsulation/release concept to RNA is not trivial: RNA could not withstand the material processing, and the silica dissolution could damage it irreversibly. In the present work we have attempted and succeeded in employing the silica encapsulation technology to generate a robust and reliable method to handle, ship and store RNA for long-time frames.
3.2 Experimental section
3.2.1 Handling RNA
A uniquely allocated lab area was designed to work with RNA; surfaces were treated with RNaseZAP (Sigma), and cleaned with ethanol. RNase-free plasticware and water (Gibco) were used for all the experiments.
3.2.2 RNA sources
A commercially available RNA (StemMACS eGFP mRNA,) was purchased from Miltenyibiotec. RNA was extracted from bovine chondrocytes (bCHs), cultivated to confluency in three T-175 flasks. Cells were trypsinized and centrifuged. The pellet was lysed and large RNA was extracted using NucleoSpin miRNA (MACHEREY-NAGEL) protocols and reagents.
3.2.3 SiO2/RNA microcapsule synthesis
Commercial silica particles (SiO2-Research Particles, 0.136 µm, Microparticles GmbH) were surface modified with ammonium groups by mixing a particle suspension in isopropanol (1 mL, 50 mg/mL) with 10 L N-trimethoxysilylpropyl-N,N,N-trimethylammonium chloride (TMAPS, 50% in MeOH, ABCR GmbH), 51, 86, 155, 172 and stirring overnight (900 rpm) at room temperature. The obtained SiO2-TMAPS particles were then washed 3 times and stored in isopropanol at ~50 mg/mL. Commercial RNA or RNA extracted from bovine chondrocytes was adsorbed on the SiO2-TMAPS particles by mixing a ~3 µg/mL RNA solution in water (1 mL) with the particle suspension (40 µL), followed by two washing cycles (without sonication) and redispersion in water (0.5 mL). Additional TMAPS (0.5 L) was added to the 55 particle dispersion, which was mixed before adding tetraethoxysilane (0.5 L, TEOS, Aldrich).51, 155 After stirring the mixture (900 rpm) for 4 h at room temperature, TEOS (4 L) was added and the reaction was the left to run for 4 days. The obtained microcapsules were then washed by centrifugation and resuspension in 100 L water. If up-concentration is not required, washing can be avoided (see Appendix, Figure A2.5).
3.2.4 Microcapsule characterization
Size analysis of the original silica particles and of the particles after the growth of the RNA protective silica layer was performed by counting the particles with a measured diameter on STEM pictures (see Appendix, Figure A2.1). Characterization by transmission electron microscopy (TEM) was performed using a Tecnai F30 TEM (300 kV, field emission gun, FEI). Scanning transmission electron microscopy (STEM) investigations were performed on a FEI nova NanoSEM 450, operated at an acceleration potential of 30 kV. Particle behavior in aqueous media was studied by zeta potential measurements (Zetasizer Nano, Malvern).
3.2.5 RNA recovery
RNA was release from the microcapsules using a buffered oxide etch solution, prepared with 51 NH4FHF (0.23 g) and NH4F (0.19 g) in water (10 mL), as described in Paunescu et al..
3.2.6 Gel electrophoresis
40 µL StemMACS eGFP mRNA particles were dissolved with 160 µL buffered oxide etch and purified using RNeasy MinElute Cleanup Kit (Qiagen). The obtained RNA solution and an unprocessed RNA solution were denaturated at 65 °C for 5 min prior to electrophoresis (2% agarose with SYBR® Gold stain, Invitrogen).
3.2.7 One-step RT-qPCR cDNA synthesis (reverse transcription) and qPCR reaction were combined in a one-step procedure performed sequentially in the same tube, simplifying the experimental setup and reducing the risk for contamination. Following reaction components were mixed together: Takyon No Rox SYBR MasterMix dTTP Blue (25 µL, Eurogentec), primer mix stock (5 µL, containing both forward and reverse primers 1 µM each), additive (0.5 µL, Eurogentec), EuroScript RT H- and RNase Inhibitor (0.25 µL, 50 u/µL EuroScript RT H-, 20 u/µL RNase Inhibitor, Eurogentec), template RNA (1 µL), RNase-free water (18.25 µL). Primer sequences for StemMACS eGFP mRNA were generously donated from Miltenyibiotec. Primers for bCH 56 mRNA were: 5’-GCCAAGATCCACTATCGGAAA-3’ (forward), 5’-AGGACCTCTGTG AATTTGCC-3’ (reverse).209-210 Each one-step RT-qPCR reaction was performed with a Roche LightCycler 96, using the following parameters: reverse transcription at 48°C for 30 min, Takyon activation and EuroScript deactivation step at 95°C for 5 min, followed by 40 cycles of 15 s at 95°C, 30 s at 60°C and 30 s at 72°C.
3.2.8 RNA absolute quantification
RNA sample concentration before and after the various treatments was determined by interpolation from standard curves (see Appendix, Figure A2.3 and A2.4), generated from RNA dilution series of known concentration (measured by Qubit RNA HS assay used with Qubit fluorometer, Invitrogen).
3.2.9 Reactive oxygen species treatment
Particle suspension or free RNA solution (5 μl) was mixed with a L-ascorbic acid solution
(2.5 μl, 20 mM, Acros Organics), H2O2 (12.5 μl, 20 mM, Merck) and CuCl2 solution (17.5 μl, 500 μM, Fluka). After 10 min the treatment was stopped by adding EDTA (17.5 μl, 100 mM, Biosolve). BOE (20 μl) was then added to the particles to dissolve them, while the same volume of water was added to the free RNA. The solutions were dialyzed against pure water (Millipore 0.025 μm VSWP) before RT-qPCR. All experiments were repeated twice and assayed in triplicates by RT-qPCR (n = 6). Data are shown as mean concentration ± standard deviation.
3.2.10 RNase treatment
5 μl of particle suspension or free RNA solution were mixed with 1X buffer (50 µL, New England Biolabs) and RNase If (2 µL, New England Biolabs), and incubated at 37°C for 20 min. Particles were then centrifuged and washed with water (50 µL) to remove RNase before adding the BOE solution (20 µL). The obtained solution was then dialyzed against water before performing RT-qPCR. Free RNA solutions were cleaned from RNase using QIAquick PCR purification kit (Qiagen).
3.2.11 Long term RNA stability
RNA particles suspension (5 µL) was pipetted in RNase-free Eppendorf tubes and dried overnight in a fume hood. Free RNA (20 µL) was added to RNase-free Eppendorf tubes or to RNAstable tubes, and dried in a vacuum concentrator (Concentrator plus, Eppendorf). 57
Prepared samples were stored (with opened lid) in a closed jar in an oven at 65°C and 50% RH, achieved with a saturated solution of sodium bromide (NaBr ≥ 99.5%, Sigma). After the long term storage, RNA was recovered by adding water (20 µL) to the dried RNA or the RNAstable tubes. Particles were dissolved with BOE (20 µL) and dialyzed against water before RT-qPCR analysis. Data are shown as mean logarithm of the concentration ± standard deviation (n = 3).
3.2.12 Capillary electrophoresis
StemMACS eGFP mRNA particles (40 μL) were treated with reactive oxygen species as described above or heated at 70°C for 20 min. Then they were dissolved with BOE and purified with RNeasy MinElute Cleanup Kit (Qiagen) before being analyzed from Microsynth AG (Agilent 2100 Bioanalyzer).
3.2.13 Sanger sequencing
5 µL bCH-mRNA particles were treated or not and later dissolved with BOE (20 µL). The obtained mRNA solution was dialyzed and then reverse transcribed and amplified by RT- qPCR. PCR products were purified with QIAquick PCR purification kit (Qiagen), diluted to 2 µg/mL, mixed with forward primer (same one used for RT-qPCR, 2 µM) and sequenced in direction 5’-3’ from Microsynth AG.
3.3 Results and discussion
A method to preserve RNA in silica microcapsules has been developed. To produce the RNA microcapsules, commercial silica particles were modified with TMAPS to obtain a positive charged surface,51, 86, 155, 172 as confirmed by zeta-potential measurements (+ 40 mV). RNA, being negatively charged, interacts electrostatically with this positive support and binds to it, decreasing the surface charge of the particles (+ 20 mV). Bound RNA was then encapsulated in a dense silica layer by a sol-gel process, using TMAPS as co-interacting species and TEOS as silicon source (Figure 3.1).51, 155
The whole procedure is extremely simple and only requires several minutes of handling (see Appendix, subsection A.2.2 and Figure A2.5). In a standard protocol, a commercially available messenger RNA (mRNA) was encapsulated. TEM micrographs in Figure 3.2a and Figure A2.1a illustrate the silica microcapsules containing the mRNA.
58
Figure 3.1 Silica comprising RNA microcapsules: synthesis scheme.
Figure 3.2 a) TEM picture of the SiO2/RNA microcapsules; b) size distribution of the SiO2 particles before and after the growth of the silica capping layer.
The presence of the protective silica coating was confirmed by zeta-potential measures (- 21 mV), and by analyzing particle size on scanning transmission electron microscopy images (Figure A2.1b and c). A mean particle diameter of 119 nm was calculated for the original silica particles, and of 137 nm for the particle after the silica growth, which corresponds to a 9 nm thick silica capping layer (Figure 3.2b).
RNA was released from the microcapsules upon rapid silica dissolution in a buffered fluoride- ─ 2─ containing buffer (BOE, 2.5 wt% F in water, pH ~ 4), forming SiF6 and leaving a clear RNA solution without solid residues. Recovered RNA was detected/measured by Qubit RNA HS Assay Kit, used with the Qubit fluorometer, and an early assessment of RNA integrity was done by gel electrophoresis (Figure A2.2). RNA quality was further characterized by capillary electrophoresis (Agilent Bioanalyser), the standard in RNA integrity assessment. RNA was also incorporated in a commercially available synthetic matrix (RNAstable),205-206 for comparison. Retrieved RNA quality was checked using capillary electrophoresis. Figure 3.3 59 shows the electropherograms of the original RNA, of the RNA recovered from RNAstable and from the microcapsules, showing similar profiles.
Figure 3.3 Assessment of RNA intactness by capillary electrophoresis (Agilent Bioanalyser). A graph of the size distribution of RNA fragments is shown for a) original RNA, b) RNA stored in RNAstable, c) RNA stored and directly released from the silica microcapsules or additionally treated with d) ROS and e) heat before the recovery. Similar RNA quality is depicted for all the samples.
This result suggests that the RNA integrity is maintained during the encapsulation and recovery processes. Thus RNA, as well as DNA, is not affected by ammonium groups, hexafluorosilicate ions, fluorides as well as by the acidity of the dissolution buffer.51, 155
A demonstration of the physical and chemical protection provided by the silica microcapsules was obtained by exposing encapsulated RNA to reactive oxygen species (ROS) or heat (70°C for 20 min). Recovered RNA integrity was assessed with Agilent Bioanalyser system (Figure 3.3). The produced electropherograms reveal approximately unvaried RNA quality.
The compatibility of the encapsulation and release procedures with the most common RNA analysis technique in research and diagnostic procedures, i.e. RT-qPCR, was then tested. In gene expression measurements, RT-qPCR has established as the benchmark for the detection and quantification of RNA samples,177-178, 204 being the most sensitive and accurate method. We could successfully analyze and quantify the RNA released from the microcapsules by a one-step RT-qPCR reaction (Figure 3.4). 60
Figure 3.4 RT-qPCR detection and quantification of unprotectd RNA and silica encapsulated RNA before and after exposure to ROS or RNase. While unprotected RNA is disintegrated by the treatment, RNA protection against ROS and RNase activity by encapsulation within silica is evidenced.
A microcapsule loading capacity of ~ 0.5 µg RNA mg─1 particles was quantified by RT-qPCR (using the standard curve in Figure A2.3, as described in the Experimental section), confirming what was measured by the Qubit fluorometer. Reverse transcription and cDNA amplification by qPCR could even be performed without additional purification if samples were diluted prior to analysis 1:10 (see Figure A2.5). This shows that the low fluoride concentrations required to dissolve the silica do not interfere with the action of the reverse transcriptase or polymerase.51-52
To evaluate the protective capability of the silica microcapsules comprising RNA, they were treated with ROS or Ribonuclease If (RNase If). For comparison, unprotected RNA solutions were also treated with the degrading agents. After exposure to ROS or RNase, the microcapsules were dissolved and retrieved RNA was analyzed by RT-qPCR. As shown in Figure 3.4, non-encapsulated RNA was almost completely disintegrated by the ROS and RNase activities, while RNA within the microcapsules could better survive the aggressive treatments. 85% of the encapsulated RNA withstood the ROS treatment. The losses measured after treating encapsulated RNA with RNase are connected to the well-known difficulties in washing away the excess RNase and in inactivating it completely. However, during storage in normal conditions we do not expect such high dose of RNase and therefore such losses become neglectable. These results therefore prove that the storage within the silica microcapsules increase the RNA chemical stability. 61
In order to estimate the degradation rate of encapsulated RNA, sample aging was accelerated by heating in the presence of air moisture. The accelerated aging study was performed in parallel on non-protected dried RNA samples and on RNA stored in RNAstable to compare degradation rates. Samples were kept at 65°C in a 50% relative humidity (RH) atmosphere for about 100 hours, and analyzed by RT-qPCR at different time points. Results are reported in Figure 3.5, as natural logarithm of the concentration versus time, according to a first order decay equation.211
Figure 3.5 RNA samples stored in silica microcapsules at 65°C were stabilized and showed slower degradation as compared to dried control samples and samples store in the RNAstable matrix.
After 104 hours, only 0.05% and 0.08% of, respectively, unprotected RNA and RNA stored in RNAstable survived. Instead, 32% of the RNA within the silica microcapsules was integer.
We therefore kept the SiO2/RNA microcapsules for a longer period of time at the accelerated aging conditions. After 1 month under these most aggressive conditions (last point of the kinetics in Figure 3.5), we could still retrieve and detect some RNA. The decay rate constants at 65°C (k65°C), calculated assuming first order kinetics (see Appendix subsection A.2.1 and table A.2.1), were 2 10─5, 2 10─5, and 2 10─6 s─1 for unprotected RNA, RNA protected within
RNAstable matrix and within SiO2, respectively. Encapsulation in SiO2 therefore affords an approximately 10-fold decrease in the RNA decay rate at 65°C with respect to unprotected RNA and RNA stored in the commercial RNAstable matrix. The degradation kinetic constant at room temperature (k25°C) for encapsulated RNA, calculated according to the Arrhenius' equation using the activation energy estimated by Fabre et al. for solid-state RNA, was ─9 ─1 7 10 s . This corresponds to a half-life (t1/2) at room temperature of 3 years for the 847 nt 3 RNA strand, i.e. a RNA phosphodiester bond has a t1/2 of ~3 10 years in silica at 25°C. 62
Although numbers on uncatalyzed RNA degradation reported in literature deviate by about a factor of 10,199, 211-212 the encapsulation within silica affords a stability increase of 100-1000 compared to RNA in aqueous solution at pH 6-7 (Figure 3.6),199 where stability is maximal under aqueous conditions.211-212
Figure 3.6 Phosphodiester bond half-life at room temperature extrapolated according to the Arrhenius’ equation for dried RNA, RNA stored in RNAstable and in silica microcapsules (see Appendix, subsection A.2.1, for detailed half-life calculation), compared to literature data (at room temperature) on RNA stability in RNase-free aqueous solution at pH 6,199 and DNA stability in silica.92
The comparison is even more remarkable when considering RNA storage in solution under strongly acidic or basic conditions (pH < 5.4 or > 8), since the half-life of the phosphodiester bond in RNA reduces by about one order of magnitude.211 Additionally, the last point of the kinetics of RNA in silica (Figure 3.5) simulates storage at 25°C for 24 years. In contrast, calculated half-lives at 25°C were as low as ~4 months for both unprotected RNA and RNA protected within RNAstable.
These results agree with other studies,92, 198 concluding that nucleic acid degradation rate is reduced by water removal. Common laboratory plastic tubes, as well as RNAstable tubes are not water tight. As a consequence of the exposure to a humid atmosphere, the dried RNA rehydrates and the occurrence of phosphodiester bond cleavage phenomena increases. In contrast, silica microcapsules hermetically isolate the RNA from the environmental moisture and nucleases, maintaining the dehydrated state and therefore stabilizing RNA against hydrolysis for longer time frames. This exceptional ability to maintain a water-free 63 atmosphere over time mimicks the protection provided to nucleic acids in ancient fossils.51, 92, 162
The stainless steel minicapsules invented by Imagene198 also aim at maintaining a water and oxygen-free environment. In contrast to the laser-welding of stainless steel under anhydrous argon needed to seal the capsules, we propose an easy protocol which does not require expensive procedures or equipment, facilitating the archiving of the increasing numbers and sizes of RNA libraries. Facile room temperature handling is achieved, as well as a reduction of cost and space during transporting and storage, since dry ice storage boxes and freezers are not needed.
RNA microcapsules present properties (silica shell thickness, nucleic acid loading capacity) comparable to those of the earlier developed encapsulated DNA systems. Encapsulated RNA resistance to ROS was analogous to what measured for DNA.51, 92, 155 When comparing the accelerated aging data, the results are even more astonishing. Surprisingly, the encapsulated RNA decay rate at 65°C is nearly identical to that of DNA,92 meaning that their stability within the silica microcapsules is equivalent (Figure 3.6), despite their intrinsic differences in chemical properties. This remarkable achievement provides a new tool in several applications where DNA has been historically preferred to RNA because of his improved stability rather than for its performance.
The compatibility of the storage system with common methods for mapping transcriptomes was further checked. Here we chose a sequence-based approach instead of a hybridization base one. The direct determination of the complementary DNA (cDNA) sequence allows to avoid cross-hybridization problems. Moreover, for low throughput, Sanger sequencing was the method of choice. For the application test we prepared RNA from biological samples (bovine chondrocytes), encapsulated it within silica, and sequenced it upon release and purification. Encapsulated RNA samples were additionally exposed to degradation sources (ROS or RNase) before retrieving the RNA. For all the samples, released RNA could be successfully reverse transcribed to cDNA, further amplified and quantify in a one-step RT- qPCR reaction, using primers for the ribosomial protein L13 gene (RPL13), a reference gene used in gene expression.209-210 Also in this case an encapsulation capacity of 0.5 µg/RNA mg particles was measured (using the standard curve in Figure A2.4). A quality control of all the obtained cDNA samples was done by gel-electrophoresis (Figure 3.7), before analyzing them by Sanger sequencing (Figure 3.8). 64
Figure 3.7 RNA samples recovered from the silica microcapsules were successfully used as templates for cDNA synthesis and subsequent amplification by PCR. Reverse transcription and PCR were possible also for samples exposed to ROS and RNase during the storage in silica.
Figure 3.8 cDNA samples obtained from a) original RNA, b) RNA directly recovered from silica microcapsules or additionally treated with c) ROS and d) RNase before the release, were successfully sequenced.
65
As shown in Figure 3.8, sequencing was successful for all the encapsulated samples (untreated and treated with ROS or RNase), producing good quality chromatograms with low background noise, comparable with the result produced from the original, non-encapsulated sample.
These results prove that the SiO2 microcapsules represent a suitable storage system for sample destined to transcriptome analysis by RT-qPCR and sequencing, overcoming the challenging and persistent problems connected to RNA fragility.
3.4 Conclusion
A novel storage system has been developed to preserve RNA for long-term periods, eliminating some of the variables associated with RNA instability. It consists of RNA embedded in silica microcapsules, grown by sol-gel chemistry. The microcapsules decrease the rate of RNA degradation by preserving a nearly anhydrous environment, which significantly reduces hydrolysis. RNA released from the microcapsules was tested for standard downstream applications (including cDNA synthesis, RT-qPCR and gene sequencing), exhibiting no inhibition or interference. It was found that the silica microcapsules, with the lowest water content, ensure better chemical stabilization against hydrolysis in comparisons to dry storage or RNAstable, with a calculated phosphodiester bond half-life at 25°C of ~3 103 years. Although RNA is intrinsically less stable than DNA, with a phosphodiester bond half-life approximately 104 times smaller, this technology makes RNA as resilient as DNA. The RNA comprising silica microcapsules are prepared with minimal effort, following a simple procedure, and utilizing basic laboratory equipment. Since freezing is not necessary, the method provides an alternative handling strategy and is particularly suited for samples shipment.
66
67
4 Magnetically deliverable calcium phosphate nanoparticles for localized gene expression
Published in parts as:
M. Puddu, N. Broguiere, D. Mohn, M. Zenobi-Wong, W. J. Stark and R. N. Grass
RSC Adv. 2015, 5, 9997-10004.
68
4.1 Introduction
The process of introducing nucleic acids into cells is crucial for gene therapy, gene function and regulation studies, tissue engineering, as well as for protein manufacturing. The ideal transfection agent, besides giving high transfection efficiency, should be biocompatible and biodegradable, non-toxic to cells, non-immunogenic, and not affecting cell physiology. Additionally, it should be cost-effective, easy to prepare and apply, and be reproducible.
Viral vectors are the oldest and most efficient tools known to deliver genes. Despite ease and effectiveness of virus-mediated transfection, such vectors have shown to provoke an immune response.213 Therefore, several non-viral vectors have been developed, that are less efficient than virus-based systems but exhibit enhanced biosafety.
Among the non-viral vectors, the most commonly-used systems are cationic polymer-based (e.g. polyethylenimine, PEI).214-217 Positively charged reagents interact electrostatically with negatively charged nucleic acids, forming complexes that are up-taken by cells via endocytosis. The technique is easy and inexpensive, but cationic components can be highly cytotoxic.218
Another simple and inexpensive option is calcium phosphate (CaP) mediated transfection.83-84, 121 DNA is mixed with CaCl2 and a saline/phosphate containing buffer to form precipitates carrying DNA into cells. DNA-calcium phosphate co-precipitates have been used for about 40 years to deliver nucleic acids. Biodegradability and biocompatibility of CaP,219-223 whose chemical composition mimicks that of natural bone mineral,219, 224 make CaP precipitates ideal transfection vehicles. However, transfection efficiency is inferior to other available non- viral agents. Additionally the method does not show enough reproducibility since size/shape and therefore transfection efficiency of the co-precipitates depend strongly on experimental factors (e.g. concentration, pH, precipitation time) and handling.84, 122 A combination of CaP and PEI transfection techniques has also been developed to maximize transfection efficiency, leading though to reduced cell viability.223
A promising transfection technique is magnetically guided gene transfection or magnetofection.44, 49, 109 Nucleic acids are associated with magnetic particles (generally composed of iron oxide) and delivery is accomplished by the application of a magnetic field gradient. The method is universally suitable for viral and non-viral vector delivery and highly efficient. Most of the commercial magnetic particles are modified with cationic molecules able to complexate DNA. Magnetofection promises to solve fundamental problems associated 69 with in vivo gene therapy, i.e. low vector availability at the target site, side effects related to high vector doses and vector distribution in non-targeted tissues. The magnetic particles together with the associated vectors can be delivered and retained by magnetic means at the disease site after injection, allowing reduction of vector doses and minimizing gene vector spreading throughout the body. However, biocompatibility in magnetofection can be compromised due to the cytotoxic nature of cationic species often used to mediate DNA binding to the particles.
Nowadays spatial control of gene delivery and expression in specific areas plays an important role not only in gene therapy, but also in synthetic biology.225 By creating spatial patterns of gene expression in a cell culture one could recreate the complex architecture of tissues and organs.225-226 Additionally, one could engineer artificial gene circuits mimicking natural networks to gain basic biological understanding of cellular processes or for practical applications.225, 227 Localized gene expression has been achieved using a robotic microarrayer to deposit gelatin-plasmid solutions,228 near infrared irradiation of gold nanorods-green fluorescent protein gene conjugates,229 lipoplex deposition using microfluidic devices,226 and lyophilization of adenovirus.230 Above all, gene expression localization using magnetic particles represent an easy and precise way to define transfection patterns.225, 231
In this study, we designed and produced a novel, polycation-free, CaP-based magnetic transfection agent which combines the above described advantages of standard CaP and magnetic beads mediated transfection, overcoming at the same time the obstacles presented by the two technologies. We produced a composite nanopowder made of iron oxide and tricalcium phosphate by flame spray synthesis. The material was characterized and tested as transfection mediator with Human Embryonic Kidney 293 cells (HEK 293) to present its effectiveness.
4.2 Experimental section
4.2.1 Nanoparticle production
Iron oxide doped tricalcium phosphate nanoparticles (Fe2O3@TCP) were produced by flame spray synthesis.232-233 Calcium-2-ethylhexanoate (superconductor grade, ABCR) and tributylphosphate (98%, Aldrich) were used as calcium and phosphor precursors, respectively. Iron precursor was prepared according to a previously described procedure.234 The liquid mixture with a Ca/P molar ratio of 1.5 and a final Fe2O3 content of 33 wt% was obtained by 70 mixing the corresponding amounts. All precursors were diluted with xylene to a final metal concentration of 0.8 mol/L. The precursor solutions were fed through a capillary into a burning methane (1.13 L/min, Pan Gas, Switzerland)/oxygen (2.4 L/min, Pan Gas) supporting flame using a gear-ring pump (HNP Mikrosysteme, Germany) adjusted to a rate of 5 mL/min. Oxygen (5 L/min, Pan Gas) was used to disperse the liquid leaving the capillary. Produced nanoparticles were collected on metal filters (G. Bopp & Co AG, Switzerland) with the aid of a vacuum pump. A detailed description of the set-up can be found in Madler et al. 2002.232
4.2.2 Nanoparticle characterization
Fe2O3@TCP nanoparticles were analyzed by nitrogen adsorption according to the Brunauer- Emmett-Teller method (BET; Tristar, Micromeritics), after degassing the nanoparticles at 150°C for 1 h. X-ray diffraction data were acquired using a X'Pert Pro-MPD diffractometer (PANalytical), by exposing the nanoparticle powder to Cu-Kα radiation (λ = 1.54060 Å), over a range of 10 - 70° 2θ with a step size of 0.05°. Iron oxide and TCP phase identification was carried out by means of the X'Pert software PANalytical High Score Plus. Characterization by transmission electron microscopy (TEM) was performed using a Tecnai F30 TEM (300 kV, field emission gun, FEI). Scanning transmission electron microscopy (STEM) investigations were performed on an aberration-corrected HD-2700CS (cold-field emitter; Hitachi), operated at an acceleration potential of 200 kV. An energy-dispersive X-ray spectrometer (EDXS; Gemini system of EDAX) attached to this microscope allowed the recording of EDX spectra and elemental maps. Magnetic properties were investigated by vibrating sample magnetometry (Princeton Measurements Corporation, model 3900). Particle hydrodynamic size distribution was measured with a X-ray disk centrifuge (XDC; Brookhaven Instruments, United States), using a 1.5 % (wt/vol) powder suspension in ethanol, ultrasonicated at 200 W for 5 min. Nanoparticle behavior in aqueous media was studied by zeta potential measurements (Zetasizer Nano, Malvern).
4.2.3 Plasmid preparation pAcGFP1-Endo plasmid (Clontech) was kindly provided by Prof. Dr. Wilfried Weber (Institute of Biology II/BIOSS Center for Biological Signalling Studies, Albert-Ludwigs- Universität Freiburg, Germany). The plasmid (5356 bp), which encodes the green fluorescent protein (GFP), was propagated in chemically competent E. coli and purified using the Qiagen Plasmid Midi kit. Obtained pDNA was diluted to a concentration of 0.6 µg/mL and stored at -20°C. 71
4.2.4 DNA binding assay
DNA (2 µg) was mixed with 5 µL 2M CaCl2, particles (8 µg) and water to a final volume of 100 µL. After incubation for 15 min at room temperature and magnetic separation of the particles, the unbound DNA in the supernatant was quantified by Qubit fluorometer (Invitrogen) using Qubit dsDNA HS assay (Invitrogen, cat. no. Q32851). Excess DNA not bound to Fe2O3@TCP particles was also visualized on agarose gel (see Appendix, Figure A3.2). Before loading the DNA solution in the gel, it was dialysed (Millipore 0.025 µm VSWP) against water to remove the salt.
4.2.5 Cell culture
HEK 293 for transfection experiments were grown in Dulbecco’s modified Eagle’s medium (DMEM, Invitrogen no. 41966) supplemented with 10% fetal bovine serum (FBS, Gibco) and
1% Penicillin/Streptomycin (Gibco). Cells were grown at 37°C and 5% CO2, and subcultivated according to standard cell culture protocols.
4.2.6 Transfection of mammalian cells
One day before transfection, cells were seeded in a 96-well plate (at a density in the range 312-937 cells/mm2) and cultivated in 100 µL media per well. Commercial transfection agents PolyMAG (Chemicell, Germany) and NeuroMag (OZ Biosciences, France), as well as traditional CaP and PEI transfection were used for comparison to the developed material. For each protocol, 0.4 µg pDNA per 96-well was used.
Transfection with PolyMAG (Chemicell, Germany) and NeuroMag (OZ Biosciences, France) was carried out according to the manufacturer’s guidelines. 0.4 µg pDNA was dissolved in 19 µL serum-free media and mixed with the transfection reagent. For Polymag a ratio of 1 µL reagent/µg pDNA was used, while for NeuroMag a ratio of 3.5 µL reagent/µg pDNA was used. The mixture was left to stand 15 min at room temperature to allow for complexation. Then 80 µL of medium supplemented with serum was added.
To perform standard transfection with CaP, DNA was mixed with 4.4 µL of 2 M CaCl2 solution and diluted with sterile ultra-pure water (SIMSV0000-Simplicity UV, Millipore) to a final volume of 35 µL. The prepared CaCl2-DNA solution was briefly vortexed and then added dropwise to 35 µL of HEPES buffered saline (BioUltra, 2X concentrate, Sigma Aldrich), while vortexing. The suspension of CaP-DNA coprecipitates was incubated at room 72 temperature for 15 min before adding 280 µL of media supplemented with serum. The prepared mixture was enough to transfect 3 wells (100 µL per 96-well).
Transfection with PEI was accomplished by mixing pDNA with PEI (branched, Mn~10000, Sigma Aldrich) at a nitrogen/phosphate (N/P) molar ratio of 10 in serum free media (20 µL per well). The mixture was incubated for 15 min at room temperature before adding media supplemented with serum (80 µL per well).
The Fe2O3@TCP powder was sterilized by heating at 200°C for 30 min. A fresh particle suspension was prepared each time before performing transfection experiments. Nanoparticles were dispersed in sterile ultra-pure water at a concentration of 1 mg/mL and sonicated at 200 W for 20 min. The following buffer was prepared in ultra-pure water: 16 g/L sodium chloride (NaCl for analysis EMSURE® ACS, ISO, Reag. Ph Eur.), 10 g/L HEPES ( ≥ 99.5% Sigma), 2 g/L D-(+)-glucose ( ≥ 99.5% Sigma) and 0.74 g/L KCl (Sigma Aldrich), with pH adjusted to ~7.5. The overall transfection procedure is illustrated in Figure 4.1.
Figure 4.1 Scheme of magnetic assisted cell transfection using Fe2O3@TCP particles.
To prepare the transfection mixture for a well, 0.4 µg pDNA was mixed with 2 M CaCl2 to achieve the desired molarity (in the range 62.5-100 mM) in the mixture and with the prepared buffer to a final volume of 20 µL. The CaCl2-DNA solution was briefly vortexed before adding the required volume (2 to 6 µL reagent/µg pDNA) of particle suspension. The transfection mixture was vortexed again and left to stand for 15 min at room temperature. 80 µL of medium supplemented with serum was then added.
The transfection volume (100 µL per 96-well), was then added to the cells. A magnetic field (MagnetoFACTOR-96 plate, Chemicell, Germany) was applied for 30 min to PolyMAG,
NeuroMag as well as to Fe2O3@TCP nanoparticles, while incubating at 37°C and 5% CO2.
After 48 h, live/dead assay and transfection visualization/quantification were performed by fluorescence microscopy. Transfection efficiencies are given as ratio of cells expressing GFP 73 to the total number of cells. Viability data are provided as ratio of live cells to the total number of cells. Proliferation data are given as ratio of average number of cells in a sample to the average number of cells in the untreated control. Quality factors (QFs) are calculated as product of transfection efficiency, proliferation rate and viability. By definition the QF is zero for the untreated control, and 1 in the ideal case (i.e. transfection efficiency = proliferation rate = viability = 100%).
4.2.7 Localization of gene expression
One day before transfection the cells were seeded in a 55 mm diameter Petri dish with 4 mL of media, at a density of 312 cells/mm2. Transfection mixture was prepared as described in the previous section by mixing DNA (8 µg) with 2 M CaCl2 to achieve a CaCl2 concentration of 100 mM in the transfection mixture volume (800 µL), saline buffer and particles (32 µL). The mixture was then diluted with media up to 4 mL. The Petri dish was placed over the MagnetoFACTOR-96 array. The transfection mixture was pipetted onto the cells overlaying the magnet positions. Cells were then incubated for 48 h onto the magnets.
4.2.8 Live/dead assay
Simultaneous determination of live and dead cells was performed by labeling live cells with Hoechst stain and dead cells with Ethidium homodimer-1 (EthD-1). Cells were incubated for 1 h in medium supplemented with 10 µg/mL Hoechst and 2 µM EthD-1 (both purchased from Life Technologies).
4.2.9 Microscopy
Cells were imaged with a Zeiss Observer microscope (Germany). Cell morphology was observed under bright field (96-well plates) or phase contrast (Petri dishes), while live/dead and transfected cells were identified by fluorescence microscopy using DAPI, FITC and DsRed filter sets. The same exposure times were used for all the conditions and images are shown with minor adjustments (color display, as well as background substraction from a reference image and assembly of tiles for petri dish fluorescence imaging, using Fiji).
4.2.10 Cell counting
After staining, cells were counted manually, using Fiji, over an area of 2.4 mm2 (i.e. four times the area illustrated in Figure A3.3 of the Appendix), with blue nuclei giving the total 74 number of cells, red nuclei giving the number of dead cells, and green cytoplasm/nuclei showing successful transfection. Cell counting data are included in Table A3.1.
4.2.11 Statistical analysis
Data are provided as mean value ± standard deviation. Experiments were performed in triplicates. Statistical analyses were performed with OriginPro 8.6. Data were analyzed by one-way analysis of variance (ANOVA), using Bonferroni test for post hoc analysis.
4.3 Results and discussion
We engineered and synthetized a new transfection composite material. Our idea was to produce non-toxic magnetic particles with a surface able to bind DNA and to deliver it inside cells. Iron oxide, widely used in magnetofection and for various biomedical applications, was the obvious choice of magnetic material. An outer biodegradable calcium phosphate layer was highly preferred to iron oxide surface modification with generally more efficient but cytotoxic polycations. Because of their biocompatibility and biodegradability,235-238 iron oxide nanoparticles are the only metal oxide particle clinically approved (e.g. Feridex, a MRI contrast agent), and used as food additives (iron oxide pigment = E172). CaP high biocompatibility and easy integration in the body,219-223 where it is present in solid form or as calcium and phosphate ions,219, 223 has determined its Food and Drug Administration approval as nutrient and its use as bone-substitute in clinical treatments.219 CaP ceramic-iron oxide nanoparticle composites have been previously tested as potential bone replacement and proved to possess good biocompatibility and ability to promote cell proliferation in vitro.239 An earlier attempt to produce magnetic CaP nanoparticles in a multi-step synthesis, comprising biomineralization of CaP on PEI-coated magnetic beads, has also been made.240 The presence of the polycation though did not eliminate the potential toxicity of the formulation.
Flame spray synthesis was used to produce the designed magnetic particles, possessing an iron oxide core surrounded by a TCP matrix. It has been previously shown that the technology is capable of producing multi-component nanoparticles in a single step,241-242 as here in the case of Fe2O3@TCP nanocomposites. Additionally, flame spray technology enables large- scale production of nanoparticles with reproducible size distribution and at low cost.57, 243-244 75
While CaP co-precipitates need to be prepared before performing each experiment,
Fe2O3@TCP powder can be stored in dry state, reducing the experimental variations and facilitating the handling.
The newly synthetized particles were fully characterized (see Experimental section). Brunauer-Emmett-Teller (BET) measurements of the nanoparticles gave a specific surface area of 68 m2/g, which corresponds to a primary particle diameter of 25 nm, calculated assuming spherical particles and using the overall density of the composite material (see Appendix, subsections A.3.1 and A.3.2 for density and size calculations). Transmission electron microscopy (Figure 4.2) and scanning transmission electron microscopy (Figure 4.3a) images supported the assumption of agglomerated spherically shaped nanoparticles.
Figure 4.2 TEM micrograph of Fe2O3@TCP particles prepared by flame spray synthesis.
Energy-dispersive X-ray mapping (Figure 4.3b-e) of the particles showed spherical iron oxide nanoparticles embedded in the TCP matrix, suggesting a core/shell-like structure. The X-ray diffraction pattern, shown in Figure 4.4a, confirmed the presence of Fe2O3 and revealed the coexistence of amorphous and α-TCP. Iron oxide nanocrystallite size, calculated from the XRD pattern by means of the Scherrer formula, was estimated to be 10 nm. As expected, after heating the material to 1000°C, X-ray diffraction pattern showed peaks ascribed to β-TCP (Figure 4.4a).233
Vibrating sample magnetometry data (Figure 4.4b) revealed a saturation magnetization of 9.3 emu/g. Mean hydrodynamic particle diameter measured by X- ray disk centrifuge was 121 nm, significantly larger than the calculated primary diameter, which can be explained by 76 the formation of aggregates. Full hydrodynamic size distribution is given in the Appendix (Figure A3.1 and subsection A.3.3). The average number of primary particles per aggregate was estimated to be 17 (Appendix subsection A.3.4). A surface zeta potential of -14.8 mV was measured.
Figure 4.3 a) STEM image of Fe2O3@TCP nanoparticles and corresponding elemental mapping of b) Fe, Ca, and P merged and of c-e) the single elements.
Figure 4.4 (a) XRD pattern and (b) hysteresis loop of Fe2O3@TCP nanoparticles. 77
After characterizing the produced powder, we proved that the outer TCP surface allows for pDNA binding in the presence of CaCl2, as evidenced by Qubit fluorometric quantitation (~0.02 µg DNA per µg of nanoparticles, changing with plasmid size), gel electrophoresis (Figure A3.2), and surface zeta potential (-18.2 mV). Most probably Ca2+ ions mediate DNA- particles binding through electrostatic interactions.
Particle size and ability to bind pDNA were not changed after powder long term storage at room temperature (see Appendix, subsection A.3.5).
To demonstrate the applicability of the particles as transfection agents we used them to deliver a GFP encoding plasmid into HEK 293 cells. GFP expression demonstrated successful cell transfection. Experimental conditions were optimized to maximize the transfection rate of
Fe2O3@TCP particles while minimizing effects on cell proliferation. Best conditions were found to be particle:DNA ratio 4:1 (4 µg particle per µg of DNA) and a cell density of 937 cells/mm2. Cell viability throughout the experiments was in the range of 92-98%, not showing marked particle induced toxicity. Transfection efficiency in HEK 293 cells increased from ~33% to ~48% with decreasing CaCl2 concentration (Figure 4.5a), when cells were seeded at a density of 312 cells/mm2. Conversely, proliferation increased with increasing
CaCl2 concentration.
Figure 4.5 Transfection efficiency, proliferation rate and viability in HEK 293 cells transfected with Fe2O3@TCP while varying a) CaCl2 concentration (fixed cell density: 2 312 cells/mm ) or b) cell seeding density (fixed CaCl2 concentration: 100 mM).
Samples incubated with naked DNA, CaCl2/saline buffer without particles (-P), and untreated samples (neg. ctr.) are shown for comparison. 78
A loss in transfection was thought as acceptable in favour of an increase in cell proliferation rate, an indicator of cell health. Therefore, a CaCl2 concentration in the range 87.5-100 mM was considered as optimal for transfection purposes. Notably, only a negligible number of transfected cells were observed in presence of CaCl2 without particles. Cell viability in the negative controls (untreated) was always about 100% and, as expected, no GFP was detected.
In Figure 4.5b data resulting from cell transfection with Fe2O3@TCP and 100 mM CaCl2 while varying starting cell density are shown. We observed less transfected cells and higher proliferation rate with increased cell density, less DNA-particle conjugates being available per 2 cell. In optimal conditions (937 cells/mm , 100 mM CaCl2), Fe2O3@TCP were only marginally affecting cell growth, with a proliferation rate of ~86%. As illustrated in Figure 4.6a, cells showed a normal, spread morphology.
Figure 4.6 GFP fluorescence microscopy (upper row) and merged fluorescence and bright field microscopy images (lower row) of HEK 293 transfected with Fe2O3@TCP (a), with the commercial transfection agents PolyMAG (b) and NeuroMag (c), with standard CaP method (d) and with PEI (e). Untreated control is also shown (f). Images are representatives of n = 3 experiments. Scale bar: 50 µm.
After optimizing transfection with Fe2O3@TCP, we compared the technique to other existing transfection methods. Fluorescence microscopy micrographs and overlay images from fluorescence and bright field microscopy (Figure 4.6) show the GFP expression in samples treated with Fe2O3@TCP particles and with the other transfection agents. Transfection, proliferation and viability data are graphically represented in Figure 4.7.
One-way ANOVA statistical test was used to estimate global differences among the methods. Subsequently, Bonferroni post hoc test was applied for transfection, proliferation, and viability mean values comparison (0.05 significance level). Statistical analysis results are summarized in Table A3.2 in the Appendix. 79
Figure 4.7 (a) Transfection efficiency, (b) proliferation rate, (c) viability and (d) quality factor for HEK 293 cells at various cell seeding density after transfection with Fe2O3@TCP and 87.5 mM CaCl2 (+) or 100 mM CaCl2 ( ++), with PolyMAG, NeuroMag, standard CaP and PEI; # data missing due to NeuroMag fluorescence; * p < 0.0001 versus Fe2O3@TCP
(CaCl2 ++) at the same density (Bonferroni post test).
With PolyMAG transfection efficiencies were comparable to results obtained with
Fe2O3@TCP and 100 mM CaCl2. Conversely, with NeuroMag we achieved significantly higher transfection rates (~60-75%) than with Fe2O3@TCP particles (p < 0.001 at all cell densities). CaP gave significantly better results than Fe2O3@TCP particles in term of transfection efficiency when cells were seeded at 312 and 625 cells/mm2 (p < 0.001 in both cases), while results were comparable when starting cell density was 937 cells/mm2. With
PEI we obtained transfection rates not significantly different from those with Fe2O3@TCP and 100 mM CaCl2.
Transfection efficiency is the key factor indicating how many cells the plasmid was able to enter, but it can be an unfair parameter: if the method significantly affects cell proliferation 80 and physiology, high efficiencies can be obtained since cells remain few in number and, thus, most of them result transfected. Even if not dead, those cells are possibly very unhealthy. Additionally, this translates in low density of transfectants (number of transfectants per surface unit), a disadvantage in many applications such as protein manufacturing. For this reasons we decided to include an additional parameter, i.e. cell proliferation, to rigorously judge how efficient a transfection agent is. An optimal transfection tool should provide simultaneously high transfection efficiency, proliferation rate and viability. The quality factor introduced (Experimental section), taking into account the three equally important variables, is therefore the most natural measure to properly evaluate the overall performance of the different methodologies. If just one of the three variables is very small, the QF value is drastically affected.
As expected, PolyMAG and NeuroMag highly affected cell proliferation, which was significantly lower when compared to results obtained with Fe2O3@TCP particles (p < 0.001 in all cases). Cells incubated with PolyMAG and NeuroMag were rounded, detached and clumped, a clear sign of induced damage (Figure 4.6b and c). Viability of cells treated with PolyMAG was in the range of 64-72% as reported elsewhere,245 and therefore significantly lower (p < 0.0001 at all cell densities) than in samples incubated with Fe2O3@TCP particles. PolyMAG cytotoxicity was most likely associated to the presence of polycations.218 NeuroMag had most probably a similar toxic effect on cells, as judged from the atypical, round morphology. Unfortunately, we could not estimate cell viability in samples transfected with NeuroMag since particles showed a strong red fluorescence which did not allow to distinguish red-fluorescent dead cells stained with ethidium homodimer-1, and represents an obstacle for in vitro fluorescence microscopy investigations. With standard CaP precipitates, proliferation rate was strongly dependent on starting cell density and significantly lower (at 312 and 625 cells/mm2, p < 0.05 in both cases) or comparable (937 cells/mm2) to that with
Fe2O3@TCP particles. Cells were considered generally healthy by morphological examination (Figure 4.6d), with a viability of ≥ 90% at each cell density. Surprisingly, PEI did not show toxic effects. Proliferation rates with PEI were significantly higher (at 312 cells/mm2, p < 0.01) or comparable (at the other two cell densities) to those in samples transfected with Fe2O3@TCP particles. The absence of cytotoxicity was possibly due to the low molecular weight of the PEI chosen, combined with the use of a not too high N:P ratio.246 Morphological inspection of cells transfected with PEI confirmed the healthy status of the cells. Cells were ≥ 99% viable. 81
As the Figure 4.7 illustrates, QF was equally high for Fe2O3@TCP particles, CaP and PEI, while Polymag had a significantly lower score. Therefore, taking into account all the analyzed factors, Fe2O3@TCP particles perform better than PolyMAG, and are generally comparable to CaP and PEI, which are though non-magnetic and therefore cannot be magnetically targeted to a desired site.
Even though the application of a magnetic field gradient did not increased transfection efficiency of Fe2O3@TCP particles (data not shown), it allowed to guide them by magnetic means and induce localized gene expression. We showed the possibility to achieve spatial control of transfection by creating the pattern illustrated in Figure 4.8, where GFP expression is observed only in areas defined by the magnetic gradients applied.
Figure 4.8 (a) Scheme of the experimental set-up used: cells in a Petri dish are placed on a magnetic array, featuring permanent magnets in a plastic mold; (b) phase contrast image illustrating cells equally spread all over the Petri dish, with zoom on non-transfected cells and on transfected ones; (c) GFP fluorescence image proving that Fe2O3@TCP magnetic particles allow localized control of gene expression in areas overlaying magnet positions and in between poles of opposite polarity, leaving not transfected spots between poles of same polarity; zoom on a green area and on a dark spot are also shown.
The magnetic array used to achieve such result is formed by cylindrical permanent magnets inserted into a plastic support in an alternating polarization manner from each adjacent permanent magnet (Figure 4.8a). Due to this arrangement, highest field gradients are produced in the region above the magnet positions, where most of the particle are guided, producing the highest density of transfected cells. Lower magnetic gradients are present between poles of opposite polarity, where a lower density of transfected cells is observed. Transfectant density approximates zero in areas between poles of same polarity repulsing each other (Figure 4.8c). 82
Such a platform allowing site-specific transfection would be suitable for gene screening as well as for engineering of complex tissues in vitro.
4.4 Conclusion
In summary, we designed and produced for the first time Fe2O3@TCP nanoparticles by flame spray synthesis. The intrinsic non-toxicity of the components and the magnetic properties of the composite powder make it a promising candidate for magnetically guided nucleic acid delivery. The particles were successfully used to transfect HEK 293 cells, without compromising cell viability or inducing changes in cell morphology. Fe2O3@TCP nanoparticles were further compared to other transfection agents: two magnetic commercial options (PolyMAG and NeuroMag), standard CaP co-precipitates and PEI. Although highest transfection efficiencies were obtained with NeuroMag, Fe2O3@TCP nanoparticles did not have the negative influence on cell growth and morphology that both commercial magnetic transfection agents induced. Overall Fe2O3@TCP nanoparticle performance, considering together transfection efficiency, proliferation rate, and viability, were similar to that of CaP and PEI. An enhancement with respect to CaP and PEI was achieved thanks to the magnetic properties of the particles, which allowed spatially controlled transfection. These results suggest possible method applications for site-specific gene expression control in synthetic biology and regenerative medicine.
83
5 Submicrometer-sized thermometer particles exploiting selective nucleic acid stability
M. Puddu, G. Mikutis, W. J. Stark and R. N. Grass
Small, 2015, DOI: 10.1002/smll.201502883.
84
5.1 Introduction
Temperature measurements are required in most research areas and everyday life practices.247 While thermometry at macroscale is firmly established, the development of thermometers working at the micro- and nanoscale is an active research field, driven by unsolved scientific and technological challenges. Conventional bulk thermometers are not suitable in areas such as microelectronics, photonics, microfluidics, where a sub-micrometer spatial resolution is required.248-249 Progress in geothermal energy extraction depends on the development of highly thermally and chemically stable nano-geothermometers for underground temperature measurements.250 Similarly, nanosensors able to perform temperature measurements in small or difficult to access spaces are required in many research fields and industrial sectors, ranging from biotechnology and medicine to automotive industry and aeronautics.
Designing a nanoscale thermometer is not only about reducing the size of the measuring device, but requires the use of novel materials and approaches since conventional techniques encounter severe limitations at the small scale, mainly deriving from the changes in material properties from the macro- to the nano-world.249 Several strategies and materials have been proposed for the development of nanoscale thermometers, including temperature sensitive molecules,251-252 polymers,253-255 proteins 256 and various nanostructures (e.g. quantum dots),247, 257-260 as well as different combinations thereof (e.g. nanoparticles functionalized with thermally sensitive entities).261-264 However, most approaches require complex and expensive equipment for temperature read-out, and the nano-thermometers often do not meet the stability requirements or are too invasive or not sensitive enough.247, 265
Nucleic acids offer unique opportunities to the development of nano-thermometry, enhanced by the ease of synthesizing artificial nucleic acid sequences and detecting them by routine bioanalytical techniques.266 Deoxyribonucleic acid (DNA) has been already exploited in the construction of thermometers, mostly based on fluorescence measurements.140, 266-268 Although the use of DNA provides exceptional versatility, these thermometers present some drawbacks such us as low sensitivity or systematic errors due to fluorescence fluctuations.247 Moreover, the measurement involves real-time temperature read-out, which is impossible in some situations, such as underground temperature measurements for geothermal reservoir exploration and monitoring. Lastly, prolonged exposure of naked DNA strands to elevated temperatures results in unwanted DNA degradation and loss of the sensing ability, thus their application is limited to low temperatures and short exposure times. 85
Although DNA damage represents a problematic side effect in most sensing approaches, the extent of damage occurring when heating could be used to quantify the temperature, which has caused it, for the development of a novel class of thermometers. However, the damage should occur in a slow and controlled way, in order to not rapidly lose sensor functionality.
In previous studies, we have developed a technique to improve nucleic acid (both DNA50-52 and RNA53) thermal and chemical stability by encapsulating them within silica and magnetic silica particles (with an iron oxide core). We have shown that the nucleic acids can be quantified by quantitative real-time polymerase chain reaction (qPCR) after particle dissolution using a fluoride buffer. This encapsulation strategy could be used to obtain the desired effect of controlling and slow down DNA degradation for sensing purposes.
However, such sensing set-up would not allow to discriminate DNA loss (= qPCR signal) generated by the heat from DNA loss originating from particle dilution or from DNA damage caused by sources different from heat. To account for these events, the sensing mechanism should employ two distinct nucleic acid molecules: a sensitive sensing sequence, and a reference sequence of higher thermal stability.
DNA, as Nature’s archive, is intrinsically much more resilient than ribonucleic acid (RNA) and its decay is slower at ambient and elevated temperatures.53 Based on this assumption, the natural choice would be to use DNA as reference, and RNA as sensing element.
In the present work the nucleic acid encapsulation within silica was combined to the newly envisioned temperature sensing mechanism to obtain a robust temperature measurement tool, based on the qPCR quantification of the damage occurred as a consequence of the accumulated temperature.
5.2 Experimental section
5.2.1 Nucleic acids
A commercially available RNA (StemMACS eGFP mRNA), purchased from Miltenyibiotec, was used as sensing sequence. An artificial dsDNA (5’-ATT CAT GCG ACA GGG GTA AGA CCA TCA GTA GTA GGG ATA GTG CCA AAC CTC ACT CAC CAC TGC CAA TAA GGG GTC CTT ACC TGA AGA ATA AGT GTC AGC CAG TGT AAC CCG AT-3’, purchased by Microsynth AG) was selected as reference sequence. 86
5.2.2 Iron oxide particle synthesis
52 Fe2O3 particles were synthetized by co-precipitation as reported from Puddu et al. 2014. 4 g
FeCl24H2O (Aldrich, 99%) and 10.8 g FeCl36H2O (Aldrich, 98%) were dissolved in 50 mL distilled water, and were added dropwise to a 1 M NH4OH solution (500 mL) under vigorous stirring. The obtained particles were washed with water and stored at 50 mg/mL.
5.2.3 Encapsulate synthesis
Silica-nucleic acid encapsulates (SiO2-NA-SiO2) and magnetic nucleic acid encapsulates 50-53 (Fe2O3-NA- SiO2) were synthesized following previous developed protocols.
Commercial SiO2 particles (0.136 µm, Microparticles GmbH) or, for the magnetic encapsulates, the produce iron oxide particles, were functionalized with ammonium groups using N-trimethoxysilylpropyl-N,N,N-trimethylammonium chloride (TMAPS, 50% in MeOH, ABCR GmbH). 1 mL of a 50 mg/mL particle suspension in isopropanol was mixed with 10 L of TMAPS, and the mixture was stirred overnight (900 rpm) at room temperature. The functionalized particles were washed 3 times and stored in isopropanol at 50 mg/mL.
1 mL of a solution containing the dsDNA and the mRNA (12 µg/mL DNA, 3 µg/mL RNA) was mixed with 40 µL of SiO2-TMAPS particle suspension or with 15 µL of Fe2O3-TMAPS particle suspension in order to adsorb the nucleic acids onto the particles, further washed three times and redispersed in 0.5 mL water.
1 L of TMAPS was added to the particle dispersion, which was then vortexed before adding 1 L of tetraethoxysilane (TEOS, ≥ 99%, Aldrich). The mixture was stirred (900 rpm) for 4 h at room temperature before adding additional 8 L of TEOS. The reaction was left to run for 4 days under stirring. The particles were then washed twice with water and once with ethanol.
An additional silica layer was grown on the obtained particles via acidic catalyzed Stöber 269 reaction (233 L ethanol, 72 L H20, 22 L TEOS, 5 L of 10 mol/L acetic acid). The reaction mixture was stirred overnight (900 rpm) at room temperature. After that, the particles were washed twice with ethanol, once with water, and redispersed in 100 L water.
5.2.4 Nucleic acid recovery
Encapsulated nucleic acids were recovered using a buffered oxide etch solution obtained by diluting 1:10 a stock solution (NH4FHF/NH4F, 0.23 g NH4FHF + 0.19 g NH4F in 10 mL water), prepared and handled as described in Paunescu et al.51 87
5.2.5 Encapsulate characterization
Morphology of magnetic and non-magnetic encapsulates was investigated by transmission electron microscopy (TEM), using a Tecnai F30 TEM (300 kV, field emission gun, FEI). Scanning transmission electron microscopy (STEM) investigations on magnetic encapsulates were performed on an aberration-corrected HD-2700CS (cold-field emitter; Hitachi), operated at an acceleration potential of 200 kV. An energy-dispersive X-ray spectrometer (EDXS; Gemini system of EDAX) attached to this microscope allowed to record EDX spectra and elemental maps.
Zeta-potential measurements with Zetasizer Nano (Zetasizer Nano, Malvern) allowed to verify core particle functionalization, nucleic acid adsorption and outer silica layer growth (Table A4.2).
Particle size distributions were obtained by an analytical photocentrifuge (dispersion analyser LUMiSizer, LUM GmbH, light source 470 nm). Transmission profiles (1000) were recorded with time intervals of 20 seconds, at a centrifuging speed of 2000 rpm. Data elaboration was -3 performed by SEPView® software (LUM GmbH), using a density of 2.8 g cm for the Fe2O3- 52 NA- SiO2 particles (calculated as in Puddu et al. 2014).
5.2.6 Reactive oxygen species treatment
The silica encapsulate suspension (5 μl) was mixed with a L-ascorbic acid solution (2.5 μl,
20 mM, Acros Organics), H2O2 (12.5 μl, 20 mM, Merck) and a CuCl2 solution (17.5 μl, 500 μM, Fluka). After 10 min, the treatment was stopped by adding EDTA (17.5 μl, 100 mM, Biosolve). 20 μl of the undiluted fluoride buffer was then added to dissolve the particles. The solutions were dialyzed against water (Millipore 0.025 μm VSWP) before qPCR (see Appendix, Figure A4.3).
5.2.7 qPCR and RT-qPCR analysis
After particle dissolution, the nucleic acid solution was directly analyzed without the need for purification.
For DNA analysis by qPCR, following components were mixed together: 10 µL Takyon No Rox SYBR MasterMix dTTP Blue (Eurogentec), 1 µL primer mix stock (containing both forward and reverse primers, 1 µM each), 8 µL water, 1 µL sample. Following primers 88 sequences were used: 5’-ATT CAT GCG ACA GGG GTA AG-3’ (forward primer) and 5’- ATC GGG TTA CAC TGG CTG AC-3’ (reverse primer), purchased from Microsynth AG.
For RNA analysis, reverse transcription and qPCR reaction were combined in a one-step procedure (RT-qPCR). Following components were mixed together: 25 µL Takyon No Rox SYBR MasterMix dTTP Blue (Eurogentec), 5 µL primer mix stock (containing both forward and reverse primers 1 µM each), 0.5 µL additive (Eurogentec), 0.25 µL EuroScript RT H- and RNase Inhibitor (50 u/µLEuroScript RT H-, 20 u/µLRNase Inhibitor, Eurogentec), 18.25 µL RNase-free water, 1 µL sample. Primer sequences for StemMACS eGFP mRNA were generously donated from Miltenyibiotec.
Both qPCR and one-step RT-qPCR reactions were performed with a Roche LightCycler 96, using the following parameters: 48°C for 30 min, 95°C for 5 min, followed by 40 cycles of 15 s at 95°C, and 1 min at 60°C.
5.2.8 Nucleic acid absolute quantification
Nucleic acid concentration was determined by interpolation from standard curves (see Appendix, Figure A4.4 and Figure A4.5), generated from RNA and DNA dilution series of known concentration (measured respectively by Qubit RNA HS and Qubit dsDNA HS assay, used with Qubit fluorometer, Invitrogen).
5.2.9 Temperature tests
Particle stocks were diluted 1:20 in tap water, achieving a concentration of 1 mg/mL and 0.5 mg/mL, respectively for the silica encapsulates and the magnetic silica encapsulates. 50 µL of the diluted suspensions were pipetted in a RNase-free Eppendorf tubes. Prepared samples were incubate (with closed lid) at the desired temperature in a Thermomixer (Eppendorf). After a given time, the samples were removed and the particles were collected by centrifugation (3 min at 20000 g) or by magnetic means, using a magnetic separator. Then 100 µL of 1:10 diluted fluoride buffer was added and qPCR/RT-qPCR was performed.
5.2.10 Kinetic studies
For the kinetics studies, two samples were analyzed at each time point, and assayed in duplicate by qPCR/RT-qPCR (n = 4). The logarithm of the concentration measured by qPCR/RT-qPCR was fitted versus the time, and linear regression analysis was applied to 89 determine the decay rates at the various temperature (Figure A4.6 and Table A4.3), according to first-order kinetics.53, 92, 270
5.2.11 Calibration curves
To obtain the calibration curves in Figure 5.4 of the manuscript, five samples were analyzed for each temperature, each assayed in triplicates by qPCR/RT-qPCR (n = 15). For each sample, the difference between the cycle threshold of RNA and DNA was calculated
(ΔCt = Ct RNA- Ct DNA).
5.3 Results and discussion
The overall design of the sub-micrometer size thermometer and the key synthetic steps are illustrated in Figure 5.1.
Figure 5.1 Sensor synthesis and working principle: a sensitive sensing sequence (RNA), and a more stable reference nucleic acid sequence (DNA) are encapsulated within silica, which is eventually exposed to a heat source, and further dissolved for the read-out of RNA and DNA damage.
Silica particles, loaded with an artificial double stranded DNA sequence and a commercially available RNA, were synthesized as previously described (see Experimental section). An additional silica layer was grown on the particles surface (see Experimental section) in order to avoid potential nucleic acid deprotection events, arising from increased silica solubility in water at high temperatures. The obtained particles were characterized by transmission electron microscopy (Figure 5.2), and in terms of particle size distribution, zeta-potential, nucleic acid loading (see Appendix A.4). 90
Figure 5.2 a) TEM image of silica encapsulates, scale bar = 50 nm; b) STEM image of magnetic silica encapsulates and c) elemental mapping of Si and Fe, scale bar = 40 nm.
Additionally, resistance of the encapsulated DNA and RNA against reactive oxygen species (ROS) was tested to validate the chemical stability of the encapsulates (Appendix, Figure A4.3). The sensing mechanism of the produced particles, with an average hydrodynamic size in water of 251 nm (Appendix, figure A4.1), is schematically depicted in Figure 5.3.
Figure 5.3 Sensing mechanism schematic: the encapsulates are exposed to a given temperature for a given time, inducing nucleic acid damage. After the event, RNA and DNA are analyzed by qPCR; the difference in qPCR readings of RNA and DNA (cycle threshold difference ΔCt = Ct RNA - Ct DNA) is calculated to quantify the damage produced by the heating, and correlated to the temperature which has produced it. 91
The exposure of the particles to elevated temperature significantly affects the RNA (sensing sequence), but only marginally the DNA (reference sequence). The occurred damage is then quantified by qPCR - expressed as a difference between the cycle threshold Ct of RNA and
DNA (ΔCt), which reflects the difference in concentration between the two - and can be correlated to the temperature which has produced it.
As opposed to previously developed DNA-based temperature probes, here the use of qPCR for the temperature read-out is introduced, a simple and inexpensive method widely used in biology, medical diagnostics and forensic. qPCR not only allows for a precise damage quantification, but also permits to detect nucleic acids at a molecular level with high specificity, thereby increasing the accuracy and the sensitivity of the method.
In a standard procedure, the synthetized encapsulates were dispersed in tap water at a concentration of 1 mg/mL, and exposed to a given temperature (see Experimental section). First, the kinetics of DNA and RNA decay within the silica particles was studied at different temperatures, in order to confirm the applicability of the hypothesized sensing mechanism, and investigate the optimal temperature range of the thermometer. Samples were exposed to temperatures ranging from 50°C to 90°C with 10°C increments. At different time points, the encapsulates were recollected by centrifugation, dissolved with a fluoride comprising buffer (0.25 wt% fluoride, releasing unharmed DNA and RNA), and directly quantified, using qPCR in the case of DNA, and real-time quantitative reverse transcription PCR (RT-qPCR) for RNA. The same thermal cycling protocol was used for DNA and RNA in order to allow for simultaneous DNA and RNA quantification in a single run. As previously reported in literature, a first-order nucleic acid decay kinetics was observed.53, 92, 270 Additionally, as predicted, RNA decayed faster than DNA at all the examined temperatures (Figure A4.6 in the Appendix). In Figure A4.7 in the Appendix, the ΔCt is reported versus time for the various temperatures, exhibiting the expected linearity. By using the obtained curves, and knowing the temperature at which the sample has been exposed to, it is therefore possible to determine the exposure time.
To show the applicability of the produced particles for temperature determination, we then constructed a calibration curve, by heating the encapsulates at various temperatures for a fixed time. The silica encapsulates were exposed to temperatures ranging from 60°C to 85°C with 5°C increments for 30 min prior to analysis by qPCR, obtaining the curve illustrated in Figure 5.4a. 92
Figure 5.4 Sensor calibration curves, obtained by exposing a) silica and b) magnetic silica encapsulates to temperatures within the range 50-80°C for 30 min, showing significant ΔCt value differences (two-sample t-test) between samples exposed at different temperatures: *) p < 0.05; **) p < 0.01.
All the samples showed a clear dependence of the ΔCt (and hence of the RNA and DNA concentration) on the applied temperature. The last data point at 85°C was not included, as the cycle threshold difference of RNA samples and RNA negative controls was too small (< 5). To statistically analyse the results, two-sample t-tests were carried out, showing that mean
ΔCt values obtained at temperatures at 5°C increments between 65°C and 80°C were significantly different (p < 0.05 in all cases), if averages were taken from five independent samples (Figure 5.4a). By using such calibration curves, it is possible to relate the encapsulated nucleic acid concentration to the temperature they have experienced, and, therefore, it proves the successful development of a novel sub-micrometer size thermometer. As compared to previous DNA-based thermometers, the present technology permits to measure generally higher temperatures as a result of the nucleic acid encapsulation. The temperature range of the developed probe is determined by the nucleic acid stability: the lower limit is represented by the temperature at which no degradation occurs, while the upper limit is fixed by the point where near to complete RNA degradation occurs. It is envisioned that by using nucleic acid sequences of different stability, the temperature range can be extended or shifted towards lower or higher temperatures (within the restrictions posed by natural nucleic acid decay). The accuracy instead depends on the number of samples analysed: a better accuracy would be possible if more independent samples were averaged, 93 even though it is limited by the intrinsic logarithmic scale of the qPCR method. The encapsulation of the nucleic acids, significantly slowing down their natural decay, ensures long-term data storage92 and sensor durability. Therefore, such thermometer can record the temperature it was exposed to and be read later on, even long time after the thermal event. Furthermore, the encapsulation allows to confine the nucleic acids within a sub-micrometer carrier which can be easily delivered and recovered. Since the system integrates multiple functions (measurement, data storage, transport) within a particle, it can be referred to as a “lab-in-a-particle”. This feature, combined with the thermal and chemical stability of the encapsulation material (silica), makes these particles suitable sensing tools in a range of challenging industrial, environmental and research settings.
Depending on the systems where the temperature measurement is conducted, sensor recollection by centrifugation could be problematic. The use of magnetic nucleic acid encapsulates as temperature sensors would facilitate the handling and would permit easy sample up-concentration before the analysis. Also, it would allow to sample bigger volumes, increasing the amount of collected encapsulates and thereby reduce the starting particle quantity required for efficient read-out. To exploit this potential, we encapsulated the DNA and RNA sequences within silica particles with an iron oxide core, following a previously developed protocol, and protected them with an additional silica layer. The produced magnetic nucleic acid encapsulates (Figure 5.1b and c) were thoroughly characterized (see Appendix A.4) and assessed for their thermo-sensitivity. The particles, having an average hydrodynamic size of 478 nm, were dispersed in water at 0.5 mg/mL and exposed for 30 min at temperature ranging from 50°C to 75°C, with 5°C increments. The encapsulates were then separated by magnetic means and dissolved by a fluoride buffer (releasing DNA and RNA) prior to analysis by qPCR, obtaining the calibration curve in Figure 4b. The temperature range where the thermometer is functional was identified as 50°C-70°C. We were able to statistically discriminate exposure temperatures between 55°C and 70°C at 5°C increments to a certainty of 95% (p < 0.05). These results show the development of a magnetically recoverable nucleic acid-based thermometer.
Because of the presence of the nucleic acids, such particles can be detected at trace levels (< ppb), being optimal tracers. This unique feature makes these objects capable of simultaneous flow tracing and temperature measurement, opening up a variety of applications, ranging from quality control, to industrial fluid monitoring or temperature and flow mapping within underground reservoirs. There, the sub-micrometer sized sensor may serve to measure 94 localized temperatures or to perform continuous distributed measurement over thousands of points simultaneously. The broadly recognized biocompatibility of constituent materials, which are approved food additives (amorphous silica = E551, iron oxide = E172), could facilitate their use within the food or food packaging industries.
5.4 Conclusion
In summary, this work demonstrates for the first time the feasibility of using nucleic acid damage quantification by qPCR to measure temperature, by employing a novel design, consisting of a RNA sensing sequence and a DNA reference sequence. With increasing temperature, the sensing sequence is degraded, while the reference is only partially damaged, and therefore allows to account for non-thermal damage and particle loss. The sensor design provides excellent stability and versatility, and warrants its application in several industrial and technological setting for the investigation of temperature at the sub-micrometer scale.
95
6 Conclusion and outlook
96
To summarise, different nucleic acid/inorganic particle hybrids were assembled and some of their applications were presented in the previous chapters. A way to encapsulate nucleic acids (both DNA and RNA) within silica and magnetic silica particles with an iron oxide core was reported. It has been shown how the encapsulated nucleic acids possess superior thermal and chemical stability as compared to naked nucleic acids. This allowed the development of a novel method to handle, store and ship RNA samples at room temperature for use in medical laboratories and life science research. The preservation strategy represents a promising alternative to current methodologies, which mostly rely on the use of freezers and low temperatures, and are therefore impractical, costly, time-sensitive and take up space and energy.
DNA encapsulates with magnetic properties were tested as tags for in-product labeling of oil products, in order to protect them against counterfeit. The developed platform, requiring trace levels of the tags and allowing the quantification of the tag concentration on a logarithmic scale, enables to discriminate adulterated products (e.g. fake, diluted items) and to determine oil origin/legitimacy. The procedure can be applied to tag/trace any liquid product or bulk fluid in many industrial or environmental settings. Many industries (e.g underground energy extraction and chemical industries, nuclear power plants, wastewater treatment plants, refineries, aeronautic and rail sectors) could benefit from highly stable and secure tags enabling the detection of leakages and the determination of fluid flow rates or ensuring authenticity and supply chain security. Similarly, their use as environmental tracers would bring several advantages in comparison to traditional tracers, not only in term of stability, but also in term of complexity of studies which could be performed (e.g. multi-tracer tests using several different DNA sequences at the same time). In this regard, the DNA-silica hybrid particles are currently being tested for multi-tracer studies in surface and subsurface hydrology, in order to obtain information on solute transport, travel times, and flow pathways, showing promising preliminary results. Similarly, the particles are soon going to be used to perform underground connectivity studies, in order to assess their suitability as tracers for geothermal reservoir characterization.
The geothermal energy extraction sector would also profit from thermosensitive tracers able to measure temperature beyond the wellbore region and within fractured rocks, since currently there is no way of doing this. To this end, the sub-micrometer sized thermometer reported in Chapter 5 would represent an ideal candidate. The sensor is based on the quantification by real-time polymerase chain reaction of the encapsulated nucleic damage produced by a 97 temperature increase. Because of its design, the system allows for simultaneous tracing and sensing. Additionally, the hybrid particles can perform measurement in small/difficult to access spaces where traditional sensing is prevented due to sensor size, as within rock fractures. However, the temperature spatial mapping of underground reservoir requires particles which can withstand the high temperatures into the formation (up to 200°C). Therefore, for such application, effort would be required in extending the temperature stability and temperature measurement range of the particles.
The developed sensing mechanism can be generalized and utilized to measure other properties than temperature such us light, oxidative stress, pH, humidity. The combination of particle and nucleic acid technology provides not just functionality but also extreme versatility, since both the materials can be tuned to achieve the optimal sensitivity towards the property to be measured and to meet the requirements for the specific system where the measurement is performed. It has been shown that by modifying DNA sequences with light-sensitive groups, a light sensor is obtained. It is envisioned that by synthesizing a porous silica layer which allows chemical species (e.g. reactive oxygen species, heavy metals ions) to diffuse to the nucleic acid compartment, a sensor able to detect such entities could be produced.
Lastly, magnetic calcium phosphate particles were designed as optimal transfection vehicles and produced for the first time by flame spray synthesis. The material, able to bind DNA in the presence of CaCl2, was tested for the transfection of a green fluorescent protein encoding plasmid with Human Embryonic Kidney 293 cells. It was shown that the nanoparticles are improved tools to deliver nucleic acids into cells and achieve spatial control of transfection, suggesting possible application in synthetic biology and regenerative medicine. Currently, the developed formulation is being tested for the transfection of HeLa cells and neuronal cell lines, exhibiting good performance.
The thesis shows how the fusion of nucleic acid technology and particle technology has not only led to significant progresses in traditional application fields of nucleic acids and particles, but has additionally opened up new opportunities. The combination of these biomolecules with particles has allowed the design and manufacturing of hybrid materials with new properties to address not only bio-medical issues but also other technological problems. By tuning the inorganic component, it has been possible to improve the properties of the biomolecule, such us nucleic acid thermal and chemical stability. Similarly, the possibility to easily purchase nucleic acids with the desired sequence and modifications has facilitated their coupling with the inorganic counterpart and has permitted to provide specific 98 function to the hybrid (e.g. sensing capability). Additionally, the development of tools and platforms based on such hybrid materials has largely benefited from the refinement of bio- analytical techniques (e.g. polymerase chain reaction) originally developed for molecular biology. The implementation of such techniques within sensing, tracing and other fields non- related to molecular biology or medicine, has allowed the development of novel advanced technological platforms, and their continuous improvement and easier access promise exciting future developments.
99
Appendix
100
A.1 Supporting information to chapter 2
A.1.1 Particle density
Magnetization data suggested that the produced material is 60 wt% SiO2 and 40 wt% Fe2O3. We estimated material density accordingly:
1 = 푚푖 ∑푖 푖 where i is the density of constituent i (silica, iron oxide), and mi the mass fraction.
3 3 SiO2 density: 2.6 g/cm Fe2O3 density: 5.6 g/cm
Calculated particle density: 3.3 g/cm3
A.1.2 Particle sedimentation velocity
Particle sedimentation velocity was calculated using the Stoke’s law:
2 (푝 − 푓) 푔 푑 푣 = 푠 18 with p and f densities respectively of the particles and of the fluid, dynamic viscosity of the fluid, g gravitational acceleration, and d particle diameter. The average hydrodynamic diameter obtained with the LUMiSizer and the calculated particle density were used. Density and viscosity values are listed in the below table.
Table A1.1 Density, dynamic viscosity values and calculated particle sedimentation velocities in toluene, gasoline, limonene and olive oil.
Density (Kg/m3)* Viscosity (Pas)* Sedimentation velocity (m/s) Toluene 865 0.00056 410-8 Gasoline 749 0.00045 810-8 Limonene 842 0.00092 410-8 Olive Oil 910 0.084 410-10 *Values taken from product data sheets and “Food Processing Technology: Principles and Practice”.
101
A.1.3 Particle aggregation time
At the extremely low concentrations of particles in the suspensions, aggregation is slow. Aggregation depends on concentration as described in the Smoluchowski equation:
푑푛 = −훽푛2 푑푡 where n is the particle concentration and β is the aggregation rate, which can be simplified for a monomodal solution to:
8푘 푇 훽 = 푏 µ
The characteristic time of aggregation (average time for two particles to meet in solution) is, therefore:
3µ 휏푎푔푔 = 4푘푏푇푛0 with µ viscosity, kb Boltzmann constant, T tempearature, n0 initial particle concentration (#/m3).271
A.1.4 Independent two-sample t-test
푋̅ − 푋̅ 푡 = 1 2 2 푠 ∙ √ 푋1푋2 푛
where
1 푠 = √ (푠2 + 푠2 ) 푋1푋2 2 푋1 푋2
X1, X2 and SX1, SX2 are, respectively, means and standard deviations of sample group 1 and 2. The degree of freedom for this test is 2n – 2.
102
Table A1.2 X1-X2, SX1X2, n and t values for bergamot oil, gasoline, and olive oil.
Bergamot Oil
Group 1 vs Group 2 X1-X2 = ΔCt SX1X2 n t 1 µg/L vs blank 6.2 1.6 5 5.989 10 µg/L vs 1 µg/L 2.8 1.8 5 2.434 100 µg/L vs 10 µg/L 3.8 1.7 5 3.530 Gasoline
Group 1 vs Group 2 X1-X2 = ΔCt SX1X2 n t 1 µg/L vs blank 4.7 1.2 5 6.411 10 µg/L vs 1 µg/L 4.3 1.1 5 6.488 100 µg/L vs 1 µg/L 5.3 0.8 5 9.843 Olive Oil
Group 1 vs Group 2 X1-X2 = ΔCt SX1X2 n t 1 µg/L vs blank 5.3 1.6 5 5.232 10 µg/L vs 1 µg/L 2.5 1.8 5 2.257 100 µg/L vs 1 µg/L 4.0 1.4 5 4.497
103
Table A1.3 Cost calculation to produce 1 g of DNA encapsulates.
Working step FeCl24H2O and TEOS TMAPS DNA C6 silane
FeCl36H2O (50% in Methanol)
Fe2O3 synthesis 0.3 g and 0.8 g Ammonium group 10 mL functionalization DNA adsorption 16 mg
SiO2 coating 9 mL 1 mL
C6 functionalization 100 mL Price (USD)* 0.9 1.0 14.1 160.0 23.8
*All basic chemical costs were taken from the corresponding material suppliers used in the methods section of the paper (lab-scale pricing) without optimizing the synthesis procedures. For the cost of DNA we considered a commercial offer for the synthesis of 1 g custom DNA from Microsynth AG (Switerland).
~ 200 USD are needed for production of 1 g of DNA encapsulates. Consequently, the cost of tagging 1 L of oil with 1 g DNA encapsulates (= 1 ppb particle concentration in oil) is 210-4 USD. This price is negligible compared to cost of barcoded oils: gasoline: 0.9 - 2 USD/L (U.S. Energy Information Administration, October 2013 - Europe’s Energy Portal, December 2013); extra virgin olive oil: 4 - 20 USD/L (IMF Primary Commodity Prices, October 2013 - branded product); bergamot oil: 35 - 70 USD/L (taken from www.alaibaba.com, December 2013 - branded product).
104
Figure A1.1 TEM micrographs of pristine Fe2O3 and of Fe2O3/TMAPS/DNA/SiO2 particles.
Figure A1.2 Standard curve used for DNA quantification. Slope and R values are provided.
Figure A1.3 Simulated long-term stability of encapsulated DNA in decalin at RT following ASTM F1980 (Accelerated Aging of Sterile Medical Device Packages, Q10=2, T=65°C). 105
Figure A1.4 Particle size distributions of Fe2O3/TMAPS/DNA/SiO2-C6 particles in toluene (dash), gasoline (short dash), limonene (dash dot).
106
A.2 Supporting information to chapter 3
A.2.1 Kinetics
The decay kinetics of solid-state RNA can be best expressed by the equation: 92, 270
dC − = kC dt where C is the RNA concentration and k is the decay rate constant, which integrated gives:
lnC = −kt + lnC0
To document sample decay during accelerated aging studies, RT-qPCR was used. Obtained Cq values were used to calculate specimen concentration using the previously produced standard curve (Figure A2.3).
The logarithm of the RNA concentrations measured by RT-qPCR was fitted versus the time and linear regression analysis was applied to determine the decay rates at 65°C (Table A2.1) according to the equation above.
The degradation kinetic constant for a given temperature kT was calculated with the Arrhenius’ equation: 92, 270
─EA kT = Ae RT
198 using an activation energy (EA) of 119 kJ/mol.
270 RNA molecule half-life (t1/2) at a certain temperature was calculated with the formula:
ln2 t = 1/2 k where k is the rate constant. Assuming that RNA degrades by the same phosphodiester bond break mechanism as in solution,198 phosphodiester bond half-lives were obtained by multiplying the molecular half-life by the number of nucleotides in a RNA molecule (in this case nt = 847).
107
A.2.2 Simplified procedure for RT-qPCR
In order to simplify the synthesis and analysis procedure for RT-qPCR we investigated if the encapsulated samples required a washing step after 4 days of synthesis, and if the usually applied dialysis step in RNA work-up could be replaced by a simple dilution step (diluting 2- RNA but also diluting contaminants such as SiF6 ). As shown in Figure A2.5 the absence of the washing step did not make any difference in performance, and the relative concentrations of RNA after dialysis and simple dilution were equivalent.
Table A2.1 Linear fit of concentration data to first order decay rate equation.
k 65° (s-1) R2 Dried RNA 2 10-5 0.97 RNAstable 2 10-5 0.96 -6 SiO2/RNA microcapsules 2 10 0.96
Figure A2.1 a) TEM micrograph of SiO2/RNA microcapsules; STEM pictures of b) the original silica particles and of c) the particles after the growth of the protective silica layer. 108
Figure A2.2 Gel-electrophoresis of the original mRNA (left) and after the encapsulation/release process (right).
Figure A2.3 StemMACS eGFP mRNA standard curve.
Figure A2.4 Bovine chondrocyte RNA standard curve. 109
Figure A2.5 Produced microcapsule suspension (500 µL, ~4 mg/mL) was washed and resuspended in 100 µL after 4 day synthesis; or no sample work-up was performed (unwashed). Equivalent amounts of the unwashed and washed particles were dissolved with fluoride buffer solution (20 µL, 2.5 F─ wt %), and further dialyzed against water or diluted 1:10 or 1:100 prior to RT-qPCR analysis. Data are shown as mean concentration ± standard deviation (n = 4).
110
A.3 Supporting information to chapter 4
A.3.1 Particle density
We estimated material density knowing that the produced composite material is 67 wt% TCP and 33 wt% Fe2O3:
1 = 푚푖 ∑푖 푖
3 3 where i is the density of constituent i (TCP: 3.14 g/cm , Fe2O3: 5.24 g/cm ), and mi the mass fraction. Calculated particle density was 3.6 g/cm3.
A.3.2 Primary particle size
The average primary particle diameter (dBET) was calculated from the measured specific surface area As and the density ρ using:
푑퐵퐸푇 = 6/(퐴푠 · 휌)
Obtained particle diameter was 25 nm.
A.3.3 Particle size dispersity
Particle size distribution data obtained by XDC were fitted with a lognormal distribution (Figure A3.1) to estimate the mean diameter and the geometric standard deviation (GSD). We obtained a mean diameter of 121 nm and a GSD of 1.2, indicating a sufficiently monodisperse distribution.
A.3.4 Number of particles per aggregate
Average number of primary particles per aggregate (np) was calculated using the following relationship:
퐷 푑푐 푛푝 = ( ) 푑푝 using dBET as primary particle diameter (dp), mean hydrodynamic size measured by XDC as 272 collision diameter (dc), and a constant D of 1.8.
111
A.3.5 Particle long-term storage
Fe2O3@TCP nanoparticles can be stored as prepared for long periods. The dry storage does not affect particle size and ability to bind pDNA: both quantities were measured again after storage at ambient temperature for 8 months, attaining the same results previously obtained (i.e. a BET particle diameter of 26 nm - BET gives an error ~5% - , and a binding capacity of 0.02 µg DNA/µg particles, as measured by Qubit fluorometer and visualized on the gel in Figure A3.2). An additional proof that particle size is not affected by long-term storage is that a BET particle diameter of 26 nm was measured after sterilization for 30 min at 200°C, a process equivalent to storage at room-temperature for about 10 years according to ASTM F1980 (Accelerated Aging of Sterile Medical Device Packages, Q10=2).
112
Table A3.1 Cell counting data.
Cell density: 312 cells/mm2 625 cells/mm2 937 cells/mm2 Blue Red Green Blue Red Green Blue Red Green 3887 110 1294 2088 178 576 2662 237 700
Fe2O3@TCP 3913 133 1248 4271 269 1200 5874 217 1582
(CaCl2 +) 3262 92 1255 4567 174 1277 4392 244 1278 3914 62 1288 6396 247 1731 7022 203 1759
Fe2O3@TCP 3756 93 1305 5623 271 1322 7328 217 1731
(CaCl2 ++) 3278 58 1034 5520 243 1348 7659 145 1799 1075 373 240 2492 1111 586 3468 1180 933 PolyMAG 1007 311 197 2228 1344 660 3378 1322 837 845 250 137 1624 1126 414 2267 1056 633 354 - 227 1559 - 932 1670 - 1236 NeuroMag 568 - 378 2485 - 1511 2869 - 2186 684 - 537 2303 - 1360 2637 - 1995 1110 74 864 4138 362 1811 6552 291 1530 CaP 1538 84 948 3746 442 1514 6152 423 1754 1287 122 964 1945 284 753 6565 251 1871 5169 39 1544 7638 28 1382 7911 28 1628 PEI 5135 57 1798 6763 39 1799 8195 39 1901 6941 16 2224 6618 22 1724 7388 16 1570 6709 1 0 8159 13 0 8366 8 0 Neg. ctr. 6620 4 0 8020 5 0 8565 5 0 6799 1 0 8054 6 0 8794 4 0
113
Table A3.2 Results of one-way ANOVA followed by Bonferroni post-test, comparing transfection, proliferation, and viability mean values obtained with Fe2O3@TCP particles
(100 mM CaCl2) and the other reagents.
Cell density: 312 cells/mm2 625 cells/mm2 937 cells/mm2 Transfection Prob. Sig. Prob. Sig. Prob. Sig.
Fe2O3@TCP vs. PolyMAG 1E-1 NS* 1 NS 1 NS
Fe2O3@TCP vs. NeuroMag 1E-4 S** 5E-7 S 7E-11 S
Fe2O3@TCP vs. CaP 7E-5 S 6E-4 S 8E-1 NS
Fe2O3@TCP vs. PEI 1 NS 1 NS 1 NS Proliferation Prob. Sig. Prob. Sig. Prob. Sig.
Fe2O3@TCP vs. PolyMAG 7E-04 S 5E-04 S 7E-06 S
Fe2O3@TCP vs. NeuroMag 2E-04 S 6E-04 S 2E-06 S
Fe2O3@TCP vs. CaP 2E-03 S 1E-2 S 4E-1 NS
Fe2O3@TCP vs. PEI 5E-03 S 6E-1 NS 1 NS Viability Prob. Sig. Prob. Sig. Prob. Sig.
Fe2O3@TCP vs. PolyMAG 1E-7 S 4E-6 S 6E-7 S
Fe2O3@TCP vs. NeuroMag ------
Fe2O3@TCP vs. CaP 3E-2 S 2E-1 NS 9E-1 NS
Fe2O3@TCP vs. PEI 1 NS 9E-1 NS 1 NS *NS = the means difference is non significant at the 0.05 level **S = the means difference is significant at the 0.05 level
114
Figure A3.1 Hydrodynamic size distribution of Fe2O3@TCP particles.
Figure A3.2 Gel electrophoresis showing DNA Ladder (1), control DNA solution (2) not treated with nanoparticles, and residual DNA in the supernatant treated with nanoparticles (3) as prepared (left) and after storage for 8 months at room temperature (right).
Figure A3.3 Fluorescence microscopy picture (left) showing Hoechst stained cells (blue), EthD-1 stained cells (red), and cells expressing GFP (green); merged image (right) of green, red, and bright field, showing cell morphology. Scale bar: 100 µm. 115
A.4 Supporting information to chapter 5
Table A4.1 Particle capacity.
µg DNA/mg particles µg RNA/mg particles
SiO2-NA-SiO2 0.1 0.1
Fe2O3-NA- SiO2 0.1 0.01
Table A4.2 Zeta-potential data.
Zeta-potential (mV)
SiO2 -38
SiO2-TMAPS 58
SiO2- TMAPS-NA -34
SiO2- TMAPS-NA-SiO2 -18
Fe2O3 0
Fe2O3-TMAPS 37
Fe2O3-TMAPS-NA -32
Fe2O3-TMAPS-NA-SiO2 -27
Table A4.3 Linear fit of concentration data to first order decay rate equation.
k DNA (min-1) k RNA (min-1) 50°C 9 10-5 0.002 60°C 8 10-4 0.01 70°C 0.007 0.07 80°C 0.004 0.1 90°C 0.02 0.2
116
Figure A4.1 Size distributions of SiO2-NA-SiO2 and Fe2O3-NA-SiO2 particles in water.
Figure A4.2 Hysteresis loops of Fe2O3 and Fe2O3-NA- SiO2 particles.
Figure A4.3 Stability against reactive oxygen species (ROS) of SiO2-NA-SiO2 particles. 117
Figure A4.4 dsDNA standard curve.
Figure A4.5 mRNA standard curve.
118
Figure A4.6 Logarithm of the DNA and RNA concentration measured by qPCR/RT-qPCR fitted versus the exposure time at temperatures ranging from 50°C to 90°C. Data are provided as mean logarithm of the concentration ± standard deviation.
Figure A4.7 Difference between the cycle threshold of RNA and DNA (ΔCt = Ct RNA- Ct DNA) versus the exposure time at temperatures ranging from 50°C to 90°C. Data are provided as mean ΔCt value ± standard deviation.
119
References 1. Watson, J. D.; Crick, F. H. C. Molecular Structure of Nucleic Acids - a Sturcture for Deoxyribose Nucleic Acid. Nature 1953, 171, 737-738.
2. D. Nelkin, M. S. L., The DNA Mystique: The Gene as a Cultural Icon. University of Michigan Press: United States, 2004.
3. Kemp, M. The Mona Lisa of Modern Science. Nature 2003, 421, 416-420.
4. Moen, I.; Jevne, C.; Wang, J.; Kalland, K. H.; Chekenya, M.; Akslen, L. A.; Sleire, L.; Enger, P. O.; Reed, R. K.; Oyan, A. M.; Stuhr, L. E. B. Gene Expression in Tumor Cells and Stroma in Dsred 4t1 Tumors in Egfp-Expressing Mice with and without Enhanced Oxygenation. BMC Cancer 2012, 12, 10.
5. Hebert, P. D. N.; Penton, E. H.; Burns, J. M.; Janzen, D. H.; Hallwachs, W. Ten Species in One: DNA Barcoding Reveals Cryptic Species in the Neotropical Skipper Butterfly Astraptes Fulgerator. Proc. Natl. Acad. Sci. U. S. A. 2004, 101, 14812-14817.
6. Ye, X.; Al-Babili, S.; Klöti, A.; Zhang, J.; Lucca, P.; Beyer, P.; Potrykus, I. Engineering the Provitamin a (Β-Carotene) Biosynthetic Pathway into (Carotenoid-Free) Rice Endosperm. Science 2000, 287, 303-305.
7. Davis, J. Microvenus. Art Journal 1996, 55, 70-74.
8. C. Gorman, D. F. M., The Rna Revolution. Scientific American 2014.
9. Seeman, N. C. Structural DNA Nanotechnology: Growing Along with Nano Letters. Nano Lett. 2010, 10, 1971-1978.
10. Seeman, N. C. DNA in a Material World. Nature 2003, 421, 427-431.
11. Pinheiro, A. V.; Han, D.; Shih, W. M.; Yan, H. Challenges and Opportunities for Structural DNA Nanotechnology. Nat. Nanotechnol. 2011, 6, 763-772.
12. Lu, C.-H.; Willner, B.; Willner, I. DNA Nanotechnology: From Sensing and DNA Machines to Drug-Delivery Systems. ACS Nano 2013, 7, 8320-8332.
13. Seeman, N. C. Nucleic-Acid Junctions and Lattices. J. Theor. Biol. 1982, 99, 237-247.
14. Junghuei, C.; Seeman, N. C. Synthesis from DNA of a Molecule with the Connectivity of a Cube. Nature 1991, 350, 631-633.
15. Winfree, E.; Liu, F. R.; Wenzler, L. A.; Seeman, N. C. Design and Self-Assembly of Two-Dimensional DNA Crystals. Nature 1998, 394, 539-544.
16. Zhang, Y. W.; Seeman, N. C. Construction of a DNA-Truncated Octahedron. J. Am. Chem. Soc. 1994, 116, 1661-1669.
17. Park, S. H.; Pistol, C.; Ahn, S. J.; Reif, J. H.; Lebeck, A. R.; Dwyer, C.; LaBean, T. H. Finite-Size, Fully Addressable DNA Tile Lattices Formed by Hierarchical Assembly Procedures. Angew. Chem.-Int. Edit. 2006, 45, 735-739. 120
18. Ke, Y.; Ong, L. L.; Shih, W. M.; Yin, P. Three-Dimensional Structures Self- Assembled from DNA Bricks. Science 2012, 338, 1177-1183.
19. Wei, B.; Dai, M.; Yin, P. Complex Shapes Self-Assembled from Single-Stranded DNA Tiles. Nature 2012, 485, 623-626.
20. Zheng, J.; Birktoft, J. J.; Chen, Y.; Wang, T.; Sha, R.; Constantinou, P. E.; Ginell, S. L.; Mao, C.; Seeman, N. C. From Molecular to Macroscopic Via the Rational Design of a Self-Assembled 3d DNA Crystal. 2009, 461, 74-77.
21. Rothemund, P. W. K. Folding DNA to Create Nanoscale Shapes and Patterns. Nature 2006, 440, 297-302.
22. Guo, P. X. The Emerging Field of Rna Nanotechnology. Nat. Nanotechnol. 2010, 5, 833-842.
23. Chworos, A.; Severcan, I.; Koyfman, A. Y.; Weinkam, P.; Oroudjev, E.; Hansma, H. G.; Jaeger, L. Building Programmable Jigsaw Puzzles with Rna. Science 2004, 306, 2068- 2072.
24. Delebecque, C. J.; Lindner, A. B.; Silver, P. A.; Aldaye, F. A. Organization of Intracellular Reactions with Rationally Designed Rna Assemblies. Science 2011, 333, 470- 474.
25. Fischler, M.; Sologubenko, A.; Mayer, J.; Clever, G.; Burley, G.; Gierlich, J.; Carell, T.; Simon, U. Chain-Like Assembly of Gold Nanoparticles on Artificial DNA Templates Via 'Click Chemistry'. Chem. Commun. 2008, 169-171.
26. Cutler, J. I.; Zheng, D.; Xu, X. Y.; Giljohann, D. A.; Mirkin, C. A. Polyvalent Oligonucleotide Iron Oxide Nanoparticle "Click" Conjugates. Nano Lett. 2010, 10, 1477- 1480.
27. Wang, Y. F.; Wang, Y.; Zheng, X. L.; Ducrot, E.; Lee, M. G.; Yi, G. R.; Weck, M.; Pine, D. J. Synthetic Strategies toward DNA-Coated Colloids That Crystallize. J. Am. Chem. Soc. 2015, 137, 10760-10766.
28. Herr, J. K.; Smith, J. E.; Medley, C. D.; Shangguan, D. H.; Tan, W. H. Aptamer- Conjugated Nanoparticles for Selective Collection and Detection of Cancer Cells. Anal. Chem. 2006, 78, 2918-2924.
29. Bagalkot, V.; Zhang, L.; Levy-Nissenbaum, E.; Jon, S.; Kantoff, P. W.; Langer, R.; Farokhzad, O. C. Quantum Dot - Aptamer Conjugates for Synchronous Cancer Imaging, Therapy, and Sensing of Drug Delivery Based on Bi-Fluorescence Resonance Energy Transfer. Nano Lett. 2007, 7, 3065-3070.
30. Day, P. J. R.; Flora, P. S.; Fox, J. E.; Walker, M. R. Immobilization of Polynucleotides on Magnetic Particles - Factors Influencing Hybridization Efficiency. Biochem. J. 1991, 278, 735-740.
31. Hilliard, L. R.; Zhao, X. J.; Tan, W. H. Immobilization of Oligonucleotides onto Silica Nanoparticles for DNA Hybridization Studies. Anal. Chim. Acta 2002, 470, 51-56. 121
32. Alivisatos, A. P.; Johnsson, K. P.; Peng, X.; Wilson, T. E.; Loweth, C. J.; Bruchez, M. P.; Schultz, P. G. Organization of 'Nanocrystal Molecules' Using DNA. Nature 1996, 382, 609-611.
33. Dubertret, B.; Calame, M.; Libchaber, A. J. Single-Mismatch Detection Using Gold- Quenched Fluorescent Oligonucleotides (Vol 19, Pg 365, 2001). Nat. Biotechnol. 2001, 19, 680-681.
34. Mirkin, C. A.; Letsinger, R. L.; Mucic, R. C.; Storhoff, J. J. A DNA-Based Method for Rationally Assembling Nanoparticles into Macroscopic Materials. Nature 1996, 382, 607- 609.
35. Park, S. J.; Lazarides, A. A.; Mirkin, C. A.; Brazis, P. W.; Kannewurf, C. R.; Letsinger, R. L. The Electrical Properties of Gold Nanoparticle Assemblies Linked by DNA. Angew. Chem.-Int. Edit. 2000, 39, 3845-3848.
36. Patolsky, F.; Ranjit, K. T.; Lichtenstein, A.; Willner, I. Dendritic Amplification of DNA Analysis by Oligonucleotide-Functionalized Au-Nanoparticles. Chem. Commun. 2000, 1025-1026.
37. Bardea, A.; Dagan, A.; Ben-Dov, I.; Willner, I.; Bardea, A.; Ben-Dov, I.; Amit, B. Amplified Microgravimetric Quartz-Crystal-Microbalance Analyses of Oligonucleotide Complexes: A Route to a Tay-Sachs Biosensor Device. Chem. Comm. 1998, 839-840.
38. Letsinger, R. L.; Elghanian, R.; Viswanadham, G.; Mirkin, C. A. Use of a Steroid Cyclic Disulfide Anchor in Constructing Gold Nanoparticle−Oligonucleotide Conjugates. Bioconjugate Chem. 2000, 11, 289-291.
39. Li, Z.; Jin, R. C.; Mirkin, C. A.; Letsinger, R. L. Multiple Thiol-Anchor Capped DNA- Gold Nanoparticle Conjugates. Nucleic Acids Res. 2002, 30, 1558-1562.
40. Shaiu, W. L.; Larson, D. D.; Vesenka, J.; Henderson, E. Atomic Force Microscopy of Oriented Linear DNA-Molecules Labeled with 5nm Gold Spheres. Nucleic Acids Res. 1993, 21, 99-103.
41. Yang, X. P.; Wenzler, L. A.; Qi, J.; Li, X. J.; Seeman, N. C. Ligation of DNA Triangles Containing Double Crossover Molecules. J. Am. Chem. Soc. 1998, 120, 9779-9786.
42. Levy, M.; Cater, S. F.; Ellington, A. D. Quantum-Dot Aptamer Beacons for the Detection of Proteins. Chembiochem 2005, 6, 2163-2166.
43. Niemeyer, C. M.; Burger, W.; Peplies, J. Covalent DNA - Streptavidin Conjugates as Building Blocks for Novel Biometallic Nanostructures. Angew. Chem.-Int. Edit. 1998, 37, 2265-2268.
44. Scherer, F.; Anton, M.; Schillinger, U.; Henkel, J.; Bergemann, C.; Kruger, A.; Gansbacher, B.; Plank, C. Magnetofection: Enhancing and Targeting Gene Delivery by Magnetic Force in Vitro and in Vivo. Gene Ther. 2002, 9, 102-109.
45. Sandhu, K. K.; McIntosh, C. M.; Simard, J. M.; Smith, S. W.; Rotello, V. M. Gold Nanoparticle-Mediated Transfection of Mammalian Cells. Bioconjugate Chem. 2002, 13, 3-6. 122
46. Sullivan, M. M. O.; Green, J. J.; Przybycien, T. M. Development of a Novel Gene Delivery Scaffold Utilizing Colloidal Gold-Polyethylenimine Conjugates for DNA Condensation. Gene Ther. 2003, 10, 1882-1890.
47. He, X. X.; Wang, K. M.; Tan, W. H.; Liu, B.; Lin, X.; He, C. M.; Li, D.; Huang, S. S.; Li, J. Bioconjugated Nanoparticles for DNA Protection from Cleavage. J. Am. Chem. Soc. 2003, 125, 7168-7169.
48. Thomas, M.; Klibanov, A. M. Conjugation to Gold Nanoparticles Enhances Polyethylenimine's Transfer of Plasmid DNA into Mammalian Cells. Proc. Natl. Acad. Sci. U. S. A. 2003, 100, 9138-9143.
49. Ang, D.; Nguyen, Q. V.; Kayal, S.; Preiser, P. R.; Rawat, R. S.; Ramanujan, R. V. Insights into the Mechanism of Magnetic Particle Assisted Gene Delivery. Acta Biomater. 2011, 7, 1319-1326.
50. Paunescu, D.; Fuhrer, R.; Grass, R. N. Protection and Deprotection of DNA-High- Temperature Stability of Nucleic Acid Barcodes for Polymer Labeling. Angew. Chem.-Int. Edit. 2013, 52, 4269-4272.
51. Paunescu, D.; Puddu, M.; Soellner, J. O. B.; Stoessel, P. R.; Grass, R. N. Reversible DNA Encapsulation in Silica to Produce Ros-Resistant and Heat-Resistant Synthetic DNA 'Fossils'. Nat. Protoc. 2013, 8, 2440-2448.
52. Puddu, M.; Paunescu, D.; Stark, W. J.; Grass, R. N. Magnetically Recoverable, Thermostable, Hydrophobic DNA/Silica Encapsulates and Their Application as Invisible Oil Tags. ACS Nano 2014, 8, 2677-2685.
53. Puddu, M.; Stark, W. J.; Grass, R. N. Silica Microcapsules for Long-Term, Robust, and Reliable Room Temperature Rna Preservation. Adv. Healthc. Mater. 2015, 4, 1332-1338.
54. Revilla-Lopez, G.; Casanovas, J.; Bertran, O.; Turon, P.; Puiggali, J.; Aleman, C. Modeling Biominerals Formed by Apatites and DNA. Biointerphases 2013, 8, 1-15.
55. Okazaki, M.; Yoshida, Y.; Yamaguchi, S.; Kaneno, M.; Elliott, J. C. Affinity Binding Phenomena of DNA onto Apatite Crystals. Biomaterials 2001, 22, 2459-2464.
56. Oyane, A.; Wang, X. P.; Sogo, Y.; Ito, A.; Tsurushima, H. Calcium Phosphate Composite Layers for Surface-Mediated Gene Transfer. Acta Biomater. 2012, 8, 2034-2046.
57. Link, N.; Brunner, T. J.; Dreesen, I. A. J.; Stark, W. J.; Fussenegger, M. Inorganic Nanoparticles for Transfection of Mammalian Cells and Removal of Viruses from Aqueous Solutions. Biotechnol. Bioeng. 2007, 98, 1083-1093.
58. Smerkova, K.; Dostalova, S.; Vaculovicova, M.; Kynicky, J.; Trnkova, L.; Kralik, M.; Adam, V.; Hubalek, J.; Provaznik, I.; Kizek, R. Investigation of Interaction between Magnetic Silica Particles and Lambda Phage DNA Fragment. J. Pharm. Biomed. Anal. 2013, 86, 65-72.
59. Melzak, K. A.; Sherwood, C. S.; Turner, R. F. B.; Haynes, C. A. Driving Forces for DNA Adsorption to Silica in Perchlorate Solutions. J. Colloid Interface Sci. 1996, 181, 635- 644. 123
60. Cai, P.; Huang, Q.; Zhang, X.; Chen, H. Adsorption of DNA on Clay Minerals and Various Colloidal Particles from an Alfisol. Soil Biol. Biochem. 2006, 38, 471-476.
61. Hou, Y. K.; Wu, P. X.; Zhu, N. W. The Protective Effect of Clay Minerals against Damage to Adsorbed DNA Induced by Cadmium and Mercury. Chemosphere 2014, 95, 206- 212.
62. Choy, J. H.; Choi, S. J.; Oh, J. M.; Park, T. Clay Minerals and Layered Double Hydroxides for Novel Biological Applications. Appl. Clay Sci. 2007, 36, 122-132.
63. Choy, J. H.; Oh, J. M.; Park, M.; Sohn, K. M.; Kim, J. W. Inorganic-Biomolecular Hybrid Nanomaterials as a Genetic Molecular Code System. Adv. Mater. 2004, 16, 1181- 1184.
64. Cutler, J. I.; Auyeung, E.; Mirkin, C. A. Spherical Nucleic Acids. J. Am. Chem. Soc. 2012, 134, 1376-1391.
65. Loweth, C. J.; Caldwell, W. B.; Peng, X. G.; Alivisatos, A. P.; Schultz, P. G. DNA- Based Assembly of Gold Nanocrystals. Angew. Chem.-Int. Edit. 1999, 38, 1808-1812.
66. Mucic, R. C.; Storhoff, J. J.; Mirkin, C. A.; Letsinger, R. L. DNA-Directed Synthesis of Binary Nanoparticle Network Materials. J. Am. Chem. Soc. 1998, 120, 12674-12675.
67. Mitchell, G. P.; Mirkin, C. A.; Letsinger, R. L. Programmed Assembly of DNA Functionalized Quantum Dots. 1999, 121, 8122-8123.
68. Pinto, Y. Y.; Le, J. D.; Seeman, N. C.; Musier-Forsyth, K.; Taton, T. A.; Kiehl, R. A. Sequence-Encoded Self-Assembly of Multiple-Nanocomponent Arrays by 2d DNA Scaffolding. Nano Lett. 2005, 5, 2399-2402.
69. Zheng, J. W.; Constantinou, P. E.; Micheel, C.; Alivisatos, A. P.; Kiehl, R. A.; Seeman, N. C. Two-Dimensional Nanoparticle Arrays Show the Organizational Power of Robust DNA Motifs. Nano Lett. 2006, 6, 1502-1504.
70. Sharma, J.; Chhabra, R.; Cheng, A.; Brownell, J.; Liu, Y.; Yan, H. Control of Self- Assembly of DNA Tubules through Integration of Gold Nanoparticles. Science 2009, 323, 112-116.
71. Mastroianni, A. J.; Claridge, S. A.; Alivisatos, A. P. Pyramidal and Chiral Groupings of Gold Nanocrystals Assembled Using DNA Scaffolds. J. Am. Chem. Soc. 2009, 131, 8455- 8459.
72. Park, S. Y.; Lytton-Jean, A. K. R.; Lee, B.; Weigand, S.; Schatz, G. C.; Mirkin, C. A. DNA-Programmable Nanoparticle Crystallization. Nature 2008, 451, 553-556.
73. Nykypanchuk, D.; Maye, M. M.; van der Lelie, D.; Gang, O. DNA-Guided Crystallization of Colloidal Nanoparticles. Nature 2008, 451, 549-552.
74. Berti, L.; Burley, G. A. Nucleic Acid and Nucleotide-Mediated Synthesis of Inorganic Nanoparticles. Nat. Nanotechnol. 2008, 3, 81-87.
75. Braun, E.; Eichen, Y.; Sivan, U.; Ben-Yoseph, G. DNA-Templated Assembly and Electrode Attachment of a Conducting Silver Wire. Nature 1998, 391, 775-778. 124
76. Gu, Q.; Cheng, C. D.; Gonela, R.; Suryanarayanan, S.; Anabathula, S.; Dai, K.; Haynie, D. T. DNA Nanowire Fabrication. Nanotechnology 2006, 17, R14-R25.
77. Wang, Z. D.; Zhang, J. Q.; Ekman, J. M.; Kenis, P. J. A.; Lu, Y. DNA-Mediated Control of Metal Nanoparticle Shape: One-Pot Synthesis and Cellular Uptake of Highly Stable and Functional Gold Nanoflowers. Nano Lett. 2010, 10, 1886-1891.
78. Gugliotti, L. A.; Feldheim, D. L.; Eaton, B. E. Rna-Mediated Control of Metal Nanoparticle Shape. J. Am. Chem. Soc. 2005, 127, 17814-17818.
79. Liu, D. G.; Gugliotti, L. A.; Wu, T.; Dolska, M.; Tkachenko, A. G.; Shipton, M. K.; Eaton, B. E.; Feldheim, D. L. Rna-Mediated Synthesis of Palladium Nanoparticles on Au Surfaces. Langmuir 2006, 22, 5862-5866.
80. Coffer, J. L.; Bigham, S. R.; Pinizzotto, R. F.; Yang, H. Characterization of Quantum- Confined Cds Nanocrystallites Stabilized by Deoxyribonucleic Acid (DNA). Nanotechnology 1992, 3, 69-76.
81. Coffer, J. L.; Bigham, S. R.; Li, X.; Pinizzotto, R. F.; Rho, Y. G.; Pirtle, R. M.; Pirtle, I. L. Dictation of the Shape of Mesoscale Semiconductor Nanoparticle Assemblies by Plasmid DNA. Appl. Phys. Lett. 1996, 69, 3851-3853.
82. Ma, N.; Dooley, C. J.; Kelley, S. O. Rna-Templated Semiconductor Nanocrystals. J. Am. Chem. Soc. 2006, 128, 12598-12599.
83. Graham, F.; Van der Eb, A. A New Technique for the Assay of Infectivity of Human Adenovirus 5 DNA. Virology 1973, 52, 456-467.
84. Jordan, M.; Schallhorn, A.; Wurm, F. M. Transfecting Mammalian Cells: Optimization of Critical Parameters Affecting Calcium-Phosphate Precipitate Formation. Nucleic Acids Res. 1996, 24, 596-601.
85. Ngourn, S. C.; Butts, H. A.; Petty, A. R.; Anderson, J. E.; Gerdon, A. E. Quartz Crystal Microbalance Analysis of DNA-Templated Calcium Phosphate Mineralization. Langmuir 2012, 28, 12151-12158.
86. Liu, B.; Cao, Y.; Huang, Z.; Duan, Y.; Che, S. Silica Biomineralization Via the Self‐ Assembly of Helical Biomolecules. Adv. Mater. 2015, 27, 479-497.
87. Numata, M.; Sugiyasu, K.; Hasegawa, T.; Shinkai, S. Sol-Gel Reaction Using DNA as a Template: An Attempt toward Transcription of DNA into Inorganic Materials. Angew. Chem. Int. Ed. 2004, 43, 3279-3283.
88. Jin, C. Y.; Han, L.; Che, S. A. Synthesis of a DNA-Silica Complex with Raire Two- Dimensional Square P4mm Symmetry. Angew. Chem.-Int. Edit. 2009, 48, 9268-9272.
89. Liu, B.; Han, L.; Che, S. A. Formation of Enantiomeric Impeller-Like Helical Architectures by DNA Self-Assembly and Silica Mineralization. Angew. Chem.-Int. Edit. 2012, 51, 923-927.
90. Church, G. M.; Gao, Y.; Kosuri, S. Next-Generation Digital Information Storage in DNA. Science 2012, 337, 1628-1628. 125
91. Goldman, N.; Bertone, P.; Chen, S. Y.; Dessimoz, C.; LeProust, E. M.; Sipos, B.; Birney, E. Towards Practical, High-Capacity, Low-Maintenance Information Storage in Synthesized DNA. Nature 2013, 494, 77-80.
92. Grass, R. N.; Heckel, R.; Puddu, M.; Paunescu, D.; Stark, W. J. Robust Chemical Preservation of Digital Information on DNA in Silica Using Error Correcting Codes. Angew. Chem. Int. Ed. 2015, 54, 2552–2555.
93. Kneuer, C.; Sameti, M.; Bakowsky, U.; Schiestel, T.; Schirra, H.; Schmidt, H.; Lehr, C. M. A Nonviral DNA Delivery System Based on Surface Modified Silica-Nanoparticles Can Efficiently Transfect Cells in Vitro. Bioconjugate Chem. 2000, 11, 926-932.
94. Kneuer, C.; Sameti, M.; Haltner, E. G.; Schiestel, T.; Schirra, H.; Schmidt, H.; Lehr, C. M. Silica Nanoparticles Modified with Aminosilanes as Carriers for Plasmid DNA. Int. J. Pharm. 2000, 196, 257-261.
95. Han, G.; Martin, C. T.; Rotello, V. M. Stability of Gold Nanoparticle-Bound DNA toward Biological, Physical, and Chemical Agents. Chem. Biol. Drug Des. 2006, 67, 78-82.
96. Jin, S.; Ye, K. M. Nanoparticle-Mediated Drug Delivery and Gene Therapy. Biotechnol. Prog. 2007, 23, 32-41.
97. Seferos, D. S.; Prigodich, A. E.; Giljohann, D. A.; Patel, P. C.; Mirkin, C. A. Polyvalent DNA Nanoparticle Conjugates Stabilize Nucleic Acids. Nano Lett. 2009, 9, 308- 311.
98. Rosi, N. L.; Giljohann, D. A.; Thaxton, C. S.; Lytton-Jean, A. K. R.; Han, M. S.; Mirkin, C. A. Oligonucleotide-Modified Gold Nanoparticles for Intracellular Gene Regulation. Science 2006, 312, 1027-1030.
99. Roy, I.; Mitra, S.; Maitra, A.; Mozumdar, S. Calcium Phosphate Nanoparticles as Novel Non-Viral Vectors for Targeted Gene Delivery. Int. J. Pharm. 2003, 250, 25-33.
100. Miyata, K.; Gouda, N.; Takemoto, H.; Oba, M.; Lee, Y.; Koyama, H.; Yamasaki, Y.; Itake, K.; Nishiyama, N.; Kataoka, K. Enhanced Transfection with Silica-Coated Polyplexes Loading Plasmid DNA. Biomaterials 2010, 31, 4764-4770.
101. Sokolova, V.; Epple, M. Inorganic Nanoparticles as Carriers of Nucleic Acids into Cells. Angew. Chem.-Int. Edit. 2008, 47, 1382-1395.
102. Ito, T.; Ibe, K.; Uchino, T.; Ohshima, H.; Otsuka, M. Preparation of DNA/Gold Nanoparticle Encapsulated in Calcium Phosphate. J. Drug Deliv. 2011, 2011, 1-7.
103. Cai, P.; Huang, Q. Y.; Zhang, X. W. Interactions of DNA with Clay Minerals and Soil Colloidal Particles and Protection against Degradation by Dnase. Environ. Sci. Technol. 2006, 40, 2971-2976.
104. Franchi, M.; Gallori, E. A Surface-Mediated Origin of the Rna World: Biogenic Activities of Clay-Adsorbed Rna Molecules. Gene 2005, 346, 205-214.
105. Scappini, F.; Casadei, F.; Zamboni, R.; Franchi, M.; Gallori, E.; Monti, S. Protective Effect of Clay Minerals on Adsorbed Nucleic Acid against Uv Radiation: Possible Role in the Origin of Life. Int. J. Astrobiol. 2004, 3, 17-19. 126
106. Bernal, J. D., The Physical Basis of Life. Routledge: London, 1951.
107. Paunescu, D.; Mora, C. A.; Puddu, M.; Krumeich, F.; Grass, R. N. DNA Protection against Ultraviolet Irradiation by Encapsulation in a Multilayered Sio2/Tio2 Assembly. J. Mat. Chem. B 2014, 2, 8504-8509.
108. Ding, Y.; Jiang, Z. W.; Saha, K.; Kim, C. S.; Kim, S. T.; Landis, R. F.; Rotello, V. M. Gold Nanoparticles for Nucleic Acid Delivery. Mol. Ther. 2014, 22, 1075-1083.
109. Dobson, J. Gene Therapy Progress and Prospects: Magnetic Nanoparticle-Based Gene Delivery. Gene Ther. 2006, 13, 283-287.
110. Zhang, Y.; Satterlee, A.; Huang, L. In Vivo Gene Delivery by Nonviral Vectors: Overcoming Hurdles? Mol. Ther. 2012, 20, 1298-1304.
111. Bisht, S.; Bhakta, G.; Mitra, S.; Maitra, A. Pdna Loaded Calcium Phosphate Nanoparticles: Highly Efficient Non-Viral Vector for Gene Delivery. 2005, 288, 157-168.
112. Nel, A. E.; Madler, L.; Velegol, D.; Xia, T.; Hoek, E. M. V.; Somasundaran, P.; Klaessig, F.; Castranova, V.; Thompson, M. Understanding Biophysicochemical Interactions at the Nano-Bio Interface. 2009, 8, 543-557.
113. Radu, D. R.; Lai, C. Y.; Jeftinija, K.; Rowe, E. W.; Jeftinija, S.; Lin, V. S. Y. A Polyamidoamine Dendrimer-Capped Mesoporous Silica Nanosphere-Based Gene Transfection Reagent. J. Am. Chem. Soc. 2004, 126, 13216-13217.
114. Yezhelyev, M. V.; Qi, L. F.; O'Regan, R. M.; Nie, S.; Gao, X. H. Proton-Sponge Coated Quantum Dots for Sirna Delivery and Intracellular Imaging. J. Am. Chem. Soc. 2008, 130, 9006-9012.
115. Duan, H. W.; Nie, S. M. Cell-Penetrating Quantum Dots Based on Multivalent and Endosome-Disrupting Surface Coatings. J. Am. Chem. Soc. 2007, 129, 3333-3338.
116. Tan, W. B.; Jiang, S.; Zhang, Y. Quantum-Dot Based Nanoparticles for Targeted Silencing of Her2/Neu Gene Via Rna Interference. Biomaterials 2007, 28, 1565-1571.
117. Mykhaylyk, O.; Antequera, Y. S.; Vlaskou, D.; Plank, C. Generation of Magnetic Nonviral Gene Transfer Agents and Magnetofection in Vitro. Nat. Protoc. 2007, 2, 2391- 2411.
118. Mykhaylyk, O.; Vlaskou, D.; Tresilwised, N.; Pithayanukul, P.; Möller, W.; Plank, C. Magnetic Nanoparticle Formulations for DNA and Sirna Delivery. J. Magn. Magn. Mater. 2007, 311, 275-281.
119. Giljohann, D. A.; Seferos, D. S.; Prigodich, A. E.; Patel, P. C.; Mirkin, C. A. Gene Regulation with Polyvalent Sirna-Nanoparticle Conjugates. J. Am. Chem. Soc. 2009, 131, 2072-2073.
120. Patel, P. C.; Giljohann, D. A.; Daniel, W. L.; Zheng, D.; Prigodich, A. E.; Mirkin, C. A. Scavenger Receptors Mediate Cellular Uptake of Polyvalent Oligonucleotide- Functionalized Gold Nanoparticles. Bioconjugate Chem. 2010, 21, 2250-2256. 127
121. Wigler, M.; Silverstein, S.; Lee, L. S.; Pellicer, A.; Cheng, Y. C.; Axel, R. Transfer of Purified Herpes Virus Thymidine Kinase Gene to Cultured Mouse Cells. Cell 1977, 11, 223- 232.
122. Epple, M.; Sokolova, V. V.; Radtke, I.; Heumann, R. Effective Transfection of Cells with Multi-Shell Calcium Phosphate-DNA Nanoparticles. Biomaterials 2006, 27, 3147-3153.
123. Olton, D.; Li, J. H.; Wilson, M. E.; Rogers, T.; Close, J.; Huang, L.; Kumta, P. N.; Sfeir, C. Nanostructured Calcium Phosphates (Nanocaps) for Non-Viral Gene Delivery: Influence of the Synthesis Parameters on Transfection Efficiency. Biomaterials 2007, 28, 1267-1279.
124. Welzel, T.; Radtke, I.; Meyer-Zaika, W.; Heumann, R.; Epple, M. Transfection of Cells with Custom-Made Calcium Phosphate Nanoparticles Coated with DNA. J. Mater. Chem. 2004, 14, 2213-2217.
125. Tyner, K. M.; Roberson, M. S.; Berghorn, K. A.; Li, L.; Gilmour, R. F.; Batt, C. A.; Giannelis, E. P. Intercalation, Delivery, and Expression of the Gene Encoding Green Fluorescence Protein Utilizing Nanobiohybrids. J. Control. Release 2004, 100, 399-409.
126. Balcomb, B.; Singh, M.; Singh, S. Synthesis and Characterization of Layered Double Hydroxides and Their Potential as Nonviral Gene Delivery Vehicles. Chemistryopen 2015, 4, 137-145.
127. Storhoff, J. J.; Elghanian, R.; Mucic, R. C.; Mirkin, C. A.; Letsinger, R. L. One-Pot Colorimetric Differentiation of Polynucleotides with Single Base Imperfections Using Gold Nanoparticle Probes. J. Am. Chem. Soc. 1998, 120, 1959-1964.
128. Elghanian, R.; Storhoff, J. J.; Mucic, R. C.; Letsinger, R. L.; Mirkin, C. A. Selective Colorimetric Detection of Polynucleotides Based on the Distance-Dependent Optical Properties of Gold Nanoparticles. Science 1997, 277, 1078-1081.
129. Taton, T. A.; Mirkin, C. A.; Letsinger, R. L. Scanometric DNA Array Detection with Nanoparticle Probes. Science 2000, 289, 1757-1760.
130. Taton, T. A.; Lu, G.; Mirkin, C. A. Two-Color Labeling of Oligonucleotide Arrays Via Size-Selective Scattering of Nanoparticle Probes. J. Am. Chem. Soc. 2001, 123, 5164- 5165.
131. Lin, L.; Zhao, H. Q.; Li, J. R.; Tang, J. A.; Duan, M. X.; Jiang, L. Study on Colloidal Au-Enhanced DNA Sensing by Quartz Crystal Microbalance. Biochem. Biophys. Res. Commun. 2000, 274, 817-820.
132. Han, S. B.; Lin, J. Q.; Satjapipat, M.; Baca, A. J.; Zhou, F. M. A Three-Dimensional Heterogeneous DNA Sensing Surface Formed by Attaching Oligodeoxynucleotide-Capped Gold Nanoparticles onto a Gold-Coated Quartz Crystal. Chem. Commun. 2001, 609-610.
133. Castaneda, M. T.; Alegret, S.; Merkoci, A. Electrochemical Sensing of DNA Using Gold Nanoparticles. Electroanalysis 2007, 19, 743-753.
134. Wang, J.; Xu, D. K.; Kawde, A. N.; Polsky, R. Metal Nanoparticle-Based Electrochemical Stripping Potentiometric Detection of DNA Hybridization. Anal. Chem. 2001, 73, 5576-5581. 128
135. Han, M. Y.; Gao, X. H.; Su, J. Z.; Nie, S. Quantum-Dot-Tagged Microbeads for Multiplexed Optical Coding of Biomolecules. Nat. Biotechnol. 2001, 19, 631-635.
136. Wang, W. J.; Chen, C. L.; Qian, M. X.; Zhao, X. S. Aptamer Biosensor for Protein Detection Using Gold Nanoparticles. Anal. Biochem. 2008, 373, 213-219.
137. Lee, J. S.; Han, M. S.; Mirkin, C. A. Colorimetric Detection of Mercuric Ion (Hg2+) in Aqueous Media Using DNA-Functionalized Gold Nanoparticles. Angew. Chem.-Int. Edit. 2007, 46, 4093-4096.
138. Xue, X. J.; Wang, F.; Liu, X. G. One-Step, Room Temperature, Colorimetric Detection of Mercury (Hg2+) Using DNA/Nanoparticle Conjugates. J. Am. Chem. Soc. 2008, 130, 3244-3245.
139. Xu, X. Y.; Daniel, W. L.; Wei, W.; Mirkin, C. A. Colorimetric Cu2+ Detection Using DNA-Modified Gold-Nanoparticle Aggregates as Probes and Click Chemistry. Small 2010, 6, 623-626.
140. Dias, J. T.; Moros, M.; del Pino, P.; Rivera, S.; Grazu, V.; de la Fuente, J. M. DNA as a Molecular Local Thermal Probe for the Analysis of Magnetic Hyperthermia. Angew. Chem.- Int. Edit. 2013, 52, 11526-11529.
141. Shen, Q. P.; Nie, Z.; Guo, M. L.; Zhong, C. J.; Lin, B.; Li, W.; Yao, S. Z. Simple and Rapid Colorimetric Sensing of Enzymatic Cleavage and Oxidative Damage of Single- Stranded DNA with Unmodified Gold Nanoparticles as Indicator. Chem. Commun. 2009, 929-931.
142. Oliveira, F. C. C.; Brandão, C. R. R.; Ramalho, H. F.; da Costa, L. A. F.; Suarez, P. A. Z.; Rubim, J. C. Adulteration of Diesel/Biodiesel Blends by Vegetable Oil as Determined by Fourier Transform (Ft) near Infrared Spectrometry and Ft-Raman Spectroscopy. Anal. Chim. Acta 2007, 587, 194-199.
143. Pereira, R. C. C.; Skrobot, V. L.; Castro, E. V. R.; Fortes, I. C. P.; Pasa, V. M. D. Determination of Gasoline Adulteration by Principal Components Analysis-Linear Discriminant Analysis Applied to Ftir Spectra. Energy Fuels 2006, 20, 1097-1102.
144. Woolfe, M.; Primrose, S. Food Forensics: Using DNA Technology to Combat Misdescription and Fraud. Trends Biotechnol. 2004, 22, 222-226.
145. Tay, A.; Singh, R. K.; Krishnan, S. S.; Gore, J. P. Authentication of Olive Oil Adulterated with Vegetable Oils Using Fourier Transform Infrared Spectroscopy. Lebensm.- Wiss. Technol.-Food Sci. Technol. 2002, 35, 99-103.
146. Konig, W. A.; Fricke, C.; Saritas, Y.; Momeni, B.; Hohenfeld, G. Adulteration or Natural Variability? Enantioselective Gas Chromatography in Purity Control of Essential Oils. Hrc-J. High Res. Chrom. 1997, 20, 55-61.
147. Bryce, D. R. Magnetic Document Validator Employing Remanence and Saturation Measurements. U.S. 5,068,519, November 26, 1991.
148. McGrew, S. P. Quantum Dot Security Device and Method. U.S. 6,692,031 B2, February 17, 2004. 129
149. Small, L. D.; Highberger, G. Thermochromic Ink Formulations, Nail Lacquer and Methods of Use. U.S. 5,997,849, December 7, 1997.
150. Welle, R. P. Isotopic Taggant Method and Composition. U.S. 5,760,394, June 2, 1998.
151. Shchegolikhin, A. N.; Lazareva, O. L.; Mel'nikov, V. P.; Ozeretski, V. Y.; Small, L. D. Raman-Active Taggants and Their Recognition. U.S. 6,610,351 B2, August 26, 2003.
152. Kydd, P. H. Polypeptides as Chemical Tagging Materials U.S. 4,441,943, April 10, 1984.
153. Lebacq, P. Method and Apparatus for High Security Crypto-Marking for Protecting Valuable Objects. U.S. 5,139,812, August 18, 1992.
154. Slater, J. H.; Minton, J. E. Method of Marking a Liquid. U.S. 5,643,728, July 1, 1997.
155. Paunescu, D.; Fuhrer, R.; Grass, R. N. Protection and Deprotection of DNA - High Temperature Stability of Nucleic Acid Barcodes for Polymer Labeling. Angew. Chem. Int. Ed. 2013, 52, 4269-4272.
156. Zhang, Z.; Balogh, D.; Wang, F.; Sung, S. Y.; Nechushtai, R.; Willner, I. Biocatalytic Release of an Anticancer Drug from Nucleic-Acids-Capped Mesoporous Sio2 Using DNA or Molecular Biomarkers as Triggering Stimuli. ACS Nano 2013, 7, 8455-8468.
157. Niemeyer, C. M. Self-Assembled Nanostructures Based on DNA: Towards the Development of Nanobiotechnology. Curr. Opin. Chem. Biol. 2000, 4, 609-618.
158. Auyeung, E.; Li, T. I. N. G.; Senesi, A. J.; Schmucker, A. L.; Pals, B. C.; de la Cruz, M. O.; Mirkin, C. A. DNA-Mediated Nanoparticle Crystallization into Wulff Polyhedra. Nature 2013, 505, 73-77.
159. Davila-Ibanez, A. B.; Salgueirino, V.; Martinez-Zorzano, V.; Mariño-Fernández, R.; García-Lorenzo, A.; Maceira-Campos, M.; Muñoz-Ubeda, M.; Junquera, E.; Aicart, E.; Rivas, J.; et al. Magnetic Silica Nanoparticle Cellular Uptake and Cytotoxicity Regulated by Electrostatic Polyelectrolytes–DNA Loading at Their Surface. ACS Nano 2011, 6, 747-759.
160. Ruiz-Hernández, E.; Baeza, A.; Vallet-Regí, M. Smart Drug Delivery through DNA/Magnetic Nanoparticle Gates. ACS Nano 2011, 5, 1259-1266.
161. Park, D. H.; Kim, J. E.; Oh, J. M.; Shul, Y. G.; Choy, J. H. DNA Core@Inorganic Shell. J. Am. Chem. Soc. 2010, 132, 16735-16736.
162. Lindahl, T. Instability and Decay of the Primary Structure of DNA. Nature 1993, 362, 709-715.
163. Zelikin, A. N.; Becker, A. L.; Johnston, A. P. R.; Wark, K. L.; Turatti, F.; Caruso, F. A General Approach for DNA Encapsulation in Degradable Polymer Microcapsules. ACS Nano 2007, 1, 63-69.
164. Smith, S.; Morin, P. A. Optimal Storage Conditions for Highly Dilute DNA Samples: A Role for Trehalose as a Preserving Agent. J. Forensic Sci. 2005, 50, 1101-1108. 130
165. Rajendram, D.; Ayenza, R.; Holder, F. M.; Moran, B.; Long, T.; Shah, H. N. Long- Term Storage and Safe Retrieval of DNA from Microorganisms for Molecular Analysis Using Fta Matrix Cards. J. Microbiol. Methods 2006, 67, 582-592.
166. Yang, H.; Zheng, K.; Zhang, Z.; Shi, W.; Jing, S.; Wang, L.; Zheng, W.; Zhao, D.; Xu, J.; Zhang, P. Adsorption and Protection of Plasmid DNA on Mesoporous Silica Nanoparticles Modified with Various Amounts of Organosilane. J. Colloid Interface Sci. 2012, 369, 317- 322.
167. Kapusuz, D.; Durucan, C. Synthesis of DNA-Encapsulated Silica Elaborated by Sol- Gel Routes. J. Mater. Res. 2013, 28, 175-184.
168. Lu, A. H.; Salabas, E. L.; Schuth, F. Magnetic Nanoparticles: Synthesis, Protection, Functionalization, and Application. Angew. Chem. Int. Edit. 2007, 46, 1222-1244.
169. Aguilar-Arteaga, K.; Rodriguez, J. A.; Barrado, E. Magnetic Solids in Analytical Chemistry: A Review. Anal. Chim. Acta 2010, 674, 157-165.
170. Hao, R.; Xing, R. J.; Xu, Z. C.; Hou, Y. L.; Gao, S.; Sun, S. H. Synthesis, Functionalization, and Biomedical Applications of Multifunctional Magnetic Nanoparticles. Adv. Mater. 2010, 22, 2729-2742.
171. Sharma, A. N.; Luo, D.; Walter, M. T. Hydrological Tracers Using Nanobiotechology: Proof of Concept. Environ. Sci. Technol. 2012, 46, 8928-8936.
172. Liu, B.; Yao, Y.; Che, S. Template-Assisted Self-Assembly: Alignment, Placement, and Arrangement of Two-Dimensional Mesostructured DNA–Silica Platelets. Angew. Chem. Int. Ed. 2013, 52, 1-6.
173. Wu, W.; He, Q.; Jiang, C. Magnetic Iron Oxide Nanoparticles: Synthesis and Surface Functionalization Strategies. Nanoscale Res. Lett. 2008, 3, 397-415.
174. Mitchell, K. K. P.; Liberman, A.; Kummel, A. C.; Trogler, W. C. Iron(Iii)-Doped, Silica Nanoshells: A Biodegradable Form of Silica. J. Am. Chem. Soc. 2012, 134, 13997- 14003.
175. Aguilar, F.; Charrondiere, U. R.; Dusemund, B.; Galtier, P.; Gilbert, J.; Gott, D. M.; Grilli, S.; Guertler, R.; Kass, G. E. N.; Koenig, J.; et al. Calcium Silicate and Silicon Dioxide/Silicic Acid Gel Added for Nutritional Purposes to Food Supplements. EFSA J. 2009, 1132, 1-24.
176. Castanotto, D.; Rossi, J. J. The Promises and Pitfalls of Rna-Interference-Based Therapeutics. Nature 2009, 457, 426-433.
177. Nolan, T.; Hands, R. E.; Bustin, S. A. Quantification of Mrna Using Real-Time Rt- Pcr. Nat. Protoc. 2006, 1, 1559-1582.
178. Bustin, S. A.; Mueller, R. Real-Time Reverse Transcription Pcr (Qrt-Pcr) and Its Potential Use in Clinical Diagnosis. Clin. Sci. 2005, 109, 365-379.
179. Marioni, J. C.; Mason, C. E.; Mane, S. M.; Stephens, M.; Gilad, Y. Rna-Seq: An Assessment of Technical Reproducibility and Comparison with Gene Expression Arrays. Genome Res. 2008, 18, 1509-1517. 131
180. Butte, A. The Use and Analysis of Microarray Data. Nat. Rev. Drug Discov. 2002, 1, 951-960.
181. Wang, Z.; Gerstein, M.; Snyder, M. Rna-Seq: A Revolutionary Tool for Transcriptomics. Nat. Rev. Genet. 2009, 10, 57-63.
182. Mortazavi, A.; Williams, B. A.; McCue, K.; Schaeffer, L.; Wold, B. Mapping and Quantifying Mammalian Transcriptomes by Rna-Seq. Nat. Methods 2008, 5, 621-628.
183. Kruger, K.; Grabowski, P. J.; Zaug, A. J.; Sands, J.; Gottschling, D. E.; Cech, T. R. Self-Splicing Rna: Autoexcision and Autocyclization of the Ribosomal Rna Intervening Sequence of Tetrahymena. Cell 1982, 31, 147-157.
184. Guerrier-Takada, C.; Gardiner, K.; Marsh, T.; Pace, N.; Altman, S. The Rna Moiety of Ribonuclease P Is the Catalytic Subunit of the Enzyme. Cell 1983, 35, 849-857.
185. Haseloff, J.; Gerlach, W. L. Simple Rna Enzymes with New and Highly Specific Endoribonuclease Activities. Nature 1988, 334, 585-591.
186. Uhlenbeck, O. C. A Small Catalytic Oligoribonucleotide. Nature 1987, 328, 596-600.
187. Sullenger, B. A.; Gilboa, E. Emerging Clinical Applications of Rna. Nature 2002, 418, 252-258.
188. Tuerk, C.; Gold, L. Systematic Evolution of Ligands by Exponential Enrichment - Rna Ligands to Bacteriophage-T4 DNA-Polymerase. Science 1990, 249, 505-510.
189. Ellington, A. D.; Szostak, J. W. Invitro Selection of Rna Molecules That Bind Specific Ligands. Nature 1990, 346, 818-822.
190. Sundaram, P.; Kurniawan, H.; Byrne, M. E.; Wower, J. Therapeutic Rna Aptamers in Clinical Trials. Eur. J. Pharm. Sci. 2013, 48, 259-271.
191. Keefe, A. D.; Pai, S.; Ellington, A. Aptamers as Therapeutics. Nat. Rev. Drug Discov. 2010, 9, 537-550.
192. Coleman, J.; Green, P. J.; Inouye, M. The Use of Rnas Complementary to Specific Mrnas to Regulate the Expression of Individual Bacterial Genes. Cell 1984, 37, 429-436.
193. Izant, J. G.; Weintraub, H. Constitutive and Conditional Suppression of Exogenous and Endogenous Genes by Anti-Sense Rna. Science 1985, 229, 345-352.
194. Cong, L.; Ran, F. A.; Cox, D.; Lin, S.; Barretto, R.; Habib, N.; Hsu, P. D.; Wu, X.; Jiang, W.; Marraffini, L. A.; Zhang, F. Multiplex Genome Engineering Using Crispr/Cas Systems. Science 2013, 339, 819-823.
195. Platt, R. J.; Chen, S.; Zhou, Y.; Yim, M. J.; Swiech, L.; Kempton, H. R.; Dahlman, J. E.; Parnas, O.; Eisenhaure, T. M.; Jovanovic, M.; Graham, D. B.; Jhunjhunwala, S.; Heidenreich, M.; Xavier, R. J.; Langer, R.; Anderson, D. G.; Hacohen, N.; Regev, A.; Feng, G.; Sharp, P. A.; Zhang, F. Crispr-Cas9 Knockin Mice for Genome Editing and Cancer Modeling. Cell 2014, 159, 440-455. 132
196. Li, Z. W.; Wu, J. H.; DeLeo, C. J. Rna Damage and Surveillance under Oxidative Stress. IUBMB Life 2006, 58, 581-588.
197. Li, Y. F.; Breaker, R. R. Kinetics of Rna Degradation by Specific Base Catalysis of Transesterification Involving the 2 '-Hydroxyl Group. J. Am. Chem. Soc. 1999, 121, 5364- 5372.
198. Fabre, A. L.; Colotte, M.; Luis, A.; Tuffet, S.; Bonnet, J. An Efficient Method for Long-Term Room Temperature Storage of Rna. Eur. J. Hum. Genet. 2014, 22, 379-385.
199. Thompson, J. E.; Kutateladze, T. G.; Schuster, M. C.; Venegas, F. D.; Messmore, J. M.; Raines, R. T. Limits to Catalysis by Ribonuclease A. Bioorg. Chem. 1995, 23, 471-481.
200. Chomczynski, P. Solubilization in Formamide Protects Rna from Degradation. Nucleic Acids Res. 1992, 20, 3791-3791.
201. Seyhan, A. A.; Burke, J. M. Mg2+-Independent Hairpin Ribozyme Catalysis in Hydrated Rna Films. RNA 2000, 6, 189-198.
202. Guo, P.; Haque, F.; Hallahan, B.; Reif, R.; Li, H. Uniqueness, Advantages, Challenges, Solutions, and Perspectives in Therapeutics Applying Rna Nanotechnology. Nucleic Acid Ther. 2012, 22, 226-245.
203. Fleige, S.; Pfaffl, M. W. Rna Integrity and the Effect on the Real-Time Qrt-Pcr Performance. Mol. Aspects Med. 2006, 27, 126-139.
204. Vermeulen, J.; De Preter, K.; Lefever, S.; Nuytens, J.; De Vloed, F.; Derveaux, S.; Hellemans, J.; Speleman, F.; Vandesompele, J. Measurable Impact of Rna Quality on Gene Expression Results from Quantitative Pcr. Nucleic Acids Res. 2011, 39, 1-12.
205. Hernandez, G. E.; Mondala, T. S.; Head, S. R. Assessing a Novel Room-Temperature Rna Storage Medium for Compatibility in Microarray Gene Expression Analysis. Biotechniques 2009, 47, 667-670.
206. Seelenfreund, E.; Robinson, W. A.; Amato, C. M.; Tan, A. C.; Kim, J.; Robinson, S. E. Long Term Storage of Dry Versus Frozen Rna for Next Generation Molecular Studies. PLoS One 2014, 9, 1-6.
207. Natarajan, P.; Trinh, T.; Mertz, L.; Goldsborough, M.; Fox, D. Paper-Based Archiving of Mammalian and Plant Samples for Rna Analysis. Biotechniques 2000, 29, 1328-1333.
208. Kievit, F. M.; Veiseh, O.; Bhattarai, N.; Fang, C.; Gunn, J. W.; Lee, D.; Ellenbogen, R. G.; Olson, J. M.; Zhang, M. Q. Pei-Peg-Chitosan-Copolymer-Coated Iron Oxide Nanoparticles for Safe Gene Delivery: Synthesis, Complexation, and Transfection. Adv. Funct. Mater. 2009, 19, 2244-2251.
209. Mhanna, R.; Kashyap, A.; Palazzolo, G.; Vallmajo-Martin, Q.; Becher, J.; Moller, S.; Schnabelrauch, M.; Zenobi-Wong, M. Chondrocyte Culture in Three Dimensional Alginate Sulfate Hydrogels Promotes Proliferation While Maintaining Expression of Chondrogenic Markers. Tissue Eng. Part A 2014, 20, 1454-1464. 133
210. Mhanna, R.; Ozturk, E.; Schlink, P.; Zenobi-Wong, M. Probing the Microenvironmental Conditions for Induction of Superficial Zone Protein Expression. Osteoarthritis Cartilage 2013, 21, 1924-1932.
211. Kua, J.; Bada, J. L. Primordial Ocean Chemistry and Its Compatibility with the Rna World. Orig. Life Evol. Biosph. 2011, 41, 553-558.
212. Linjalahti, H.; Mikkola, S. Intra- and Intermolecular Interactions Influence the Reactivity of Rna Oligonucleotides. Chem. Biodivers. 2007, 4, 2938-2947.
213. Tripathy, S. K.; Black, H. B.; Goldwasser, E.; Leiden, J. M. Immune Responses to Transgene-Encoded Proteins Limit the Stability of Gene Expression after Injection of Replication-Defective Adenovirus Vectors. Nat. Med. 1996, 2, 545-550.
214. Godbey, W. T.; Wu, K. K.; Mikos, A. G. Poly(Ethylenimine) and Its Role in Gene Delivery. J. Control. Release 1999, 60, 149-160.
215. Hsu, C. Y. M.; Uludag, H. A Simple and Rapid Nonviral Approach to Efficiently Transfect Primary Tissue-Derived Cells Using Polyethylenimine. Nat. Protoc. 2012, 7, 935- 945.
216. Hu, Y.; Zhao, N. N.; Yu, B. R.; Liu, F. S.; Xu, F. J. Versatile Types of Polysaccharide- Based Supramolecular Polycation/Pdna Nanoplexes for Gene Delivery. Nanoscale 2014, 6, 7560-7569.
217. Skandrani, N.; Barras, A.; Legrand, D.; Gharbi, T.; Boulahdour, H.; Boukherroub, R. Lipid Nanocapsules Functionalized with Polyethyleneimine for Plasmid DNA and Drug Co- Delivery and Cell Imaging. Nanoscale 2014, 6, 7379-7390.
218. Fischer, D.; Li, Y. X.; Ahlemeyer, B.; Krieglstein, J.; Kissel, T. In Vitro Cytotoxicity Testing of Polycations: Influence of Polymer Structure on Cell Viability and Hemolysis. Biomaterials 2003, 24, 1121-1131.
219. Dorozhkin, S. V.; Epple, M. Biological and Medical Significance of Calcium Phosphates. Angew. Chem.-Int. Edit. 2002, 41, 3130-3146.
220. Tadic, D.; Epple, M. A Thorough Physicochemical Characterisation of 14 Calcium Phosphate-Based Bone Substitution Materials in Comparison to Natural Bone. Biomaterials 2004, 25, 987-994.
221. Theiss, F.; Apelt, D.; Brand, B. A.; Kutter, A.; Zlinszky, K.; Bohner, M.; Matter, S.; Frei, C.; Auer, J. A.; von Rechenberg, B. Biocompatibility and Resorption of a Brushite Calcium Phosphate Cement. Biomaterials 2005, 26, 4383-4394.
222. Rezwan, K.; Chen, Q. Z.; Blaker, J. J.; Boccaccini, A. R. Biodegradable and Bioactive Porous Polymer/Inorganic Composite Scaffolds for Bone Tissue Engineering. Biomaterials 2006, 27, 3413-3431.
223. Hu, J.; Kovtun, A.; Tomaszewski, A.; Singer, B. B.; Seitz, B.; Epple, M.; Steuhl, K.- P.; Ergün, S.; Fuchsluger, T. A. A New Tool for the Transfection of Corneal Endothelial Cells: Calcium Phosphate Nanoparticles. Acta Biomater. 2012, 8, 1156-1163. 134
224. LeGeros, R. Z. Calcium Phosphate-Based Osteoinductive Materials. Chem. Rev. 2008, 108, 4742-4753.
225. Isalan, M.; Santori, M. I.; Gonzalez, C.; Serrano, L. Localized Transfection on Arrays of Magnetic Beads Coated with Pcr Products. Nat. Med. 2005, 2, 113-118.
226. Houchin-Ray, T.; Whittlesey, K. J.; Shea, L. D. Spatially Patterned Gene Delivery for Localized Neuron Survival and Neurite Extension. Mol. Ther. 2007, 15, 705-712.
227. Hasty, J.; McMillen, D.; Collins, J. J. Engineered Gene Circuits. Nature 2002, 420, 224-230.
228. Ziauddin, J.; Sabatini, D. M. Microarrays of Cells Expressing Defined Cdnas. Nature 2001, 411, 107-110.
229. Chen, C. C.; Lin, Y. P.; Wang, C. W.; Tzeng, H. C.; Wu, C. H.; Chen, Y. C.; Chen, C. P.; Chen, L. C.; Wu, Y. C. DNA-Gold Nanorod Conjugates for Remote Control of Localized Gene Expression by near Infrared Irradiation. J. Am. Chem. Soc. 2006, 128, 3709-3715.
230. Hu, W. W.; Wang, Z.; Hollister, S. J.; Krebsbach, P. H. Localized Viral Vector Delivery to Enhance in Situ Regenerative Gene Therapy. Gene Ther. 2007, 14, 891-901.
231. Santori, M. I.; Gonzalez, C.; Serrano, L.; Isalan, M. Localized Transfection with Magnetic Beads Coated with Pcr Products and Other Nucleic Acids. Nat. Protoc. 2006, 1, 526-531.
232. Madler, L.; Kammler, H. K.; Mueller, R.; Pratsinis, S. E. Controlled Synthesis of Nanostructured Particles by Flame Spray Pyrolysis. J. Aerosol Sci. 2002, 33, 369-389.
233. Loher, S.; Stark, W. J.; Maciejewski, M.; Baiker, A.; Pratsinis, S. E.; Reichardt, D.; Maspero, F.; Krumeich, F.; Gunther, D. Fluoro-Apatite and Calcium Phosphate Nanoparticles by Flame Synthesis. Chem. Mater. 2005, 17, 36-42.
234. Schumacher, C. M.; Herrmann, I. K.; Bubenhofer, S. B.; Gschwind, S.; Hirt, A.-M.; Beck-Schimmer, B.; Guenther, D.; Stark, W. J. Quantitative Recovery of Magnetic Nanoparticles from Flowing Blood: Trace Analysis and the Role of Magnetization. Adv. Funct. Mater. 2013, 23, 4888-4896.
235. Weissleder, R.; Stark, D. D.; Engelstad, B. L.; Bacon, B. R.; Compton, C. C.; White, D. L.; Jacobs, P.; Lewis, J. Superparamagnetic Iron Oxide: Pharmacokinetics and Toxicity. Am. J. Roentgenol. 1989, 152, 167-173.
236. Gupta, A. K.; Gupta, M. Synthesis and Surface Engineering of Iron Oxide Nanoparticles for Biomedical Applications. Biomaterials 2005, 26, 3995-4021.
237. Yu, M. K.; Jeong, Y. Y.; Park, J.; Park, S.; Kim, J. W.; Min, J. J.; Kim, K.; Jon, S. Drug-Loaded Superparamagnetic Iron Oxide Nanoparticles for Combined Cancer Imaging and Therapy in Vivo. Angew. Chem.-Int. Edit. 2008, 47, 5362-5365.
238. Jain, T. K.; Reddy, M. K.; Morales, M. A.; Leslie-Pelecky, D. L.; Labhasetwar, V. Biodistribution, Clearance, and Biocompatibility of Iron Oxide Magnetic Nanoparticles in Rats. Mol. Pharm. 2008, 5, 316-327. 135
239. Wu, Y.; Jiang, W.; Wen, X.; He, B.; Zeng, X.; Wang, G.; Gu, Z. A Novel Calcium Phosphate Ceramic-Magnetic Nanoparticle Composite as a Potential Bone Substitute. Biomed. Mater. 2010, 5, 1-7.
240. Tang, Z.; Zhou, Y.; Sun, H.; Li, D.; Zhou, S. Biodegradable Magnetic Calcium Phosphate Nanoformulation for Cancer Therapy. Eur. J. Pharm. Biopharm. 2014, 87, 90-100.
241. Grass, R. N.; Albrecht, T. F.; Krumeich, F.; Stark, W. J. Large-Scale Preparation of Ceria/Bismuth Metal-Matrix Nano-Composites with a Hardness Comparable to Steel. J. Mater. Chem. 2007, 17, 1485-1490.
242. Teleki, A.; Suter, M.; Kidambi, P. R.; Ergeneman, O.; Krumeich, F.; Nelson, B. J.; Pratsinis, S. E. Hermetically Coated Superparamagnetic Fe2o3 Particles with Sio2 Nanofilms. Chem. Mat. 2009, 21, 2094-2100.
243. Stark, W. J.; Baiker, A.; Pratsinis, S. E. Nanoparticle Opportunities: Pilot-Scale Flame Synthesis of Vanadia/Titania Catalysts. Part. Part. Syst. Char. 2002, 19, 306-311.
244. Osterwalder, N.; Capello, C.; Hungerbuhler, K.; Stark, W. J. Energy Consumption During Nanoparticle Production: How Economic Is Dry Synthesis? J. Nanopart. Res. 2006, 8, 1-9.
245. Huang, S.-J.; Ke, J.-H.; Chen, G.-J.; Wang, L.-F. One-Pot Synthesis of Pdmaema- Bound Iron Oxide Nanoparticles for Magnetofection. J. Mater. Chem. B 2013, 1, 5916-5924.
246. Lungwitz, U.; Breunig, M.; Blunk, T.; Gopferich, A. Polyethylenimine-Based Non- Viral Gene Delivery Systems. Eur. J. Pharm. Biopharm. 2005, 60, 247-266.
247. Kucsko, G.; Maurer, P. C.; Yao, N. Y.; Kubo, M.; Noh, H. J.; Lo, P. K.; Park, H.; Lukin, M. D. Nanometre-Scale Thermometry in a Living Cell. Nature 2013, 500, 54-58.
248. Brites, C. D. S.; Lima, P. P.; Silva, N. J. O.; Millan, A.; Amaral, V. S.; Palacio, F.; Carlos, L. D. Thermometry at the Nanoscale. Nanoscale 2012, 4, 4799-4829.
249. Lee, J.; Kotov, N. A. Thermometer Design at the Nanoscale. Nano Today 2007, 2, 48- 51.
250. Alaskar, M.; Ames, M.; Liu, C.; Li, K. W.; Horne, R. Temperature Nanotracers for Fractured Reservoirs Characterization. J. Pet. Sci. Eng. 2015, 127, 212-228.
251. Brites, C. D. S.; Lima, P. P.; Silva, N. J. O.; Millan, A.; Amaral, V. S.; Palacio, F.; Carlos, L. D. A Luminescent Molecular Thermometer for Long-Term Absolute Temperature Measurements at the Nanoscale. Adv. Mater. 2010, 22, 4499-4504.
252. Ross, D.; Gaitan, M.; Locascio, L. E. Temperature Measurement in Microfluidic Systems Using a Temperature-Dependent Fluorescent Dye. Anal. Chem. 2001, 73, 4117- 4123.
253. Tsuji, T.; Yoshida, S.; Yoshida, A.; Uchiyama, S. Cationic Fluorescent Polymeric Thermometers with the Ability to Enter Yeast and Mammalian Cells for Practical Intracellular Temperature Measurements. Anal. Chem. 2013, 85, 9815-9823. 136
254. Okabe, K.; Inada, N.; Gota, C.; Harada, Y.; Funatsu, T.; Uchiyama, S. Intracellular Temperature Mapping with a Fluorescent Polymeric Thermometer and Fluorescence Lifetime Imaging Microscopy. Nat. Commun. 2012, 3, 1-9.
255. Chen, C. Y.; Chen, C. T. A Pnipam-Based Fluorescent Nanothermometer with Ratiometric Readout. Chem. Commun. 2011, 47, 994-996.
256. Donner, J. S.; Thompson, S. A.; Kreuzer, M. P.; Baffou, G.; Quidant, R. Mapping Intracellular Temperature Using Green Fluorescent Protein. Nano Lett. 2012, 12, 2107-2111.
257. Yang, J. M.; Yang, H.; Lin, L. Quantum Dot Nano Thermometers Reveal Heterogeneous Local Thermogenesis in Living Cells. ACS Nano 2011, 5, 5067-5071.
258. Li, S.; Zhang, K.; Yang, J. M.; Lin, L. W.; Yang, H. Single Quantum Dots as Local Temperature Markers. Nano Lett. 2007, 7, 3102-3105.
259. Maestro, L. M.; Jacinto, C.; Silva, U. R.; Vetrone, F.; Capobianco, J. A.; Jaque, D.; Sole, J. G. Cdte Quantum Dots as Nanothermometers: Towards Highly Sensitive Thermal Imaging. Small 2011, 7, 1774-1778.
260. Vetrone, F.; Naccache, R.; Zamarron, A.; de la Fuente, A. J.; Sanz-Rodriguez, F.; Maestro, L. M.; Rodriguez, E. M.; Jaque, D.; Sole, J. G.; Capobianco, J. A. Temperature Sensing Using Fluorescent Nanothermometers. ACS Nano 2010, 4, 3254-3258.
261. Riedinger, A.; Guardia, P.; Curcio, A.; Garcia, M. A.; Cingolani, R.; Manna, L.; Pellegrino, T. Subnanometer Local Temperature Probing and Remotely Controlled Drug Release Based on Azo-Functionalized Iron Oxide Nanoparticles. Nano Lett. 2013, 13, 2399- 2406.
262. Gustafson, T. P.; Cao, Q.; Wang, S. T.; Berezin, M. Y. Design of Irreversible Optical Nanothermometers for Thermal Ablations. Chem. Commun. 2013, 49, 680-682.
263. Oyama, K.; Takabayashi, M.; Takei, Y.; Arai, S.; Takeoka, S.; Ishiwata, S.; Suzuki, M. Walking Nanothermometers: Spatiotemporal Temperature Measurement of Transported Acidic Organelles in Single Living Cells. Lab Chip 2012, 12, 1591-1593.
264. Brites, C. D. S.; Lima, P. P.; Silva, N. J. O.; Millan, A.; Amaral, V. S.; Palacio, F.; Carlos, L. D. Ratiometric Highly Sensitive Luminescent Nanothermometers Working in the Room Temperature Range. Applications to Heat Propagation in Nanofluids. Nanoscale 2013, 5, 7572-7580.
265. Choi, J. H.; Burns, A. A.; Williams, R. M.; Zhou, Z. X.; Flesken-Nikitin, A.; Zipfel, W. R.; Wiesner, U.; Nikitin, A. Y. Core-Shell Silica Nanoparticles as Fluorescent Labels for Nanomedicine. J. Biomed. Opt. 2007, 12, 1-11.
266. Jonstrup, A. T.; Fredsoe, J.; Andersen, A. H. DNA Hairpins as Temperature Switches, Thermometers and Ionic Detectors. Sensors 2013, 13, 5937-5944.
267. Ebrahimi, S.; Akhlaghi, Y.; Kompany-Zareh, M.; Rinnan, A. Nucleic Acid Based Fluorescent Nanothermometers. ACS Nano 2014, 8, 10372-10382.
268. Tashiro, R.; Sugiyama, H. A Nanothermometer Based on the Different Pi Stackings of B- and Z-DNA. Angew. Chem.-Int. Edit. 2003, 42, 6018-6020. 137
269. Pope, E. J. A.; Mackenzie, J. D. Sol-Gel Processing of Silica .2. The Role of the Catalyst. J. Non-Cryst. Solids 1986, 87, 185-198.
270. Allentoft, M. E.; Collins, M.; Harker, D.; Haile, J.; Oskam, C. L.; Hale, M. L.; Campos, P. F.; Samaniego, J. A.; Gilbert, M. T. P.; Willerslev, E.; Zhang, G. J.; Scofield, R. P.; Holdaway, R. N.; Bunce, M. The Half-Life of DNA in Bone: Measuring Decay Kinetics in 158 Dated Fossils. Proc. R. Soc. B-Biol. Sci. 2012, 279, 4724-4733.
271. Smoluchowski, M. Zur Kinetischen Theorie Der Brownschen Molekularbewegung Und Der Suspensionen. Ann. Phys. 1906, 21, 756-780.
272. Brunner, T. J.; Wick, P.; Manser, P.; Spohn, P.; Grass, R. N.; Limbach, L. K.; Bruinink, A.; Stark, W. J. In Vitro Cytotoxicity of Oxide Nanoparticles: Comparison to Asbestos, Silica, and the Effect of Particle Solubility. Environ. Sci. Technol. 2006, 40, 4374- 4381.