Article pubs.acs.org/Biomac
Silica Precipitation by Synthetic Minicollagens Felix Weiher,† Michaela Schatz,‡ Claudia Steinem,*,‡ and Armin Geyer*,† †
Faculty of Chemistry, Philipps University Marburg, Hans-Meerwein-Straße, 35032 Marburg, Germany Institute of Organic and Biomolecular Chemistry, University of Göttingen, Tammannstraße 2, 37077 Göttingen, Germany
‡
S Supporting Information *
ABSTRACT: Oligomeric Pro-Hyp-Gly- (POG-) peptides, wherein the collagenous triple helix is supported by C-terminal capping, exhibit silica precipitation properties (O, Hyp = (2S,4R)hydroxyproline). As quantified by a molybdate assay, the length of the covalently tethered triple helix (number of POG units) determines the amount of amorphous silica obtained from silicic acid solution. Although lacking charged side chains, the synthetic collagens precipitate large quantities of silicic acid resulting in micrometersized spheres of varying surface morphologies as analyzed by scanning electron microscopy. Similar precipitation efficiencies on a fast time scale of less than 10 min were previously described only for biogenic diatom proteins and sponge collagen, respectively, which have a considerably higher structural complexity and limited accessibility. The minicollagens described here provide an unexpected alternative to the widely used precipitation conditions, which generally depend on (poly-)amines in phosphate buffer. Collagen can form intimate connections with inorganic matter. Hence, silica-enclosed collagens have promising perspectives as composite materials.
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INTRODUCTION Products derived from molecular recognition of soluble biomolecules and inorganic materials have attracted a lot of interest during recent years.1−9 Nanopattering of silica with protein templates, for example, can produce objects of arbitrary shape.10 Further tailoring of material properties is expected from mineralization products obtained from functional proteins. These are characterized by peptide repeats displaying complementary surface polarities to bind the charge pattern of the inorganic matter. Yet, diatom proteins and synthetic mimics thereof are typically used in the presence of phosphate ions.11,12 Here, we present a new class of synthetic peptides that combines synthetic accessibility with an extraordinary silicaprecipitating ability, which is independent of divalent anions during the silification process. Biogenic silica is an in/organic composite material that is known from the microporous frustules of diatoms and from the flexible spicules produced by glass sponges and demosponges. Such hierarchically assembled composites of silica associated with biopolymers are much more elaborate than current technical applications of nanosilica,13 which are obtained from precipitation of silicic acid with cationic tensides.14,15 Diatoms invest considerable effort in the biosynthesis of peptides and proteins of their biosilification machinery.16 Twelve out of 15 amino acids in silaffin1-A1,12 for example, are post-translationally modified either as phosphoserines or as oligopropylene aminolysines.17 The currently accepted molecular mechanism of silica precipitation relies primarily on electrostatic interactions. Components such as amphiphilic silaffins, acidic silacidins, basic oligopropylene amines, and chitin steer the localized silica precipitation in vivo.18,19 The secondary © 2013 American Chemical Society
structure of the polycationic polylysine is known to influence the morphology of the silica precipitate.20 Successful silification experiments were also performed with spider proteins, fibers, nematic phases, and other mesoscopic structures.21−27 Macroscopic collagen fibers have received much attention, but the role of charged amino acid side chains, which are indispensable for the activity of the soluble components listed above, had not been systematically investigated until now.28−31 Glass sponges produce spicules that are a composite material of silicic acid and collagen (among other peptides) and which act as glass fibers due to their extraordinary flexibility and optical properties. A controversially discussed subject in this context is the question of whether collagen is causatively involved in the biosilification process or whether collagen is only associated with the mineral in the biosilica spicules, while the mineralization process itself is catalyzed by silicatein and other biopolymers.31,32 Our in vitro experiments with synthetic collagens were not intended to address this question but to identify the principal requirements that separate peptides with biomineralization properties from incompetent peptides. Synthetic peptides with at least five Pro-Hyp-Gly repeats show temperature dependent cooperative triple helix formation and are therefore well-accepted minimal models of natural collagens.33,34 Covalently capped collagens, as described in the literature, have flexible spacers between the peptides and the capping (Figure 1).35,36 In order to keep the spacing between POG repeats and cap as short as possible, the three POG Received: November 8, 2012 Revised: January 10, 2013 Published: January 31, 2013 683
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Scheme 1. Compounds 1−9 Synthesized by SPPS on Chlorotrityl Resin (Black Sphere)a
Figure 1. (a) Peptides consisting of Pro-Hyp-Gly repeats form collagen-type triple helices that disintegrate cooperatively above the melting temperature Tm. The cooperativity of the process can be quantified from circular dichroism (CD) spectra (available in the Supporting Information). (b) A carboxy-terminal capping that fits the register of the collagen-type triple helix accelerates the kinetics of helix formation and prevents the fraying of terminal amino acids. This stabilizing effect is measured as a significant increase of Tm and a fast response in the CD spectra without melting hysteresis. (c) A fourth peptide strand does not further raise Tm.
strands were linked directly to a Lys−Lys (or Orn−Orn; ornithine) dipeptide (Scheme 1). Melting temperatures characterize the structural fitting between capping and the register of the C3-skrew axis of collagen.37 In the following, we will describe the analysis of the precipitation properties of canonical (three) and noncanonical (two or four) Pro-Hyp-Gly peptide chains of various lengths. We observe a fast and efficient silica precipitation with synthetic small molecules of molecular weights below 10 kDa and without being restricted to highly charged peptides in phosphate buffers.
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EXPERIMENTAL SECTION
One SPPS cycle consisted of fluorenylmethoxycarbonyl (Fmoc) deprotection (25% piperidine in dimethylformamide; DMF), coupling of Fmoc-Pro-Hyp(OtBu)-Gly-OH (coupling reagent 2-(1H-benzotriazole-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate; HBTU) and intermediate washing operations. 4,4-Dimethyl-2,6dioxo-cyclohexylidene (Dde) served as a second amino protecting group for the trimeric minicollagens 3−8. The capping is assembled from diamino acids lysine (m = 1) or ornithine (m = 0), respectively. a
Fmoc Solid Phase Peptide Synthesis (SPPS) of Minicollagen 1. Fmoc-Pro-Hyp(OtBu)-Gly-OH (1.5 eq to resin loading) was dissolved in dimethylformamide (DMF) and added to the preswelled resin. 2-Chlorotrityl chloride resin with a loading of 0.59 mmol/g of glycine was used for the synthesis. Diisopropylethylamine (DIPEA, 7 equiv) and HBTU (1.5 equiv, dissolved in DMF) were added to the reaction mixture. The reaction was swirled by nitrogen for 30 min. After completion, the solution was drained, and the resin was washed three times with DMF. The Fmoc group was removed shortly after by treatment of the still swelled-resin with 25% piperidine in DMF for 15 min. Then the deprotected resin was washed six times with DMF. This cycle was repeated seven times. After additional washing steps with dichloromethane (DCM), the final cleavage of the peptide from the resin was performed with trifluoroacetic acid (TFA):H2O 95:5 for 20 min. The filtrate was precipitated in Et2O(abs.). After centrifugation, the residue was washed with Et2O and again centrifuged. Fmoc Solid Phase Peptide Synthesis of Minicollagen 2. Fmoc-Lys(Fmoc)−OH (1.5 equiv to resin loading), 1-hydroxybenzotriazole (HOBt, 1.5 equiv) were dissolved in DMF and added to the preswelled resin. 2-Chlorotrityl chloride resin with a loading of 0.60 mmol/g of aminohexanoic acid was used for the synthesis. DIPEA (7 equiv) and HBTU (1.5 equiv dissolved in DMF) were added to the reaction mixture. The reaction was swirled by nitrogen for 30 min. After completion, the solution was drained, and the resin was washed three times with DMF. The Fmoc groups were removed shortly after by treatment of the still swelled-resin with 25% piperidine in DMF for 15 min. Finally, the resin was washed six times with DMF. Fmoc-Pro-Hyp(OtBu)-Gly-OH (2.6 equiv) was dissolved in DMF and added to the still-swelled resin. DIPEA (20 equiv) and HBTU (2.6 equiv, dissolved in DMF) were added to the reaction mixture. The reaction was swirled by nitrogen for 30 min. After completion, the
solution was drained, and the resin was washed three times with DMF. The Fmoc groups were removed shortly after by treatment of the still swelled-resin with 25% piperidine in DMF for 15 min. Then the deprotected resin was washed six times with DMF. This cycle was repeated four times. Afterward, NMP:DMF (1:1), 2.5 M ethylene carbonate, and 1% Triton X100 (NMP = N-methyl-2-pyrrolidone) was used as solvent, and NMP:DMF:piperidine (1:1:1), 1.7 M ethylene carbonate, and 0.7% Triton X100 was used for Fmoc-removal. The reaction cycle was repeated three times. After additional washing steps with DCM, the final cleavage of the peptide from the resin was performed with TFA:H2O (95:5) for 20 min. The filtrate was precipitated in Et2O(abs.). After centrifugation, the residue was washed with Et2O and again centrifuged. General Procedure for Minicollagens 3, 4, and 5 as Fmoc Solid Phase Peptide Synthesis. Fmoc-Lys(Dde)−OH (1.5 equiv to resin loading), HOBt (1.5 equiv) were dissolved in DMF and added to the preswelled resin. 2-Chlorotrityl chloride resin with a loading of 0.69 mmol/g of aminohexanoic acid was used for the synthesis. DIPEA (7 equiv) and HBTU (1.5 equiv, dissolved in DMF) were added to the reaction mixture. The reaction was swirled by nitrogen for 30 min. After completion, the solution was drained, and the resin was washed three times with DMF. The Fmoc group was removed shortly after by 684
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incubated at 20 °C for exactly 15 min and immediately used as a source of monosilicic/disilicic acid. b. In Vitro Precipitation of Silicic Acid and Scanning Electron Microscopy (SEM) Analysis. Silica formation was initiated by the addition of 25 μL of 90 mM silicic acid, prepared as described above, to the peptide in 30 mM phosphate buffer, pH 6.0, if not indicated otherwise. After 10 min at room temperature, the silica was spinned down by centrifugation (5 min, 14.000 × g). The precipitate was washed three times with water, then suspended in water, applied to an aluminum sample holder, and air-dried. Silica precipitates were analyzed without sputter-coating with a LEO Gemini 1530 fieldemission scanning electron microscope (Zeiss) using 5kV acceleration voltage. c. Concentration Assay. Precipitated silica was dissolved in 2 M NaOH for 60 min at room temperature and quantified by a modified β-silicomolybdate method. HCl (1.35 mL; 37%) was dissolved in 40.3 mL of H2O, and 774.2 mg of [(NH4)6Mo7O24 × 4 H2O] was dissolved in 9.7 mL of water. Both solutions were mixed, and the pH was adjusted to 1.04 with 2 M NaOH (molybdate solution). The sample solution containing dissolved silica (0, 20, or 40 μL) and a corresponding volume of 2 M NaOH (0, 20, or 40 μL), 160 μL H2O, and 800 μL molybdate solution was added, and the absorbance of the solution was monitored at a wavelength of 370 nm. A silicon atomic absorption standard solution was used to generate calibration curves.
treatment of the still swelled-resin with 25% piperidine in DMF for 15 min. Finally, the resin was washed six times with DMF. Fmoc-Lys(Fmoc)−OH (1.5 equiv to resin loading) and HOBt (1.5 equiv) were dissolved in DMF and added to the still-swelled resin. DIPEA (7 equiv) and HBTU (1.5 equiv, dissolved in DMF) were added to the reaction mixture. The reaction was swirled by nitrogen for 30 min. After completion, the solution was drained, and the resin was washed three times with DMF. The Dde group was removed by treatment of the resin with a solution of 2% hydrazine in DMF, which was added to the still swelled-resin and swirled for 2 × 20 min. Afterward, the resin was washed six times with DMF. The Fmoc groups were removed shortly after by treatment of the still swelled-resin with 25% piperidine in DMF for 15 min. Finally, the resin was washed six times with DMF. Fmoc-Pro-Hyp(OtBu)-Gly-OH (4.0 equiv to resin loading) was dissolved in DMF and added to the still-swelled resin. DIPEA (20 equiv) and HBTU (4.0 equiv, dissolved in DMF) were added to the reaction mixture. The reaction was swirled by nitrogen for 30 min. After completion, the solution was drained, and the resin was washed three times with DMF. The Fmoc groups were removed shortly after by treatment of the still swelled-resin with 25% piperidine in DMF for 15 min. Then the deprotected resin was washed six times with DMF. This cycle was repeated four times. Afterward NMP:DMF (1:1), 2.5 M ethylene carbonate, and 1% Triton X100 was used as solvent, and NMP:DMF:piperidine (1:1:1), 1.7 M ethylene carbonate, and 0.7% Triton X100 was used for Fmocremoval. The reaction cycle was repeated as often as necessary. After additional washing steps with DCM, the final cleavage of the peptide from the resin was performed with TFA:H2O (95:5) for 20 min. The filtrate was precipitated in Et2O(abs.). After centrifugation, the residue was washed with Et2O and again centrifuged. Fmoc Solid Phase Peptide Synthesis of Minicollagen 9. Fmoc-Lys(Fmoc)−OH (1.5 equiv to resin loading) and HOBt (1.5 equiv) were dissolved in DMF and added to the preswelled resin. 2Chlorotrityl chloride resin with a loading of 0.46 mmol/g of aminohexanoic acid was used for the synthesis. DIPEA (7 equiv) and HBTU (1.5 equiv dissolved in DMF) were added to the reaction mixture. The reaction was swirled by nitrogen for 30 min. After completion, the solution was drained, and the resin was washed three times with DMF. The Fmoc group was removed shortly after by treatment of the still swelled-resin with 25% piperidine in DMF for 15 min. Finally, the resin was washed six times with DMF. Fmoc-Lys(Fmoc)−OH (3 equiv to resin loading) and HOBt (3 equiv) were dissolved in DMF and added to the still-swelled resin. DIPEA (12 equiv) and HBTU (3 equiv, dissolved in DMF) were added to the reaction mixture. The reaction was swirled by nitrogen for 30 min. After completion, the solution was drained, and the resin was washed three times with DMF. The Fmoc groups were removed shortly after by treatment of the still swelled-resin with 25% piperidine in DMF for 2 × 15 min. Finally, the resin was washed six times with DMF. Fmoc-Pro-Hyp(OtBu)-Gly-OH (5.2 equiv) was dissolved in DMF and added to the still-swelled resin. DIPEA (25 equiv) and HBTU (5.2 equiv, dissolved in DMF) were added to the reaction mixture. The reaction was swirled by nitrogen for 30 min. After completion, the solution was drained, and the resin was washed three times with DMF. The Fmoc group was removed shortly after by treatment of the still swelled-resin with 25% piperidine in DMF for 15 min. Then the deprotected resin was washed six times with DMF. This cycle was repeated four times. Afterward, NMP:DMF (1:1), 2.5 M ethylene carbonate, and 1% Triton X100) was used as solvent, and NMP:DMF:piperidine (1:1:1), 1.7 M ethylene carbonate, and 0.7% Triton X100 for Fmoc-removal. The reaction cycle was repeated three times. After additional washing steps with DCM, the final cleavage of the peptide from the resin was performed with TFA:H2O (95:5) for 20 min. The filtrate was precipitated in Et2O(abs.). After centrifugation, the residue was washed with Et2O and again centrifuged. Silica Precipitation Assays. a. Preparation of Silicic Acid. A freshly prepared solution of 1 M tetramethoxysilane in 1 mM HCl was
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RESULTS AND DISCUSSION The peptides were obtained by solid-phase peptide synthesis with Fmoc strategy (Experimental Section). CD spectroscopy revealed a linear dependence of the melting point, estimated according to the two state model,38,39 on the length of the peptides. Each additional POG unit increases the melting temperature by approximately 10 °C. The melting temperature of covalently tethered derivative 3 is 30 °C higher than the melting temperature of monomer 1. The slightly more constrained ornithine based oligomer 6 shows a further increase of 4 °C. These Tm are explained by a mainly entropic influence of the cap. Capping increases Tm of the longer POG repeats by a constant value but fails to promote triple helix formation for very short POG repeats (n < 4) because it does not fit precisely to the register of the triple helix. The noncanonical dimer 2 and the tetramer 9 exhibit Tm values similar to 1 and trimer 5, respectively, thus supporting the idea that an extra strand does not participate in the triple helix formation. Yet, all melting temperatures are high enough to ensure a more than 90% triple helical fold during the silica precipitation experiments, which were performed at room temperature. The pH was kept at a low value of 6.0. At this pH, the acidity of the solution does not affect the melting point of synthetic collagens. To compensate for the different molecular weights of compounds 1−9, the silica precipitation experiments were carried out at similar POG concentrations. Phosphate buffer was chosen to make the experimental results comparable with previous precipitation experiments. Compounds 1−9 were incubated with silicic acid solution at pH 6.0 at room temperature for 10 min. The amount of precipitated silica was subsequently determined by the β-silicomolybdate method.40,41 The amount of SiO2 (mSi) was significantly influenced by the molecular structure (Figure 2). For 1, 2, and 3, which contain the same number of seven POG units, the amount of precipitated silica increases with Tm. An additional forth strand in 9 has a minor influence on Tm, yet further increases the amount of SiO2 precipitation. The number of POG repeats significantly increases Tm (3−8) but has only a minor influence on the amount of precipitated silica. 685
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By SEM images, the structure of the silica precipitates was further elucidated. Figure 3 shows SEM images of silica
Figure 2. Silica precipitation experiments were conducted at 0.28− 0.56 mg POG/mL in 30 mM phosphate buffer with the same precipitation time of 10 min after which the silica mass was quantified. The amount of silica was determined by the molybdate-method, and the Tm was found by CD-spectroscopy.40 Different precipitation conditions were tested for peptide 3. A reduction of the incubation time to 5 min significantly lowers the amount of silica (mSi = 77 μg), while longer incubation times (30 min) do not further increase the quantity of silica (mSi = 118 μg). Peptide 3 in MES buffer yields mSi 115 μg. A mSi less than 10 μg could not be safely quantified and is included as an open rectangle. Figure 3. SEM images of the silica precipitates obtained in the presence of the different minicollagens. (A) 2, (B) 3, (C) 9, (D) 8, (E) 7, (F) 6. The scale bars are 1 μm. The insets are magnifications (10×) illustrating that the silica precipitates appear slightly more irregular in the cases of compounds 2 and 8. The marginal amount of silica precipitated by 1 does not lead to a representative SEM image.
The quantities of precipitated SiO2 in Figure 2 differ significantly, although all peptides are expected to be folded at the precipitation temperature. In order to compensate for different folding kinetics, all peptides were stored in solution at 4 °C and not warmed above room temperature before the SiO2 precipitation experiments. Folding hysteresis of monomers like 1 were extensively characterized in the literature.42 Peptide 2 behaves in a similar way, while hysteresis is not observed in the temperature-dependent nuclear magnetic resonance (NMR) and CD spectra of the trimers 3−8 and the tetramer 9. Tethered peptides 3−9 reach thermodynamic equilibrium much faster than single stranded compound 1. 1 and 2 differ from the other peptides mainly because of their slower kinetics of folding. The amount of silica precipitate linearly scales with the amount of dissolved peptide. Compound 3 for example, which precipitates 125 μg at 0.37 mg POG/mL (0.14 mM POG concentration), precipitates 50 μg at 0.06 mM POG and only 20 μg at the highest investigated dilution of 0.03 mM POG. Cationic charges are known to positively influence the amount of precipitate. The minor contribution of cationic charges makes the mineralization process less dependent on the solvent conditions and by this expands its applicability. Minicollagens precipitate silica independently of the buffer, as shown exemplarily for 2-(N-morpholino)ethanesulfonate (MES) buffer (pH 6.0). Even though it is difficult to compare the absolute amount of precipitated silica obtained for different molecules under different conditions, it is worth mentioning that mSi is in the range of 100 μg per 0.14 mM POG units at pH 6.0, while mSi was determined to be in the 10 μg range per 1 mM nitrogen in the case of polyamines, even though the pH was larger (pH 6.8),12 which generally increases the amount of precipitated silica. In previous studies, it was shown that synthetic silaffin peptides precipitate silica in a concentration-dependent manner in the 10−100 μg regime,12 yet they did not precipitate relevant amounts of silica at pH 5.5.
structures obtained in the presence of 0.14 mM POG units at pH 6.0 at room temperature. In all cases, rather spherical, partly fused micrometer-sized structures were observed. The surface of the silica precipitates shown in Figure 3B obtained in the presence of trimer 3 are more roundish compared to those obtained in the presence of the noncanonical dimer 2. A slightly irregular shape is also found for the precipitates in the presence of the compounds with only five POG repeats like 5 and 8. Compared to silica precipitates obtained in the presence of synthetic long chain polyamines, which are spherical with a size distribution of the spheres in the 100 nm range, the silica structures found here are not perfectly spherical and strongly interconnected. From the size of the structures, which is in the micrometer range, it can be concluded that larger aggregates of the molecules are responsible for silica formation. It is well conceivable that the silification process drives this aggregation leading to interconnected silica structures. Organic molecules can accelerate the condensation of silicic acid to siloxanes. The distribution of oligoamines within the precipitate was investigated before. 21 Other classes of molecules, e.g., collagens, are less detailed characterized. Our results indicate the requirement of supramolecular aggregations during the precipitation process.
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CONCLUSIONS Collagen-type peptides with C-terminal caps precipitate silica in phosphate buffer and in MES buffer. The peptides presented here mediate biosilica formation similar to the in vitro experiments with natural silaffins,43 in spite of a distinctly different composition. In contrast to the excessively charged and highly modified peptides and proteins from diatoms, the 686
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(19) Brunner, E.; Richthammer, P.; Ehrlich, H.; Paasch, S.; Simon, P.; Ueberlein, S.; Van Pée, K.-H. Angew. Chem., Int. Ed. 2009, 48, 9724− 9727. (20) Patwardhan, S. V.; Maheshwari, R.; Mukherjee, N.; Kiick, K. L.; Clarson, S. J. Biomacromolecules 2006, 7, 491−497. (21) Bernecker, A.; Wieneke, R.; Riedel, R.; Seibt, M.; Geyer, A.; Steinem, C. J. Am. Chem. Soc. 2010, 132, 1023−1031. (22) Heinemann, S.; Heinemann, C.; Jäger, M.; Neunzehn, J.; Wiesmann, H. P.; Hanke, T. ACS Appl. Mater. Interfaces 2011, 3, 4323−4231. (23) Niu, L.-N.; Jiao, K.; Qi, Y.-P.; Yiu, C. K. Y.; Ryou, H.; Arola, D. D.; Chen, J.-H.; Breschi, L.; Pashley, D. H.; Tay, F. R. Angew. Chem., Int. Ed. 2011, 50, 11688−11691. (24) Bernecker, A.; Ziomkowska, J.; Heitmüller, S.; Wieneke, R.; Geyer, A.; Steinem, C. Langmuir 2010, 26, 13422−13428. (25) Ehrlich, H.; Worch, H. In Handbook of Biomineralization; Bäuerlein, E., Ed.; Wiley-VCH: Weinheim, Germany, 2007; pp 23−41. (26) Ehrlich, H.; Janussen, D.; Simon, P.; Bazhenov, V. V.; Shapkin, N. P.; Erler, C.; Mertig, M.; Born, R.; Heinemann, S.; Hanke, T.; Worch, H.; Vournakis, J. N. J. Nanomater. 2008, 1−8. (27) Canabady-Rochelle, L. L. S.; Belton, D. J.; Deschaume, O.; Currie, H. A.; Kaplan, D. L.; Perry, C. C. Biomacromolecules 2012, 13, 683−690. (28) Li, Y.; Thula, T. T.; Jee, S.; Perkins, S. L.; Aparicio, C.; Douglas, E. P.; Gower, L. B. Biomacromolecules 2012, 13, 49−59. (29) Eglin, D.; Mosser, G.; Giraud-Guille, M.-M.; Livage, J.; Coradin, T. Soft Matter 2005, 1, 129−131. (30) Hoyer, B.; Bernhardt, A.; Heinemann, S.; Stachel, I.; Meyer, M.; Gelinsky, M. Biomacromolecules 2012, 13, 1059−1066. (31) Ehrlich, H.; Deutzmann, R.; Brunner, E.; Cappellini, E.; Koon, H.; Solazzo, C.; Yang, Y.; Ashford, D.; Thomas-Oates, J.; Lubeck, M.; Baessmann, C.; Langrock, T.; Hoffmann, R.; Wörheide, G.; Reitner, J.; Simon, P.; Tsurkan, M.; Ereskovsky, A. V; Kurek, D.; Bazhenov, V. V; Hunoldt, S.; Mertig, M.; Vyalikh, D. V; Molodtsov, S. L.; Kummer, K.; Worch, H.; Smetacek, V.; Collins, M. J. Nat. Chem. 2010, 2, 1084− 1088. (32) Wang, X.; Schloßmacher, U.; Wiens, M.; Batel, R.; Schröder, H. C.; Müller, W. E. G. FEBS J. 2012, 279, 1721−1736. (33) Shoulders, M. D.; Raines, R. T. Annu. Rev. Biochem. 2009, 78, 929−958. (34) Cai, W.; Wong, D.; Kinberger, G. A.; Kwok, S. W.; Taulane, J. P.; Goodman, M. Bioorg. Chem. 2007, 35, 327−337. (35) Fields, C. G.; Grab, B.; Lauer, J. L.; Fields, G. B. Anal. Biochem. 1995, 231, 57−64. (36) Rump, E. T.; Rijkers, D. T. S.; Hilbers, H. W.; De Groot, P. G.; Liskamp, R. M. J. Chem.Eur. J. 2002, 8, 4613−4621. (37) Li, Y.; Mo, X.; Kim, D.; Yu, S. M. Biopolymers 2011, 95, 94−104. (38) Persikov, A. V; Xu, Y.; Brodsky, B. Protein Sci. 2004, 13, 893− 902. (39) Erdmann, R. S.; Wennemers, H. Org. Biomol. Chem. 2012, 10, 1982−1986. (40) Iler, R. K. The Chemistry of Silica; Wiley-Interscience: New York, 1979. (41) Hurfordl, T. R.; Boltz, D. F. Anal. Chem. 1968, 40, 1966−1969. (42) Mizuno, K.; Boudko, S. P.; Engel, J.; Bächinger, H. P. Biophys. J. 2010, 98, 3004−3014. (43) Belton, D. J.; Deschaume, O.; Perry, C. C. FEBS J. 2012, 279, 1710−1720.
title compounds are assembled from ubiquitous readily accessible amino acids. Minicollagens not only give access to phosphate-free precipitation environments but also add new facets to the currently discussed molecular mechanisms of silica precipitation. Molecular bionics is aimed to reproduce biomimetic function with synthetic organic structures. Further engineering toward larger collagen scaffolds might provide access to successful in vitro production of unique nanostructured silicabased composites.
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ASSOCIATED CONTENT
S Supporting Information *
Details on the synthesis 6−8, analytical data (NMR, MS, and CD spectra) of the peptides and Tm and mSi of 1−9. This material is available free of charge via the Internet at http:// pubs.acs.org.
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected] (C. S.); geyer@staff.uni-marburg.de (A.G.). Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS We would like to thank M. Klingebiel for technical assistance. REFERENCES
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