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WIREs Nanomed Nanobiotechnol
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Self‐assembly and applications of nucleic acid solid‐state films

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Abstract While most nucleic acid (NA)–lipid or NA–polymer complexes are studied in solution, there is growing interest in understanding their properties as naturally derived, biodegradable, biocompatible, solid‐state materials with tailorable properties influenced by environmental parameters. Therapeutic and cell programming applications comprise an important new research field, particularly in gene transfection and silencing using plasmid DNA and siRNA with targeted local delivery for use in cell culture. Dried solid films have lower nuclease degradation, fewer barriers to long term storage, and allow localized delivery by direct implantation in combination with controlled release and dosage adjustment. In contrast to particulate complexes or other methods of drug delivery which are prepared and must remain in solution, films can regain their biological activity once wetted. However our understanding of the types of cationic agents that predictably form self‐standing films with NA is still limited. The self‐assembly and structural, physical, and chemical properties of these materials are of key importance to maintaining their activity. We therefore discuss the material properties of NA–lipid and NA–polymer films as the focus of this article. Recent studies have indicated that there is also growing interest in NA films beyond bioengineering and medical applications in the fields of nano‐ and optoelectronics. We survey the self‐assembly of solid‐state materials composed of NA complexed with lipids, surfactants, or polymers, and summarize investigations of nanoscopic assembly, structure, optical, and macroscopic material properties. We further evaluate the current and future applications of NA–lipid and NA–polymer films and the benefits and drawbacks of each type. WIREs Nanomed Nanobiotechnol 2011 3 479–500 DOI: 10.1002/wnan.148 This article is categorized under: Therapeutic Approaches and Drug Discovery > Emerging Technologies Implantable Materials and Surgical Technologies > Nanomaterials and Implants Nanotechnology Approaches to Biology > Nanoscale Systems in Biology

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Depiction of common structures of DNA, lipids, and DNA–lipid complexes in solution. (a) DNA, formed from a sequence of the four nucleotides adenine, cytosine, thymine, and guanosine, (b) can adopt several double‐helical structural states known as A‐, B‐, C‐, and Z‐form depending on the surrounding conditions. (Reprinted with permission from Ref 14. Copyright 2003 Nucleic Acids Research). (c) DNA oligonucleotides with programmed sequences can self‐assemble into tiles and other complex 2D and 3D shapes. The displayed DNA oligonucleotide nanostructures form a lamellar sheet construct by self‐assembly from eight sequences shown in the AFM image at right. (Reprinted with permission from Ref 17. Copyright 2005 American Chemical Society). (d) Lipids and surfactants adopt various structural assemblies depending on the shape of the molecule. Rod‐like lipids tend to self‐assemble into lamellar sheets while (inverted) cone‐like lipids prefer a hexagonal assembly. (Reprinted with permission from Ref 26. Copyright 1997 National Academy of Sciences, USA; Reprinted with permission from Ref 27. Copyright 2009 PMC Biophysics). (e) Structure of complexes of the same lipid types shown in (d) complexed with DNA in solution. The overall structure type is essentially unchanged, leading to the conclusion that the lipid structure has primary influence over the complex structure. (Reprinted with permission from Ref 27. Copyright 2009 PMC Biophysics)

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(a) Cartoon of the steps leading to gene expression in cells exposed to a solid film containing plasmid DNA: (1) release of vector from film, (2) interaction of vector with cell membrane, (3) vector is endocytosed, (4) formation of early endosome, (5) late endosome transport, (6) endosomal escape, (7) transport to nucleus and dissociation of delivery agent, (8) nuclear entry, (9) tRNA transcription, (10) transport of RNA into the cytoplasm, and (11) translation of RNA into protein. (Reprinted with permission from Ref 42. Copyright 2005 MRS Bulletin). (b) Cross‐sectional view of 3 : 1 PLG sponge fabricated by a gas foaming procedure. (c) and (d) In vitro transfection by plasmid DNA released at different times from the polymer sponge. (c) The percentage of cells transfected by calcium precipitating unincorporated plasmid DNA (+ control) and plasmid DNA that was collected at different times of release. (d) Quantification of enzymatic activity of the produced protein by a mock transfection (–control), transfection using unincorporated plasmid DNA (+ control), and plasmid DNA collected at different times of release. The relative light units were recorded from a luminometer and normalized per milligram of total protein. Values represent mean and standard deviation. (Reprinted with permission from Ref 36. Copyright 1999 Nature Publishing Group). (e) Demonstration of localized transfection as measured by enhanced green fluorescent protein (eGFP) expression in COS‐7 cells from a multilayered DNA–polymer film cast on a quartz slide. The top scheme shows the idealized layer‐by‐layer fabrication of GFP‐encoding plasmid DNA (green) and polymer (gray) film which degrades slowly under physiological conditions to release DNA. Polymer 1 is a custom‐synthesized poly(β‐aminoester). The bottom scheme shows the direct localized transfection of cells placed underneath such a film coated on a quartz slide. (Reprinted with permission from Ref 37. Copyright 2005 Elsevier BV).

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(a) Transmissivity of a 3 µm thick film made of DNA and the cationic surfactant CTMA indicating that the film is highly transparent over a wide range of wavelength. (b) Variation of the film refractive index with wavelength. The DNA–CTMA film is considered a low‐loss optical material as its refractive index varies between 1.526 and 1.540 over a broad range of wavelengths making it suitable for many applications. (c) Energy level diagram of the green‐emitting [tris‐(8‐hydroxyquinoline) aluminum] (Alq3) DNA–CTMA BioLED. The wide band gap of the DNA–CTMA films (4.7 eV) allows the use of an electron blocking layer (EBL) and tuning with different voltages which result in a shift from green to red of the irradiated wavelength. In the blue‐emitting DNA–CTMA BioLED structure the Alq3 emitter layer (EML) was replaced with a single 40 nm (N,N‐bis(naphthalene‐1‐yl)‐N,N‐bis(phenyl)benzidine) (NPB) EML. Alternative structures replaced the DNA–CTMA with polyvinyl carbazole and PMMA as EBLs. ‘Baseline’ green and blue emitting OLEDs were also fabricated with the same structure, except without the EBL. (d) (Left) Current density/voltage relationship and photographs of an OLED prepared from DNA/ polyaniline/Ru(bpy) complex as the hole transport layer and tris(8hydroxyquinolinato)aluminum as the electron transport layer, with inset showing the OLED schematic. The luminescence color changed dramatically with increasing voltage above the bias voltage of 5 V. (Right) Electroluminescence (EL) spectra of OLED at various bias voltages confirming the multicolor voltage‐dependent emission. The inset shows changes in the International Commission on Illumination emission coordinates of the OLED for various bias voltages. (e) EBL DNA–CTMA allows a higher yield in efficiency as evidenced by EL photographs of light emission from (I) Green Alq3 baseline OLED at 25V (707 mA cm−2) −590 cd m−2, 0.35 cd A−1. (II) Green Alq3 BioLED with DNA–CTMA EBL at 25 V (308 mA cm−2) −21, 100 cd m−2, 6.56 cd A−1. (III) Blue NPB baseline OLED at 20 V (460 mA cm−2) −700 cd m−2, 0.14 cd A−1. (IV) Blue NPB BioLED with DNA–CTMA EBL at 20 V (200 mA cm−2) −1500 cd m−2, 0.76 cd A−1. The device emitting area is 2 × 2 mm2. Inclusion of DNA–CTMA into the BioLEDs (II and IV) enhanced efficiency 10× and brightness by 30× compared to traditional OLEDs (I and III). (Reprinted with permission from Ref 101. Copyright 2010 Springer‐Verlag; Reprinted with permission from Ref 102. Copyright 2010 Applied Physics Letters)

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Conductivity of aligned DNA–lipid films with applied mechanical force: (a) 30 kbp DNA strands in a DNA–lipid film were aligned by mechanical stretching after casting and measured for their electro‐conductive properties using two gold electrodes arranged parallel to the direction of alignment. Films were cast with dimensions of 20 × 10 mm, thickness: 30 ± 5 pm, stretched to align (e indicates the direction of elongation), and clamped on a combtype electrode at 25°C in a dry box such that the DNA strands overlapped the distance between electrodes, thus enabling the current to flow. Electrode dimensions were as shown. (b) Measurement of current in the film relative to temperature revealed that the film conductivity decreased dramatically at 40°C. (c) Electric current of a DNA‐aligned cast film at different voltage levels over time on comb‐shaped electrodes at 25°C. (d) A linear dependence of current to voltage at constant temperature was also illustrated. (I) Two electrodes were aligned perpendicular to the direction of DNA strand alignment and measured in atmosphere, (II) the same film of (I) was measured in vacuum at 0.1 mm Hg, and (III) two electrodes were placed parallel to the direction of DNA strand alignment both in vacuum and in atmosphere. The linear increase of current with voltage was only observed in the direction of the applied electrical force (circles). When the electrodes were arranged perpendicular to the direction of alignment little to no current was measured (squares). (Reprinted with permission from Ref 56. Copyright 2005 Springer‐Verlag; Reprinted with permission from Ref 78. Copyright 2009 Hindawi Publishing Corporation; Reprinted with permission from Ref 82. Copyright 1998 American Chemical Society; Reprinted with permission from Ref 88. Copyright 2001 The Royal Society of Chemistry; Reprinted with permission from Ref 89. Copyright 1998 Elsevier Science Limited)

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Studies of DNA‐surfactant films cast from organic solvents. (a) Scheme of film preparation from a DNA–DDAB complex cast from an isopropanol solution. (b)–(d) DNA–DDAB film analysis by (b) FT‐IR, (c) AFM, and (d) WAXS indicating a structural transition dependent on the environmental water content. (b) Full FT‐IR spectrum of a DNA–DDAB film in the dry state. Specific absorbance bands are given for DDAB (left insert) and the DNA bases (right insert) in the DNA–DDAB film in the dry and wet states as well as control spectra for ssDNA, dsDNA, and DDAB alone. (c) AFM image of a dry DNA–DDAB film. (d) WAXS data of a DNA–DDAB film in the dry and wet states indicating the change in lamellar spacing that takes place. (e) Schematic of the film structure indicating the structural change that takes place upon wetting and drying the film. (Reprinted with permission from Ref 48. Copyright 2009 American Chemical Society). (f) Preparation of DNA–lipid films and X‐ray analysis by Okahata et al. (Reprinted with permission from Ref 56. Copyright 2005 Topics in Current Chemistry). X‐ray diffraction patterns are shown of (I) the as‐cast film, (II), the stretched film exposed to moisture, and (III) the stretched dry film. Closed arrows show the incident X‐ray beam while the open, two‐headed arrows show the direction of film stretching. X‐ray photographs of (I) and (III) were enlarged to show the diffraction in greater detail. The DNA strands in as‐cast films were found to align with respect to each other in both wet and dry states as the film was stretched. (Reprinted with permission from Ref 57. Copyright 1996 American Chemical Society; Reprinted with permission from Ref 60. Copyright 2000 Japan Academy of Sciences; Reprinted with permission from Ref 78. Copyright 1998 American Chemical Society)

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Novel honeycomb structures which can be prepared with DNA–lipid or DNA–polymer films. (a) Proposed mechanism of the formation of honeycomb‐structured porous polymer films. (I) Solvent from a polymer solution evaporates; (II) water droplets condense due to the cold solution surface temperature; (III) water droplets assume hexagonal closed packing on the solution surface; (IV) polymer precipitates at the interface, preventing coagulation of water droplets; (V) scanning electron microscope (SEM) image of a porous star polymer film after evaporation of solvent and water, inset: surface removed by adhesive tape. (Reprinted with permission from Ref 63. Copyright 2002 CSIRO). (b) Honeycomb‐patterned films created by casting a DNA–lipid complex from organic solvent under high humidity using the breath figure technique. (VI–X) SEM images of the film prepared at different concentrations: (VI) 0.5 mg/mL, (VII) 1 mg/mL, and corresponding atomic force microscopy (AFM) images of (VIII) 0.5 mg/mL, (IX) 1 mg/mL, and (X) the local magnification of (IX). (Reprinted with permission from Ref 62. Copyright 2009 American Chemical Society)

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Types of surfactants and film preparation methods. (a) and (b) So far only a limited number of surfactants (analogous to lipids) are known to be able to form self‐standing DNA–lipid/surfactant films cast from organic solvents. These can be separated into two main categories: (a) surfactants with a quaternary amine headgroup and (b) amino acid‐derived surfactants. Esters of amino acids like alanine, glycine, and glutamic acid are known to be less toxic compared to the quaternary amine‐based surfactants but are more easily degraded in vivo due to hydrolysis by enzymes.50,52–58 (c) Dry samples of high molecular weight DNA (30 kbp, left) and low (500 bps right). (Reprinted with permission from Ref 56. Copyright 2005 Springer‐Verlag). The length or type of NA used is not thought to have an appreciable effect on the solubility of complexes.47 (d) Scheme for the formation of a DNA–lipid complex in aqueous media. (e) To prepare films the dry DNA–lipid complex (left) is either dissolved in organic solvent(s) and cast on a solid substrate (bottom right) or formed by hot embossing under pressure at increased temperature (top right), allowing custom shaping of the films, similar to what can be achieved for plastic materials, without the use of organic solvents which may cause some toxicity in vivo. The films produced are similar in appearance, although the embossing method is considered superior for producing a thick uniform film. (Reprinted with permission from Ref 56. Copyright 2005 Springer‐Verlag)

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