ISSN 00063509, Biophysics, 2011, Vol. 56, No. 5, pp. 795–802. © Pleiades Publishing, Inc., 2011. Original Russian Text © U.S. Shvyreva, M.N. Tutukina, O.N. Ozoline, 2011, published in Biofizika, 2011, Vol. 56, No. 5, pp. 821–830.
NANOTECHNOLOGIES IN VIVO
Bacterioferritin: Properties, Structural and Functional Organization of the dps Gene Regulatory Region U. S. Shvyrevaa,b, M. N. Tutukinaa, and O. N. Ozolinea a
Institute of Cell Biophysics, Russian Academy of Sciences, Pushchino, Moscow Region, 142290 Russia b Pushchino State University, Pushchino, Moscow Region, 142290 Russia Email:
[email protected] Received June 9, 2011
Abstract—The paper considers the properties of the gene encoding bacterioferritin Dps, which is involved in sequestering iron ions, forms a ferrihydrite core inside the protein cavity, and is a major nucleoid protein. Experimental evidence is presented for the effect of microwave irradiation on the dps gene expression. The structural and functional organization of its regulatory region is analyzed, and the technological prospects of bacterioferritin application for designing new materials with desired properties are discussed. Keywords: bacterioferritin, metalloprotein, nanoparticle, dps promoter, gene expression regulation, micro wave irradiation. DOI: 10.1134/S0006350911050204
INTRODUCTION In the course of evolution, nature has accumulated infinitely many molecules that form complex 3D structures and perform unique functions in the living cell. Study of these molecules open broad possibilities of their use in nanobiotechnology. It is quite natural that ferritins have become objects of such research. These are spherical proteins consisting of 24 subunits of ~18 kDa (Fig. 1). Ferritins are widespread in nature and play an important part in iron homeostasis [1, 2]. These proteins bind the iron ions, which are necessary but toxic for the cell as Fe2+, oxidize them to Fe3+, and accumulate them inside the oligomer cavity as spa tially oriented hydrated oxides (5 Fe2O3 ⋅ H2O) known as ferrihydrites. The overall number of iron oxide mol ecules in the cavity of one ferritin in different estimates is 2500 to 4500, and the size of the ferrihydrite core reaches 8 nm (Fig. 1). Possessing an intrinsic magnetic moment, the ferrihydrite core of ferritins is usually antiferromagnetic [3], but its magnetic properties can be changed upon additional (or alternative) saturation with oxides of other metals. It is for this reason that ferritins may prove very promising in microelectronics. Furthermore, they can be used to obtain some alloys such as CoPt; apoferritin (the protein without a ferri hydrite core) has been proposed for addressed delivery of drugs to cells and tissues. Figure 2 outlines the fields of ferritin application discussed in the literature. Escherichia coli is known to have at least three pro teins of the ferritin family (FtnA, Bfr, and Dps). Struc turally and functionally, FtnA more than others resembles the ferritins of higher organisms [1] but con sists of 24 identical subunits, whereas the 3D structure
of eukaryotic ferritins is usually formed by polypeptide chains of two types, only one type being able to oxidize two Fe2+ ions to Fe3+ using O2. Bfr is also composed of 24 identical subunits and along with the usual ferroxi dase sites has 12 binding sites for heme iron [14]. These sites are at the boundaries between monomers and appear to partake in releasing Fe3+ from the min eralized core. The third bacterioferritin, Dps, in con trast, consists of only 12 identical subunits (Fig. 1). Each Dps subunit also oxidizes two Fe2+ ions to Fe3+ but mainly uses a hydrogen peroxide molecule rather than O2. Apart of mineralization of cytotoxic Fe2+, this favors removal of H2O2 from the cytoplasm and thereby attenuates the production of reactive oxygen species. The ferrihydrite core of Dps consists of ~500 iron oxides [15] and is considerably smaller than in other ferritins (Fig. 1). The Dps monomer comprises 167 amino acid residues (vs. 165 in FtnA and 158 in Bfr) and has a molecular wt of 18712. The unique property of Dps versus other ferritins is its ability to tightly bind with DNA. During exponential growth, one E. coli cell contains ~6000 Dps molecules, whereas in the late stationary phase their number rises to ~180000 [16] and practically all of them are associ ated with the genome. This high affinity for DNA offers a prospect of immobilizing bacterioferritin on shapeforming DNA scaffolds, i.e., enabling self assembly of any preset spatial arrays of ferromagnetic particles of specified composition and size. We have found that microwave irradiation of sta tionaryphase bacterial cells results in accumulation of dps mRNA [17]. Most probably, this is due to compen satory activation of dps gene transcription in response
795
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SHVYREVA et al. Bfr
~4500 Fe3+
80 Å
120 Å Dps
~500 Fe3+
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90 Å Fig. 1. The 3D structures of bacterioferritins Bfr and Dps of E. coli (RCSB PDB codes 1brf and 1dps respectively), with sizes of their molecules and ferrihydrite cores.
to microwaveinduced alterations in nucleoid struc ture rather than to the influence of the electromag netic field on dps mRNA stability. If so, formation of DNA–protein complexes can be made controllable. Here we present data on the influence of microwaves on dps expression and discuss the regulatory mecha nisms that may be involved. EXPERIMENTAL Bacterial culture and irradiation. A laboratory strain Escherichia coli Top 10 (Invitrogen, USA) was grown in LB medium at 37°C to late stationary phase (16 h). Two 20mL aliquots of culture from one flask were trans ferred to special microwavetransparent vials and placed into a special thermostat [18] with two chambers and a common bottom shaker to ensure identical incu bation conditions. The control sample was placed into the electromagnetically shielded chamber; the test sam ple in the exposure chamber was 80 cm away from the horn of a sweep frequency generator Ya2R76/2 (fre quency range 8.15–18 GHz, sweep time 1 ms,) the mean energy flux density in the center of the vial was 1 μW/cm2, the exposure lasted 1–3 h. Effect of microwave irradiation on dps expression was assayed by the transcriptional activity of the gene promoter region. DNA fragments containing the dps regulatory region of different size were inserted before
the green fluorescent protein (GFP) gene in pET28b EGFP between BglII and XbaI sites, using enzymes from Fermentas (Lithuania) according to manufac turer’s protocols. All inserts were checked by direct sequencing. Plasmidtransformed E. coli Top10 cells were exposed to microwaves, and the promoter activ ity was judged about by accumulation of GFP. Fluo rescence of the latter was measured in cell suspension (MultiScanPlus, LabSystems, Finland, λ = 492 nm) and in the gel upon SDSPAGE; the samples were SDStreated but not boiled, to spare the GFP chro mophore. Control and microwaveexposed cells were harvested at 10000 g for 10 min, suspended in TE buffer (20 mM TrisHCl, pH 8.0, 0.5 mM EDTA), treated with lysozyme (1 mg/mL) on ice for 20 min, and sonicated 4 × 15 s, avoiding tube heating. The lysate was centrifuged at 10000 g for 10 min, and solu ble proteins were precipitated with saturated ammo nium sulfate. The pellet was dissolved in TE buffer, equalizing the protein concentrations in control and test samples. Aliquots with 10 μg protein were loaded onto 7.5% PAG with 0.1% SDS. Right after electro phoresis the gels were photographed in transmitted light (λ = 254 nm) and stained with Coomassie R250. The electrophoregrams were quantitated densitomet rically. BIOPHYSICS
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BACTERIOFERRITIN: DPS REGULATORY REGION Biomedical nanorobots for drug delivery [4, 5]
Bioreactor for making a CoPt alloy, a roomtemperature ferromagnetic [13]
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Liquid protein favoring oxygen delivery to tissue lesion and fast healing [6] Immunomagnetic cell markers [7]
Ferritin
Nanoparticles of 7–12 nm diameter Semiconducting materials [12]
Nanomagnetic memory devices [Nanomagnetics Ltd, UK]
Data Ink for hard disk coating [Nanomagnetics Ltd, UK]
Multilayer bionanobatteries [8]
Novel logic elements for supercomputers [11] Quantum electronic devices [10]
Filters for terahertz electromagnetic radiation [9]
Fig. 2. Fields of application of ferritin molecules, indicating publications or companies engaged in research.
Analysis of the dps regulatory region. To find poten tial transcription initiation sites, use was made of the promoter search algorithm PlatProm [19]. The ability of the predicted promoters to interact with RNA poly merase (RNAP) in vitro was assessed by a retardation assay in 4% PAG. Formation of “open” complexes was checked by permanganate footprinting. For this pur pose, PCR with primers 5'ATGCAGATCT TCTCGCTACTTTTC3' (1), 5'TCCTCTAGAT GTTATGTCCCAGT3' (2), 5'GGAAGATCTTC CTCGGAGAAACACT3' (3) was run to produce promotercontaining DNA fragments, which were 5' endlabeled in one chain with 32P using T4 polynucle otide kinase (Fermentas). Amplicons were extracted from gel routinely [20]. RNAP complexes were allowed to form for 30 min at 37°C in a standard buffer of 50 mM TrisHCl (pH 8.0), 0.1 mM EDTA, 0.1 mM DTT, 10 mM MgCl2, 50 mM NaCl, 5 mg/mL BSA (Sigma) containing 0.2 pmol of [32P]DNA fragment and 0.4–0.8 pmol of σ70RNAP (Sigma). For the gel retardation assay, the sample before loading was sup plemented with heparin (20 μg/mL). Permanganate footprinting revealing unpaired thymines was per formed as in [21]. The size markers were obtained by Maxam–Gilbert Gspecific cleavage [20] of the cor responding fragments. The products were resolved in 8% PAG with 8 M urea and autoradiographed. To determine the promoter activity in vivo, four frag ments were synthesized using the above primers (1, 2, 3) and 5'ATATCTAGATATATAAAGACGGT GTA3' (4), 5'CTAATCTAGATTCAATGAGTTA GATA3' (5), and then cloned in pET28bEGFP as above. GFP fluorescence was measured in E. coli col BIOPHYSICS
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onies grown on standard LB Plates with kanamycin (20 μg/mL), using a Leica DM6000B fluorescence microscope excitation at 480 and visualization at 510 nm. The data were processed with ImageJ. Primer extension. Total cell RNA was isolated as in [22]. Extension of a 32Plabeled primer was performed as in [23]. Then samples were treated with RNase A (Fermentas; 10 U, 37°C, 30 min) and precipitated with 3 vols. of icecold 96% ethanol, 0.3 M sodium acetate. The pellet was washed with 70% ethanol, dried, and dissolved in 5 μL of 98% formamide with 8 mM NaOH, 4 mM EDTA. The cDNA products were resolved in 8% PAG with 8 M urea and autorad iographed. RESULTS It has earlier been established that an electromag netic field in the 8.15–18 GHz range of 1 μW/cm2 intensity differentially affects the synthesis of a num ber of proteins [18] and mRNAs [17] in stationary phase E. coli. The amount of dps mRNA thereby increased by some 70%. This could be due to induced dps transcription or to inhibited hydrolysis of mRNA, the halflife of which depending on growth conditions is 8.6–12.5 min [24]. If the former suggestion is true, microwaves should stimulate the dps promoter activity. To check this, we integrated the dps regulatory region in pET28bEGFP before a GFP reporter gene. Irradiation of E. coli Top10 transformed with this plas mid indeed led to accumulation of GFP (Fig. 3a). This effect did not appreciably depend on the duration of exposure (varied from 1 to 3 h) but in different experi
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Fig. 3. Examples of electrophoregrams of soluble proteins from E. coli transformed with pET28bEGFP, where gfp was tran scribed from (a) dps or (b) hns promoters. In panel (a), the promoter fragments were obtained with (left) primers 2 and 3 or (right) primers 1 and 2. Lanes 1, 3, 5, nonirradiated cells; lanes 2, 4, 6, cells exposed to microwaves for 3 h. Gels were photographed in UV light and stained as indicated above.
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Fig. 4. The functional organization of the dps regulatory region. Gray bars represent genes (reading direction indicated). Vertical bars mark the known (Pdps) and PlatPrompredicted (p < 0.00005) transcription initiation points for sense (P1, P '1 , P2, P3) and antisense (Pa1, Pa2, Pa3) RNAs, bar height corresponding to the calculated similarity with idealized promoter (ordinate). Short arrows mark the position of primers (5' to 3'). Empty boxes denote binding sites for specified regulatory proteins [25, 26].
ments ranged from 6 to 41% (mean 23.8% after 3 h) by densitometric data. Irradiation did not change the GFP production if the plasmid contained a promoter of the hns gene encoding another nucleoid protein (Fig. 3b). Thus, although the microwaveinduced activation of reporter protein production was weaker than that of dps mRNA synthesis [17], this is evidence in favor of a specific change in dps expression. The promoter sequences inserted into the plasmid proved sufficient to respond to a regulatory signal. As the next
step, we explored the structural and functional organi zation of the dps regulatory region. According to the literature data, dps is transcribed monocistronically from the start point located 39 bp upstream of the initiation codon [27]. Transcription from Pdps can be initiated by RNAP with σ38 or σ70 as the promoter specificity factor. However, in the gene regulatory region the PlatProm algorithm predicts four more promoters for dps mRNA and three (Pa1, Pa2, Pa3) for antisense transcription (Fig. 4). The most BIOPHYSICS
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BACTERIOFERRITIN: DPS REGULATORY REGION (a) G
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Fig. 5. (a) RNAP complexes formed with a linear fragment obtained by PCR with E. coli genomic DNA and primers 2+3. The enzyme/DNA molar ration is indicated above the lanes. (b) Lane 2, cDNA obtained by reverse transcription with total cell RNA and a 32Plabled primer; lanes 3–5, permanganate footprinting of RNAP–promoter complexes; arrows and wavy line mark bands corresponding to modified thymines. Lanes 1 and 6, Gspecific cleavage of the corresponding DNA fragments; numbers give the positions of the Gs relative to the ATG codon.
probable start sites for sense transcription are 85 (P1), 116 ( P '1 ), 196 (P2), and 261 (P3) bp upstream. Since the observed induction of gene expression could be due to any of these promoters, we examined their abil ity to interact with RNAP in vitro and in vivo. A linear DNA fragment of the dps regulatory region containing all potential promoters (except Pa1) effi ciently interacted with RNAP even at a very low enzyme/DNA ratio (< 0.25 per promoterlike site) to yield several complexes differing in electrophoretic mobility (Fig. 5a), which was indicative of RNAP binding with the predicted promoters. Footprinting with potassium permanganate (Fig. 5b, lanes 3–6) revealed several locally denatured regions; this is usu ally regarded as evidence for transition of RNAP–pro moter complexes into an “open” state ready for rapid transcription initiation. One of these regions corre sponds to the known Pdps whereas two others, to pre dicted P1 and P '1. In the opposite strand, we also regis tered permanganateaccessible thymines near the pre dicted P2 (not shown). Nonetheless, reverse transcription revealed no cDNA products correspond ing to P1, P '1 , or P2 start points (Fig. 5b, lane 2). No such products were also found among cDNAs sub jected to direct sequencing [28]. In accord with this, we could not detect any synthesis of the reporter GFP (Fig. 6а) when its gene was put under control of P3 or BIOPHYSICS
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P3–P2. Insertion of the regulatory region comprising P2, P '1 , P1, and Pdps before gfp resulted in efficient expression: fluorescence of the colonies of thus trans formed cells exceeded the control 5.5 times. However, addition of P3 resulted in more than 9fold activation of reporter expression (Fig. 6b). Thus, the genome region between primers 1 and 3 (Fig. 4) interacts with RNAP (footprinting data) but does not provide gene transcription (RT data) either in the sense (Fig. 5b, Fig. 6a [28]) or in the antisense direction [28]. There with, it is important for maximal gene expression (Fig. 6b). DISCUSSION Dps was first reported in E. coli as DNAbinding protein of starved cells [29]. Until recently, the great interest in this protein has been due just to this func tion of maintaining the genomic DNA conformation corresponding to the growth rate. Along with Dps, this process involves several more proteins: IHF, Fis, HU, StpA, HNS, CbpB, CbpA, DnaA, Lrp, and IciA [30], as well as the proteins of the transcription apparatus and the factors controlling genome expression. They all influence the nucleoid structure, but in the station ary phase it is Dps that is the main architectural factor for DNA [16]. Therefore, a change in its expression
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Fig. 6. Fluorescence of colonies of cells carrying pET28bEGFP without a promoter before gfp (control) or the plasmid contain ing the true and predicted promoters in the dps regulatory region. Fragments with promoter sets specified above the photographs were obtained with the following primer pairs: (P3) 3+4; (P3P2) 3+5; (P2 P '1 P1Pdps) 1+2; (P3P2 P '1 P1Pdps) 3+2 (see Fig. 4). Numbers on the photographs reflect the fluorescence intensity. Pronounced fluorescence hinders focusing, so in the last two panels the colonies are seen less sharply.
under the action of microwaves must necessarily tell on the nucleoid structure. It is believed that the main functional unit of Dps like proteins is a dodecamer (Fig. 1), though upon a small shift of pH from neutral it breaks down into dimers, and in acidic medium, even into monomers [31], which at least partly retain ferroxidase and DNAbinding activity. Interaction with DNA in most cases involves the unordered Nterminal segment. In the E. coli Dps, it protrudes from the hydrophobic core and, owing to three lysine residues, can bind with the negatively charged DNA. Upon removal of all three lysines, Dps loses the ability to cause DNA condensa tion and to selfaggregate at slightly acidic pH [32]. Dps interaction with DNA is to all appearances not sequencespecific, so its genomic binding sites are unknown, and its ability to act as a specific regulator is doubted [33]. Yet the ability of Dps to nonspecifically interact with DNA is very important for nucleoid compactization. In the stationary phase, the nucleoid becomes resistant to detergents and heating. The DNAbound Dps does not dissociate for 16 h at 65°C, and remains bound even when heated to 100°C [29]. Technologi cally, this property may prove very useful. There are several models of Dps–DNA interaction in a compact nucleoid. In one model, Dps in the presence of DNA
forms two superimposed rings of ~9 nm diameter, each comprising six monomers; such rings may further be packed into higherorder structures [29]. This model implies dodecamer reshaping from a sphere into a tor oid, with the monomer hydrophobic surface turning insideout, which appears hardly likely though the DNAbinding ability of Dps dimers does not exclude such a way of interaction. Another model places the DNA into the space between three adjacent dodecam ers [34]. Therewith the Ntails of monomers in one dodecamer may interact with different DNA regions, bringing them closer together or with other dodecam ers, which would apparently hinder ordered conden sation of the nucleoid. In still another model, a central role in Dps–DNA interaction is assigned to salt bridges formed by Mg2+ ions with the negatively charged protein surfaces and DNA [35]. Its drawback is the inability to explain the obvious dependence of the DNAbinding activity of Dps on the structure of its Nterminal segment, which carries a positive rather than a negative charge. The applied significance of bacterioferritin as a basis for ferromagnetic constructs selfassembling on a DNA scaffold substantially depends on the way it interacts with DNA. Yet the very ability of Dps to bind with DNA as a dodecamer and as a dimer implies a possibility of regulatory influence. It is not excluded BIOPHYSICS
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that the microwave effect on dps expression found by us is associated with destabilization of the ferrihydrite core of dodecameric Dps and the ensuing alteration of the oligomeric state of the protein. The dps expression control involves at least five reg ulatory proteins (Fig. 4). HNS (Histonelike Nucle oid Structuring protein) is a major nucleoid protein during exponential growth [16] and a regulator of more than 130 genes. It specifically interacts with DNA, activating or inhibiting the transcription of genes in its regulon. Binding to its site overlapping with the Pdps start site, HNS displaces from the pro moter the RNAP with σ70 but not the enzyme with σ38 [36]. Another nucleoid protein Fis (Factor of inver sion stimulation) also inhibits dps transcription depending on the RNAP subunit composition, but in this case a σ70 enzyme is not displaced but is kept in inactive complexes [36]. Pdps contains two Fis sites, both overlapping with the promoter core, and in the E. coli genome over 200 genes are under positive or negative Fis control [25]. One more dps inhibitor is MntR, which affects a small number of genes by a yet unknown mechanism, depending on Mn2+ concentra tion [26]. Specific activators of dps expression are OxyR and IHF. OxyR is produced in response to oxidative stress in rapidly growing cells; its regulon includes just a score of genes. The dps promoter contains two OxyR sites, one of them in the region of interaction with the RNAP σsubunit, which usually favors transcription complex formation. IHF (Integration Host Factor) is a histonelike nucleoid protein, its cell content reaches a maximum of ~55000 molecules in the early stationary phase [16], and its regulon includes more than 100 genes. Its binding site in Pdps is remote from the region of contact with RNAP, so its activatory action must be mediated either by a structural change in DNA or by other transcription factors. OxyR and IHF can simultaneously bind with the dps regulatory region [27]. Thus, dps expression is controlled by at least three major nucleoid proteins (HNS, Fis, and IHF), which adjust it to the cell growth rate. All of them, as well as the two regulators responding to oxidative stress and Mn2+ homeostasis, inhibit the transcription of their own genes [25]. Such a feedback is typical of most reg ulatory proteins. Therewith, proteins that are repres sors for many genes are usually transcription activators for their own genes, and vice versa. Most probably this principle also applies to dps. Since in the stationary phase the transcription of most genes is inhibited while Dps becomes the main nucleoid protein, we may sup pose that Dps is a negative regulator for most genes and a positive factor for the synthesis of its own mRNA. The vast quantity (180000 molecules) of Dps in a stationary cell can cover the genome at a statistical density of a dimer per 52 bp (or a dodecamer per 310 bp). Clearly, several dimers or at least one BIOPHYSICS
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dodecamer may bind to the dps regulatory region, but the character of such interaction is still totally obscure. The dependence of gene expression on the DNA stretch between primers 1 and 3 found here implies that there is a site for some additional activator. We cannot exclude that the latter is Dps itself. To add, unusual regulation may come from RNAP molecules interacting with the promoterlike sites that do not initiate transcription. In any case, the data obtained suggest that regulation of dps expression is more com plex than it is currently supposed, and that it is influ enced by electromagnetic radiation, which may affect the bacterioferritin structure through altering the state of its ferrihydrite core. Other ferritins accumulating iron ions inside their cavity can also act as magnetore ceptors, but only the DNAcondensing Dps can trans form the signal received into an altered gene expres sion profile, i.e., into an adaptive biological response that is relatively easy to register. This gives premises for using the dps regulatory region in biosensors of back ground electromagnetic radiation, and the bacteriof erritin itself in designing a technology of selfassem bling constructs with a DNA scaffold binding a preset number of Dps dodecamers into a spatially fixed array of ferrite cores. ACKNOWLEDGMENTS The authors thank S.V. Chernyshov (IBC Branch) for kindly donating pET28bEGFP. The work was supported by the Russian Founda tion for Basic Research (100401218a). REFERENCES 1. T. J. Stillman, P. P. Connolly, C. L. Latimer, et al., J. Biol. Chem. 278, 26275 (2003). 2. A. Marchetti, M. S. Parker, L. P. Moccia, et al., Nature 457 (7228), 467 (2009). 3. F. M. Michel, V. Barron, J. Torrent, et al., Proc. Natl. Acad. Sci. USA 107, 2787 (2010). 4. M. C. Garnett and P. Kallinteri, Occup. Med. 56, 307 (2006). 5. J. F. Hainfeld, M. J. O’Connor, F. A. Dilmanian, et al., British J. Radiol. 84, 526 (2011). 6. A. W. Perriman, H. Colfen, R. W. Hughes, et al., Angew. Chem. Int. Ed. Engl. 48, 6242 (2009). 7. A. Jaaskelainen, R. R. Harinen, T. Soukka, et al., Ana lyt. Chem. 80, 583 (2008). 8. S. H. Chu, S. H. Choi, J. W. Kim, et al., United States patent application publication no. US 0134552 A1 (2007). 9. B. D. F. Casse, H. O. Moser, L. K. Jian, et al., J. Phys ics: Conference Series 34, 885 (2006). 10. H. Goronkin, P. Allmen, R. K. Tsui, and T. X. Zhu, in Nanostructure Science and Technology, Ed. by W. Siegel, E. Hu, M. C. Roco (Report of The Interagency Work ing Group on NanoScience, Engineering and Technol ogy and National Science and Technology Council
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