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WIREs Nanomed Nanobiotechnol
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Complex‐shaped microbial biominerals for nanotechnology

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Single‐celled microorganisms such as diatoms and coccolithophores produce inorganic microparticles with genetically controlled hierarchical nanopatterns. Besides serving as paradigms to inspire new routes for materials synthesis, biominerals themselves, particularly diatom biosilica, are increasingly utilized as templates for the synthesis of novel functional materials. Over the past decade, a large variety of methods have been established that allow not only for the attachment or coating of desired materials onto diatom biosilica but also for complete chemical conversion without altering the characteristic micro‐ and nanoscale morphology. Examples include the synthesis of materials for photonics (surface‐enhanced Raman spectroscopy,SERS, extraordinary optical transmission, EOT), ultraresponsive and sensitive gas sensors, gas storage materials, and highly active catalysts. More recently, emerging insight into the cellular mechanisms of biosilica formation has enabled the in vivo functionalization of diatom biosilica through advanced cultivation techniques and genetic engineering. As a naturally renewable material, biominerals hold the promise of serving as an inexpensive and easily available resource for a future nanotechnology‐based industry. WIREs Nanomed Nanobiotechnol 2014, 6:615–627. doi: 10.1002/wnan.1284 This article is categorized under: Nanotechnology Approaches to Biology > Nanoscale Systems in Biology
Electron microscopy images of biominerals produced by eukaryotic microorganisms. The top row shows biominerals of individual cells, in the bottom row details of the biomineral are shown. Diatoms: Stephanopyxis turris, Eucampia zodiacus, Thalassiosira pseudonana; chrysophyte: Mallomonas cf. crassisquama (courtesy of http://www.chrysophytes.eu/), Coccolithophorid: Emiliania huxleyi [the cell image (top) is courtesy of A. Scheffel, MPI of Molecular Plant Physiology, Potsdam, Germany; the image of the scale (bottom) is courtesy of Jeremy Young, Natural History Museum, London, UK].
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LiDSI method for immobilization of enzymes in diatom biosilica frustules. The diatom cell is shown in cross section. Silica elements are depicted in black, and the protoplast is shown in gray. For simplicity no intracellular organelles are shown. Genomic DNA is depicted by a chromosome symbol. For incorporation of a desired enzyme into the frustule, the genome of the diatom is genetically engineered to incorporate a synthetic fusion gene (orange + green bar) into the genome. The gene encodes a protein consisting of two domains: the enzyme of interest (green oval) and a silaffin‐3‐derived fragment (orange oval). When the fusion protein is expressed by the cell the sillaffin‐derived segment is recognized by the cell for incorporation into the silica thus anchoring the fusion protein stably into the frustule.
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Hydrogen gas detection with a single titania‐coated diatom frustule. (a) Secondary electron image of a titania‐coated Aulacoseira frustule with Pt electrode connections. (b) Temporal current response upon exposure to gas mixtures with different hydrogen concentrations. (Reprinted with permission from Ref . Copyright 2009 American Chemical Society.)
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Catalytic activity of glucose oxidase attached to diatom frustules, frustule‐derived replicas, and reference materials. The schematic shows the chemical reaction catalyzed by frustule attached glucose oxidase molecules. The glucose oxidase was modified with the peptide protamine to enable electrostatic attachment to negatively charged surfaces. The table lists for each material the enzyme‐binding capacity (‘enzyme loading’) and the catalytic activity per gram of bound enzyme (‘specific activity’) in continuous flow‐through catalysis. The materials were unmodified diatomite (‘Diatom‐SiO2’), carbon replicas of diatomite functionalized with carboxylate groups (‘Diatom‐C’), gold‐bearing replicas of diatomite functionalized with mercaptopropionic acid (‘Diatom‐Au’), commercially available carbon black functionalized with carboxylate groups (‘com‐C’), and commercially available gold NPs functionalized with mercaptopropionic acid (‘com‐Au’). (Reprinted with permission from Ref )
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Biosynthesis of the frustule (schematic). Diatom cells are shown in cross section. For simplicity, intracellular organelles other than the SDV are not shown. Cells arranged in the circle show different stages of the cell cycle: (1) Shortly before cytokinesis (i.e., division of the protoplast) the cell has the maximum number of girdle bands; (2) immediately after cytokinesis new biosilica (red) is formed in each sibling protoplast inside a valve SDV (yellow); the SDV expands as more and more silica is deposited; (3) at the final stage, each SDV contains a fully developed valve; (4) the newly formed valves are deposited in the cleavage furrow on the surface of each protoplast by exocytosis of the SDVs; (5) the sibling cells have separated (for simplicity only one of the sibling cells is shown); (6, 7) expansion of the protoplast volume requires the synthesis of new silica (red) inside girdle band SDVs (yellow); each girdle band is synthesized in a separate SDV, and after exocytosis is added to the newly formed valve until the maximum cell volume is reached.
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General schemes showing the structure of the biosilica‐NP arrays: layer‐by‐layer assembly (left) and covalent coupling (right). Both assembly methods have been applied in order to prepare diatom‐templated arrays of noble metal NPs (Pt, Au, Ag) as well as semiconductor NPs (CdTe). Surface‐modified diatom biosilica exhibits promising catalytic properties. Pt‐NPs attached to diatom biosilica exhibited a 10 times higher catalytic activity than the same NPs in pure colloidal solution (see Table ). The assembly of silver NPs on diatom biosilica enabled the sensitive detection of surface‐attached organic molecules through SERS. Maximum signal enhancement factors of about 2 × 103 were observed. (Reprinted with permission from Ref . Copyright 2012 Wiley‐VCH.)
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Silica formation in vitro by mixtures of silaffins and long‐chain polyamines from T. pseudonana. The experiments have been performed at pH 5.5 in sodium acetate buffer in the absence of inorganic phosphate. Silicic acid (100 mM final concentration) was added from a freshly hydrolyzed solution of teramethoxysilane (in 1 mM HCl). At these conditions, neither long‐chain polyamines alone nor silaffins alone are able to form silica within the assay time (10 min). (a) Quantity of silica produced after 10 min reaction time using a constant amount of long‐chain polyamines and varying amounts of silaffins tpSil1/2L, tpSil1/2H, and tpSil3. (b) Secondary electron microscopy images of the silica formed by mixtures of long‐chain polyamines with the silaffin indicated on the image. (Reprinted with permission from Ref . Copyright 2004 The American Society for Biochemistry and Molecular Biology, Inc.)
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