PERSPECTIVE pubs.acs.org/NanoLett
Biomolecule-Based Nanomaterials and Nanostructures Itamar Willner* and Bilha Willner Institute of Chemistry, The Hebrew University of Jerusalem, Jerusalem 91904, Israel ABSTRACT Biomolecule-nanoparticle (or carbon nanotube) hybrid systems provide new materials that combine the unique optical, electronic, or catalytic properties of the nanoelements with the recognition or biocatalytic functions of biomolecules. This article summarizes recent applications of biomolecule-nanoparticle (or carbon nanotubes) hybrid systems for sensing, synthesis of nanostructures, and for the fabrication of nanoscale devices. The use of metallic nanoparticles for the electrical contacting of redox enzymes with electrodes, and as catalytic labels for the development of electrochemical biosensors is discussed. Similarly, biomoleculequantum dot hybrid systems are implemented for optical biosensing, and for monitoring intracellular metabolic processes. Also, the self-assembly of biomolecule-metal nanoparticle hybrids into nanostructures and functional nanodevices is presented. The future perspectives of the field are addressed by discussing future challenges and highlighting different potential applications. KEYWORDS Nanoparticles, quantum dots, sensor, DNA, nanowires, nanostructures, nanodevices
N
anomaterials such as metal nanoparticles (NPs) or nanowires, semiconductor quantum dots (QDs), carbon nanotubes, or inorganic nanowires exhibit unique electronic, optical, and catalytic properties. The dimensional similarities of these nanomaterials and biomolecules, such as enzymes, antibodies (antigens), or DNA, suggest that the integration of biomolecules with these nano-objects may generate hybrid systems that combine the properties of the nanomaterials with the natural recognition and catalytic functions of biomolecules.1,2 These biomolecule-NPs (QDs, nanotubes, etc.) hybrids may then provide a new class of nanomaterials that reveal new tailored functionalities. Also, nanotechnology provides new tools to image and manipulate chemically modified surfaces and to probe chemical interactions at the molecular level. This enables not only the application of these microscopic tools to characterize the structure-function relationships of biomolecule-nanoobjects hybrids, but also to functionalize the nanometric tools with biomolecules and use them for the precise positioning of biomolecules and as biocatalytic manipulators at the nanoscale.3 Biomolecules themselves represent nanoscale materials with encoded structural and functional information. The ability to modify proteins by genetic means or chemical methods, to elicit antibodies or catalytic antibodies, to select and amplify nucleic acid structures with tailored specific binding properties (aptamers) or catalytic functionalities (DNAzymes), paves the way to implement the modified biomolecules or the man-made biomolecular analogs as building blocks to self-assemble and organize
Biomolecules themselves represent nanoscale materials with encoded structural and functional information.
functional nanostructures. Examples include the use of nucleic acid templates to activate enzyme cascades,4 the ingenious self-assembly of nanoscale nucleic acid building blocks into mesoscale structures,5 or the self-organization of supramolecular DNA nanostructures acting as nanomachines.6 Indeed, tremendous progress was accomplished in the past decades in developing biomolecule-based nanomaterials and nanostructures and their application for sensing, nanocircuitry, nanoscale machinery, logic operations, and the fabrication of nanodevices. These advances were summarized in many review articles addressing different facets of nanobioscience.1,2 The present article aims to discuss the broad perspectives of biomolecule-based nanomaterials by introducing the topics with representative examples. Since our laboratory was actively involved in developing the area of nanobioscience, we highlight our accomplishments within the broad scope of the area, while discussing the future perspectives of the field. The electrical contacting of the redox-active sites of enzymes with electrodes is one of the most fundamental issues in bioelectrochemistry and provides the basis for the development of amperometric biosensors and biofuel
* To whom correspondence should be addressed. E-mail:
[email protected]. Tel: +972-2-6585272. Fax: +972-2-6527715. Published on Web: 09/15/2010
© 2010 American Chemical Society
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FIGURE 1. The development of electrical sensor based on metallic NPs or carbon nanotubes. (A) The bioelectrocatalytic activation of redox enzymes by Au NPs. (B) Electrical contacting of reconstituted GOx with an electrode by means of carbon nanotubes. (C) The use of Pt NPs as electrocatalytic labels for the amplified detection of the aptamer-protein (thrombin) complex. (D) Detection of DNA through the generation of a Au-NP-labeled nucleic acid/DNA recognition complex in an insulating gap separating two microelectrodes. The catalytic growth of the Au NPs yields conductivity paths between the microelectrodes and a decrease in the resistivity of the gap region. (E) Time-dependent resistance changes upon analyzing the target DNA (a) and single-base mismatched DNA (b) by the catalytic enlargement of the Au NPs labels. (Part B Reproduced with permission from ref. 14. Copyright Wiley-VCH Verlag GmbH & Co. KGaA. Part C - Reprinted in part with permission from ref. 17. Copyright 2006 American Chemical Society. Parts D and E - From ref. 22. Reprinted with permission from AAAS.)
cells.7,8 Most of the redox enzymes lack, however, direct electron transfer communication between the enzyme redox centers and the electrode due to the spatial separation of the redox sites from the conductive support by the protein shell. Different methods to electrically “wire” redox enzymes with electrodes, by shortening electron transfer distances, were developed, and these included the modification of proteins with electron relays,9 the immobilization of the biocatalysts in redox polymers,10 and the reconstitution of apo-enzymes on relay-cofactor monolayers associated with electrodes.11 The unique electronic properties of metal NPs or CNTs were implemented to electrically wire redox enzymes with electrodes. The conductive nano-objects act in these systems as implanted nanoelectrodes that shorten electron transfer distances and act as relays for transporting electrodes between the redox sites of the enzyme and the electrode. For example, Au NPs (1.2 nm diameter) were functionalized with cofactor units, such as the N6-aminoethyl flavin adenine dinucleotide, NH2-FAD (1), cofactor or the pyrroloquinoline quinone, PQQ (2), cofactor and apo-glucose oxidase, apo-GOx, or apo-glucose dehydrogenase, apoGDH (proteins lacking the native FAD or PQQ cofactors) were reconstituted on the FAD or PQQ cofactor-functionalized Au NPs.12,13 The resulting Au NP-enzyme hybrids © 2010 American Chemical Society
were then linked to Au-surfaces by dithiol bridges, and the integrated enzyme electrodes revealed effective electrical communication with the electrodes, leading to the bioelectrocatalytic activation of the enzyme. The effective electrical wiring of the enzyme (ket ∼ 5000 electron·s-1) was attributed to the structural alignment of the redox cofactors in respect to the Au NPs relays implanted in the proteins, Figure 1A. Similarly, carbon nanotubes, modified at the ends with the amino-FAD cofactor (1), were implanted into apo-GOx via the reconstitution process, and this led to electrically contacted enzyme electrodes, Figure 1B.14 The carbon nanotubes acted in this system as charge carriers from the enzyme redox center to the electrode, resulting in the bioelectrocatalytic activation of the enzyme. Interestingly, the effectiveness of electrical “wiring” of the biocatalyst with the electrode was found to inversely relate to the length of the connecting nanotubes (ket ∝ 1/l where l is the length of the CNTs). While this result seems to be inconsistent with the ballistic conductance of the carbon nanotubes connectors, the phenomenon was attributed to defects introduced into the sidewalls of the CNTs in the course of preparing the CNTs/ enzyme hybrid nanostructures. That is, the prerequisite to cut the CNTs and modify their ends with carboxylic acid residues was essential to construct the hybrid nanostruc3806
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PERSPECTIVE tures. This oxidative process resulted in the formation of defects in the sidewalls of the tubes, and these perturbed the conductance of electrons through the CNTs. As the probability for forming defects relates directly to the length of the tubes, the effectiveness of “wiring” of the biocatalysts relates inversely to the length of the tubes. The implant of metallic NPs into proteins is, also, important for the photonic wiring of proteins with electrodes, and the activation of their photoelectrochemical functions.15 This was demonstrated by the photochemical synthesis of Pt nanoclusters at the photoactive center of the photosynthetic reaction center PS-I, and the assembly of the Pt nanoclusters/PS-I hybrid on electrodes. The trapping of the photoexcited electrons by the Pt nanoclusters was found to facilitate charge separation, and the transport of the trapped electrons to the electrode, thus leading to the effective generation of photocurrents. The use of metal NPs/protein hybrids was further extended to yield electrically contacted three-dimensional electrically contacted matrices on electrodes,16 and these enzyme/NPs composites hold great promise in future sensing applications and the design of biofuel cells or photobiofuel cell systems. Metallic NPs exhibit catalytic properties, and these may be used for amplified biosensing. For example, Pt NPs were found to exhibit horseradish peroxidase-like catalytic activities, such as the electrocatalytic reduction of H2O2 or the catalytic generation of chemiluminescence in the presence of luminol and H2O2.17,18 Accordingly, nucleic acid-functionalized Pt NPs were used as conjugates for the amplified electrochemical or chemiluminescence detection of DNA or aptamer-substrate complexes. For example, the amplified detection of thrombin, that includes two binding sites for its aptamers, was achieved by the use of the aptamer-functionalized Pt NPs conjugate as amplifying reporter, Figure 1C. Binding of thrombin to the aptamer-functionalized electrode was followed by the secondary association of the aptamer-modified Pt NPs that acted as electrocatalytic or chemiluminescent reporter units. The catalytic growth of metallic NPs on metal nanoclusters seeds was also used for optical or electronic biosensing. Different enzymes that generate reducing products, such as oxidases (generating H2O2),19 hydrogenases (generating the NAD(P)H cofactors),20 or proteases (such as alkaline phosphatase that generates cathecol products),20 were assayed by the biocatalytic growth of Au NPs. For example, glucose oxidase19and alcohol dehydrogenase and their substrates20 were optically analyzed by following the plasmon absorbance of the grown NPs. Similarly, tyrosinase was assayed by spectroscopically following the oxidation of tyrosine to L-DOPA and the subsequent growth of Au NPs.21 The catalytic growth of Au NPs also was used for the electrical detection of DNA, Figure 1D.22 A capturing nucleic acid was immobilized on a gap-surface separating two electrodes. The hybridization © 2010 American Chemical Society
of the target analyte DNA and the subsequent binding of nucleic acid-functionalized Au NPs to the resulting hybrid generated a catalytic composite for the growth of the NPs with a silver shell in the presence of hydroquinone. The enlarged particles then generated conductivity paths that bridged the microelectrodes thus decreasing the resistance across the gap, Figure 1E. The resistance was found to decrease upon the increase in the target concentration, accompanied by the increase in the coverage of the catalytic labels. By the thermal control of the hybridization between the capture nucleic acid and the target analyte DNA, single-base mismatch detection in the analyte sequence was demonstrated. Semiconductor nanoparticles (quantum dots, QDs) provide nanoscale materials exhibiting unique optical properties reflected by size-controlled fluorescence features, high luminescence quantum yields, narrow luminescence spectra and large Stokes shifts, and stability against photobleaching. The size-controlled luminescence features of QDs are particularly attractive for sensing, as they enable the use of the variable-sized same materials as different labels for multiplexed analyses. Indeed, tremendous progress was achieved in the past decade in the use of biomolecule-QDs hybrid systems for bioanalytical applications.23 While early studies have implemented antibodyor nucleic acid-functionalized QDs of variable sizes for the multiplexed analysis of pathogens24 or DNAs,25 later studies have used biomolecule-QDs hybrids together with fluorescence resonance energy transfer (FRET) or electron transfer (ET) as photophysical mechanisms to probe biocatalytic transformations as well as to follow recognition events. For example, the ribonucleoprotein telomerase that catalyzes the elongation of the telomer chains is known as a versatile biomarker for cancer cells,26 and it was analyzed by CdSe QDs, Figure 2A.27 The QDs were functionalized with the nucleic acid primer, (3), that is recognized by telomerase. Upon the treatment of the modified QDs with a HeLa cancer cell extract that included telomerase, and in the presence of the nucleotide mixture dNTPs that contained the Texas-Red dUTP, (4), telomerization proceeded on the QDs. The resulting Texas-Red-labeled telomer triggered-on the FRET process from the QDs to the dye units, thus enabling the timedependent analysis of the enzyme activity, Figure 2B. A related FRET process was used to follow the activity of caseine kinase (CK-2), as a representative of kinases that play key roles in signal transduction and the regulation of intracellular processes, Figure 2C.28 The serine-containing peptide sequence, (5), was linked to CdSe QDs, and it acted as substrate for CK-2 that stimulates the phosphorylation of the serine residue to the phosphorylated product, (6). By the association of the Atto-590-labeled antibody, specific for the phosphorylated peptide, the FRET process from the QDs to the Atto-590 acceptor was triggered-on, resulting in the fluorescence of the acceptor dye, provid3807
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FIGURE 2. Biomolecule-quantum dots (QDs) hybrids for sensing. (A) Following the activity of telomerase by the incorporation of Texas-Redfunctionalized dUTP into the biocatalytically grown telomers and using a FRET process. (B) Time-dependent luminescence changes of the QDs upon telomerization. The luminescence of the QDs at λ ) 570 decrease, while the FRET signal of the incorporated dye, λ ) 610 nm is intensified. (C) Optical QDs-based detection of CK2 through the FRET process between the QDs and the Atto-590-labeled antibody that associates to the phosphorylated product. (D) Multiplexed analysis of Hg2+ and Ag+-ions using nucleic acid-functionalized QDs of different sizes. Detection of Hg2+ by 560 nm luminescent CdSe/ZnS QDs and of Ag+ by 620 nm luminescent QDs. (E) Luminescence changes upon the multiplexed analysis of different concentrations of Hg2+ and Ag+ (a) before addition of the ions, (b) 10 µM, (c) 20 µM, (d) 30 µM, and (e) 50 µM of the two ions. (Part B - Adapted with permission from ref. 27. Copyright 2003 American Chemical Society. Part E - Reproduced with permission from ref. 32. Copyright Wiley-VCH Verlag GmbH & Co. KGaA.)
QDs.32 CdSe/ZnS QDs emitting at 560 nm were functionalized with oligo thymine-rich nucleic acid chains, and QDs emitting at 620 nm were modified with oligo-cytosine chains. As the thymine bases form T-Hg2+-T complexes, and the cytosine bases yield specific C-Ag+-C complexes, the formation of the respective Hg2+-thymine nucleic acid structures and the Ag2+-cysteine nucleic acid complexes on the respective QDs resulted in the electrontransfer quenching of the respective QDs, Figure 2D, and the multiplexed analysis of the two ions, Figure 2E. The progress in designing artificial heterocyclic bases or ligand-conjugated bases that bind other ions in duplex DNA nanostructures33 suggests that other ions could be specifically sensed by appropriate, differently sized, nucleic acid-functionalized QDs. Surface-modified QDs hold great promise as optical labels for the monitoring of intracellular processes. For example, Nile-Blue (NB)-functionalized QDs revealed the luminescence quenching of the QDs by a FRET mechanism.34 The reduction of the NB-units by 1,4-dihydronic-
ing a quantitative optical signal for the activity of CK-2. The modification of different-sized QDs with peptide substrates specific for different kinases then could provide an attractive approach for the multiplexed analysis of different kinases. Similar FRET-based assays were used to follow the activities of different proteolytic enzymes.29 QDs were functionalized with dye-labeled peptides specific to different proteases, and the FRET process from the QDs to the dye units proceeded in the peptide-functionalized nanocrystals. The protease-induced cleavage of the peptides removed the dye acceptor units, thus switching off the FRET process, while restoring the luminescence of the QDs. Such FRET processes between QDs and acceptor dyes provide a general and versatile optical readout for sensing platforms that follow the formation of biorecognition complexes, such as DNA hybridization30 or the formation of aptamer-substrate complexes.31 The specific binding of ions to purine and pyrimidine bases of DNA was used to develop ion-specific and multiplexed sensing of ions using nucleic acid-functionalized © 2010 American Chemical Society
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PERSPECTIVE activation of the intracellular processes generated the NAD(P)H cofactors, resulting in the enhanced luminescence from the QDs, Figure 3B. A control experiment that used L-glucose, that is not recognized by the cells, did not alter the luminescence of the QDs, implying that the luminescence changes originated, indeed, from the activation of the cell metabolism. The HeLa cells were then cultured in the presence of Taxol, (7), a drug known to inhibit the metabolic processes occurring in the cytoplasm. Indeed, the Taxol-treated cells revealed only minute increase in the luminescence of the QDs, as compared to the nontreated HeLa cells, consistent with the inhibition of the intracellular metabolism in the drug-treated cells, Figure 3C. Besides the significant progress in using hybrid biomolecule/metal NPs or semiconductor QDs for sensing, these nanobiohybrids find growing interest as building units of nanocircuits and nanodevices. Different biomolecular templates, such as duplex DNA35,36 or protein nanotubes,37 acted as templates for the growth of metallic or semiconductor nanowires. For example, the chemical growth of Ag nanoclusters associated with duplex DNA,35 or the catalytic enlargement of Ag NPs incorporated into β-amyloid nanotubes37 generated metallic nanowires. The base sequence in DNA provides encoded instructive information for programmed self-assembly of nucleic acid nanostructures that reveal dictated functionalities. Guided hybridization, binding of specific low molecular weight or macromolecular substrates (by aptamers), or catalytic functions of nucleic acids represent some of these unique properties of DNA. Indeed, intriguing one-dimensional,38 two-dimensional,39 and three-dimensional40 DNA or DNAprotein nanostructures were designed in the past few years. By the appropriate design of the DNA nanostructures, hybrid metal nanoparticles/DNA nanostructures were synthesized. For example, a two-dimensional DNA array was organized by the self-assembly of 21 single strands of DNA that generate four different double-crossover tiles, Figure 4A.41 The hybridization of Au NPs functionalized with a nucleic acid tether complementary to one of the tiles enabled then the organization of two-dimensional Au NPs arrays with programmed spacing, Figure 4A,B. This methodology was extended to generate tubular nanostructures of DNA carrying ordered configurations of 5 nm Au NPs.42 The folding of a two-dimensional array generated by the assembly of four doublecrossover tiles, where the Au NPs were tethered to one of the tiles, yielded the tubular ordering of the NPs. Interestingly, four types of tubular Au NPs structures were identified, where the symmetrical folding led to stacked rings of the particles, and the nonsymmetrical linkage of the edges of the tiles led to single-spiral, double-spiral, and nested-spiral nanotubes, Figure 4C. A different approach to construct DNA metal nanoparticle hybrid systems has involved the programmed encapsulation of Au NPs in
FIGURE 3. (A) Nile-Blue-functionalized CdSe/ZnS QDs for the analysis of the NAD(P)H. (B) Fluorescence intensity changes upon the activation of the intracellular metabolism in HeLa cancer cells with incorporated Nile-Blue-functionalized QDs upon the addition of (a) D-glucose (b) L-glucose. (C) Effect of taxol on the intracellular metabolism of HeLa cancer cells, monitored by the fluorescence features of the QDs incorporated in the cancer cells, (a) Taxolnontreated cancer cells. (b) Taxol-treated cancer cells. (Reproduced with permission from ref. 34. Copyright Wiley-VCH Verlag GmbH & Co. KGaA.)
otineamide adenine dinucleotide (phosphate), NAD(P)H, yielded the colorless NBH2 units, that prohibited the FRET process, thus triggering-on the luminescence of the particles, Figure 3A. This process was used to sense the NAD(P)H cofactors, and a series of NAD(P)+-dependent enzymes and their substrates,34 but more importantly, the modified QDs were implemented to follow intracellular metabolism. This latter process demonstrated the use of the functionalized QDs as optical labels for the screening of anticancer drugs. The NB-modified QDs were introduced into the cytoplasm of HeLa cancer cells that were grown under starvation, and subjected to the addition of D-glucose that activated the intracellular metabolism. The © 2010 American Chemical Society
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FIGURE 4. Self-assembly of DNA/Au NPs hybrid structures. (A,B) Self-assembly of double-crossover tiles and programmed tethering of metallic NPs on the two-dimensional nanostructures. (C) The folding of four double-crossover tiles where Au NPs are tethered to one tile to yield tubular ordering of nanoparticle consisting of stacked rings of particles, single-spiral, double-spiral, and nested nanotubes of NPs. (D,E) Selfassembly of longitudinal nanotubes of DNA with the programmed ordered encapsulation of Au NPs. (Parts A and B - Reprinted with permission from ref. 41. Copyright 2004 American Chemical Society. Part C - From ref. 42. Reprinted with permission from AAAS. Parts D and E Reprinted by permission from Macmillan Publishers Ltd: Nature Chemistry (Ref. 43), copyright 2010.)
DNA nanotubes exhibiting controlled longitudinal variation.43 Two different sized DNA triangles, functionalized each with three single-strand DNA arms, acted as the building blocks of the DNA nanotubes, Figure 4D. The interlinkage of the differently sized triangles in the presence of Au NPs, by means of three duplex DNA strands with appropriate sticky-ends for hybridization to the respective DNA tethers associated with the triangles, “stapled” the triangles into nanotubes that incorporated the Au NPs, Figure 4E. By varying the sizes of the Au NPs, the NPs could be addressed either to the small or the large capsule domains of the nanotubes. Furthermore, by © 2010 American Chemical Society
the displacement of the “staple” DNA strands by complementary nucleic acids, the nanotubes were separated to the triangle building blocks while releasing the Au NPs. This exemplifies one of the future potential applications of such DNA/metal nanoparticle hybrids in the triggeredrelease of nanocargos from the organized nanostructures. The biocatalytic growth of metallic NPs provided a method to synthesize metallic nanowires.44 Glucose oxidase (GOx) or alkaline phosphatase (AlkPh) were functionalized with Au NPs (1.2 nm diameter) and deposited onto Si surfaces by dip-pen nanolithography (DPN), Figure 5A. The glucose oxidase-mediated oxidation of glucose 3810
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FIGURE 5. (A) The dip-pen nanolithography patterning of Si-surfaces with Au NPs (1.4 nm) functionalized GOx or AlkPh and the biocatalytic growth of Au or the orthogonal biocatalytic growth of Au and Ag nanowires. (B) Biocatalytic growth of Au nanowires by the hybridization of nucleic acid-functionalized glucose oxidase with a DNA scaffold generated by the rolling circle amplification method. (Part A - Reproduced with permission from ref. 45. Copyright Wiley-VCH Verlag GmbH & Co. KGaA. Part B - Reprinted with permission from ref. 46. Copyright 2009 American Chemical Society.)
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FIGURE 6. (A) Synthesis of an actin-Au nanowire-actin nanostructure through the stepwise polymerization g-actin units modified with Au NPs (1.4 nm), and subsequently actin units, followed by the catalytic growth of the NPs to the Au nanowire. (B) AFM image of the actin-Au nanowire-actin structure. (C) Reflectance microscopy images following the ATP-fueled motility of the actin-Au-nanowire-actin structures on a myosin-functionalized glass surface. The time-interval for recording the different frames I to IV is 5 s. (Reprinted by permission from Macmillan Publishers Ltd: Nature Materials (ref. 47), copyright 2004.)
yielded H2O2 that acted as a reducing agent for the enlargement of the Au NPs with gold, resulting in the synthesis of long Au nanowires.45 Similarly, the alkaline phosphatase-driven hydrolysis of p-aminophenol phosphate yielded p-aminophenol that reduced Ag+-ions on the Au catalytic seeds, and this enabled the synthesis of Ag° nanowires on the enzyme template structure. By the sequential deposition of the two enzymes on the surface, the orthogonal synthesis of two different metallic nanowires (Au° and Ag°) was demonstrated. Similarly, a single-stranded DNA template that was synthesized by the rolling circle amplification process (RCA) and consisted of constant repeat units was used as a template for the synthesis of metallic nanowires, Figure 5B. The hybridization of Au NP-modified glucose oxidase, functionalized with nucleic acid tethers complementary to the repeat units of the template, generated an integrated biocatalytic protein/ DNA nanowire structure, and this acted as a template for growing the metallic nanowires.46 The formation of metallic nanowires on biomolecular templates enabled the fabrication of nanodevices and nanomotors. For example, the polymerization of Au NPfunctionalized g-actin units generated the Au NPs-modified f-actin filaments, and these were further extended at their ends by nonmodified actin units, Figure 6A. The catalytic enlargement of the Au NPs yielded Au-nanowires flanked by actin filament tethers, Figure 6B.47 Upon the deposition of the actin-metal nanowire-actin structures on a myosin-functionalized glass surface, the respective actin-myosin motor protein complexes were © 2010 American Chemical Society
generated, and upon the addition of the ATP fuel, the motility of the actin-Au nanowires on the myosin surface was imaged, Figure 6C. The random motility of the nanostructures on the surface at a speed of 25 nm·sec-1 was demonstrated. Such motor nanostructures, consisting of biomolecule-metal nanowire hybrids, hold promise as future nanotransporting elements. (For example, the thermal release of drugs immobilized on the metallic nanowires). The deposition of metal nanoclusters on composite biomolecular templates and the catalytic enlargement of the nanoclusters into conductive domains was implemented to fabricate a nanostructured field-effect transistor device.48 The sequence-specific winding of a homologous nucleic acid into duplex DNA by the RecA protein carrier was used to address the nucleic acid/protein complex in a long duplex DNA scaffold, Figure 7A. The binding of the anti-Rec A antibody to the protein, followed by the association of the biotinylated anti-antibody, shielded the respective DNA scaffold domain. The binding of streptavidin-coated carbon nanotubes to the protein domain, followed by cation exchange and functionalization of the DNA phosphate residues with Ag+-ions assembled the composite nanostructure for the fabrication of the field-effect transistor device. The deposition of the composite nanostructure in the gap separating two microelectrodes, followed by the reduction of the Ag+-ions associated with the nonprotein-coated DNA segments, and their catalytic enlargement with gold, generated the metallic contacts that bridged the single3812
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FIGURE 7. (A) Assembly of a DNA-templated carbon-nanotube-based field-effect transistor device. (B) SEM image of a carbon nanotube bridging the grown metallic contacts of the device. (C) Electrical circuit and electrical characterization corresponding to the nanoscale FET device. The drain-source current is given versus the gate voltage at different drain-source VDS bias potentials: (a) 0.5, (b) 1, (c) 1.4, (d) 2 V. (From ref. 48. Reprinted with permission from AAAS.)
ric sensors, and efficient biofuel cell elements.49 Similarly, the unique size-controlled optical properties of metallic nanoparticles or semiconductor quantum dots found broad applicabilities in bioanalytical science. Different sensing platforms implementing the plasmonic effects of metallic nanoparticles were developed.50 The progress in the synthesis of shaped metallic nanoparticles, such as tripod, tetrapod, or hollow metal NPs, exhibiting new plasmonic features, suggests that extensive theoretical and experimental efforts will be directed to characterize these nanostructures and use them to develop new sensor systems. Also, the substitution of organic dyes by semiconductor QDs as optical labels for biorecognition events finds growing interest. Specifically, the size-controlled luminescence features of QDs facilitate the design of fluorescence resonance energy transfer (FRET) pairs, and their application to follow the dynamics of biorecognition or biocatalytic processes is anticipated to attract growing interest. Also, the size-controlled luminescence properties of QDs are expected to lead to new optical chips for multiplexed analysis. Important future applications of semiconductor QDs are still ahead of us. The incorporation of biomolecule-QDs nanostructures into cells could introduce new possibilities to target specific intracellular domains, thus enabling the imaging of biotransformations with nanoscale precision.
wall carbon nanotubes on the silicon substrate, Figure 7B. The resulting device acted as a field-effect nanotransistor device, where the gold contacts acted as the source and drain electrodes, and the current flow through the device was controlled by the potential applied on the carbon nanotube gate, Figure 7C. Conclusions and Perspectives. The article has discussed recent advances in nanobioscience by addressing different applications of hybrid nanostructures consisting of biomolecules and metallic or semiconductor nonparticles (or carbon nanotubes). The article did not attempt to provide a comprehensive survey of the research area, but aimed to highlight the accomplishments of our laboratory within the general scope of the topic, while addressing the future potential applications of the different functional nanostructures. Indeed, impressive progress was achieved in the application of hybrid biomolecular nanostructures for the development of sensors, nanocircuitry and devices, and the future perspectives of the field are bright. The integration of biomolecules with nanoelements, such as metallic or semiconductor nanoclusters, introduced new directions to the field of bioanalysis and sensor design. The tailoring of reconstituted redox-enzyme/metallic NPs (or carbon nanotubes) hybrids on electrodes not only resolves the fundamental problem of electrical wiring of redox proteins with electrodes, but it provides new opportunities to design miniaturized, implantable, amperomet© 2010 American Chemical Society
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The integration of biomolecules with nanoelements, such as
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metallic or semiconductor (7)
nanoclusters, introduced new direction to the field of bioanalysis
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and sensor design.
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The advances in the self-assembly of programmed biomolecule-NPs nanostructures adds new dimensions to the rapidly developing field of nanobiotechnology. Highthroughput methods to synthesize metal or semiconductor nanocircuitries are emerging, and the use of these hybrid systems to construct nanoscale devices holds great promises. The progress in fabricating nanoscale gaps between electrodes and the development of methods that deposit biomolecule-NPs hybrids in these gaps paves the way to construct new bioelectronic devices. Furthermore, the programmed deposition of stimuli-activated biomolecule/NPs hybrids on patterned nanostructures could lead to nanorobots or nanotransporting devices. Such devices have great potential in future nanomedicine and drug delivery. In fact, recent reports51 indicated that such systems can be constructed, and future activities along these directions should be encouraged. While substantial progress in the assembly of functional biomolecule/NPs was accomplished, numerous scientific challenges are ahead of us. These provide a rich playground for interdisciplinary future research efforts.
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Acknowledgment. The skilled and highly motivated research team members of our laboratory should be acknowledged. Their names appear in the list of references. The continuous support of the Israel Science Foundation is gratefully acknowledged.
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REFERENCES AND NOTES (1) (2) (3)
(4) (5)
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Katz, E.; Willner, I. Angew. Chem., Int. Ed. 2004, 43, 6042–6108. (b) Niemeyer, C. M. Angew. Chem., Int. Ed. 2001, 40, 4128–4158. (c) Gu, H.; Xu, K.; Xu, C.; Xu, B. Chem. Commun. 2006, 941–949. (a) Baron, R.; Willner, B.; Willner, I. Chem. Commun. 2007, 323– 332. (b) Willner, I.; Basnar, B.; Willner, B. FEBS J. 2007, 274, 302– 309. (a) Clausen-Schaumann, H.; Grandobois, M.; Gaub, H. E. Adv. Mater. 1998, 10, 949–952. (b) Hyun, J.; Kim, J.; Craig, S. L.; Chikoti, A. J. Am. Chem. Soc. 2004, 126, 4770–4771. (c) Takeda, S.; Nakamura, C.; Miyamoto, C.; Nakamura, N.; Kageshima, M.; Tokumoto, H.; Niyake, J. Nano Lett. 2003, 3, 1471–1474. Wilner, O. I.; Weizmann, Y.; Gill, R.; Lioubashevski, O.; Freeman, R.; Willner, I. Nat. Nanotechnol. 2009, 4, 249–254. (a) Seeman, N. C. J. Theor. Biol. 1982, 99, 237–247. (b) Mao, C.; Sun, W.; Seeman, N. C. J. Am. Chem. Soc. 1999, 121, 5437–5443. © 2010 American Chemical Society
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(30)
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(c) Winfree, E.; Liu, F.; Wenzler, L. A.; Seeman, N. C. Nature 1998, 394, 539–544. (d) Rothemund, P. W. K. Nature 2006, 440, 297– 302. (e) Simmel, F. C. Angew. Chem., Int. Ed. 2008, 47, 5884– 5887. (f) Niemeyer, C. M. Curr. Opin. Chem. Bio. 2000, 4, 609– 618. (a) Bath, J.; Turberfield, A. J. Nat. Nanotechnol. 2007, 2, 275–284. (b) Simmel, F. C.; Dittmer, W. U. Small 2005, 1, 284–299. (c) Beissenhirtz, M. K.; Willner, I. Org. Biomol. Chem. 2006, 4, 3392– 3401. (a) Willner, I.; Willner, B. Trends Biotechnol. 2001, 19, 222–230. (b) Willner, B.; Katz, E.; Willner, I. Curr. Opin. Biotechnol. 2006, 17, 589–596. (c) Heller, A. J. Phys. Chem. 1992, 96, 3579–3587. (a) Barton, S. C.; Gallaway, J.; Atanassov, P. Chem. Rev. 2004, 104, 4867–4886. (b) Heller, A. Phys. Chem. Chem. Phys. 2004, 6, 209– 216. (a) Schuhmann, W.; Ohara, T. J.; Schmidt, H.-L.; Heller, A. J. Am. Chem. Soc. 1991, 113, 1394–1397. (b) Willner, I.; Lapidot, N.; Riklin, A.; Kasher, R.; Zahavy, E.; Katz, E. J. Am. Chem. Soc. 1994, 116, 1428–1441. (a) Maidan, R.; Heller, A. Anal. Chem. 1992, 64, 2889–2896. (b) Gregg, B. A.; Heller, A. J. Phys. Chem. 1991, 95, 5970–5975. (c) Aoki, A.; Heller, A. J. Phys. Chem. 1993, 97, 11014–11019. (d) Rajagopalan, R.; Aoki, A.; Heller, A. J. Phys. Chem. 1996, 100, 3719–3729. (a) Willner, I.; Heleg-Shabtai, V.; Blonder, R.; Katz, E.; Tao, G.; Bu¨ckmann, A. F.; Heller, A. J. Am. Chem. Soc. 1996, 118, 10321– 10322. (b) Zayats, M.; Katz, E.; Willner, I. J. Am. Chem. Soc. 2002, 124, 14724–14735. Xiao, Y.; Patolsky, F.; Katz, E.; Hainfeld, J. F.; Willner, I. Science 2003, 299, 1877–1881. Zayats, M.; Katz, E.; Baron, R.; Willner, I. J. Am. Chem. Soc. 2005, 127, 12400–12406. Patolsky, F.; Weizmann, Y.; Willner, I. Angew. Chem., Int. Ed. 2004, 43, 2113–2117. Yehezkeli, O.; Wilner, O. I.; Tel-Vered, R.; Roizman-Sade, D.; Nechushtai, R.; Willner, I. J. Phys. Chem. B [Online early access]. (DOI: 10.1021/jp100454u) . Yehezkeli, O.; Yan, Y. -M.; Baravik, I.; Tel-Vered, R.; Willner, I. Chem.sEur. J. 2009, 15, 2674–2679. Polsky, R.; Gill, R.; Kaganovsky, L.; Willner, I. Anal. Chem. 2006, 78, 2268–2271. Gill, R.; Polsky, R.; Willner, I. Small 2006, 2, 1037–1041. Zayats, M.; Baron, R.; Popov, I.; Willner, I. Nano Lett. 2005, 5, 21–25. Xiao, Y.; Pavlov, V.; Levine, S.; Niazov, T.; Markovitch, G.; Willner, I. Angew Chem., Int. Ed. 2004, 43, 4519–4522. Baron, R.; Zayats, M.; Willner, I. Anal. Chem. 2005, 77, 1566– 1571. Park, S.-J.; Taton, T. A.; Mirkin, C. A. Science 2002, 295, 1503– 1506. (a) Medintz, I. L.; Uyeda, H. T.; Goldman, E. R.; Mattoussi, H. Nat. Mater. 2005, 4, 435–446. (b) Costa-Fernandez, J. M.; Pereiro, R.; Sanz-Medel, A. Trends Anal. Chem. 2006, 25, 207–218. (c) Gill, R.; Zayats, M.; Willner, I. Angew. Chem., Int. Ed. 2008, 47, 7602– 7625. Goldman, E. R.; Anderson, G. P.; Tran, P. T.; Mattoussi, H.; Charles, P. T.; Mauro, J. M. Anal. Chem. 2002, 74, 841–847. Gerion, D.; Chen, F. Q.; Kannan, B.; Fu, A. H.; Parak, W. J.; Chen, D. J.; Majumdar, A.; Alivisatos, A. P. Anal. Chem. 2003, 75, 4766– 4772. (a) Shay, J. W.; Bacchetti, S. Eur. J. Cancer 1997, 33, 787–791. (b) Sommerfeld, H. J. Cancer Res. 1996, 56, 218–222. Patolsky, F.; Gill, R.; Weizmann, Y.; Mokari, T.; Banin, U.; Willner, I. J. Am. Chem. Soc. 2006, 125, 13918–13919. Freeman, R.; Finder, T.; Gill, R.; Willner, I. Nano Lett. 2010, 10, 2192–2196. (a) Medintz, I. L.; Clapp, A. R.; Brunel, F. M.; Tiefenbrunn, T.; Uyeda, H. T.; Chang, E. L.; Deschamps, J. R.; Dawson, P. E.; Mattoussi, H. Nat. Mater. 2006, 5, 581–589. (b) Shi, L.; De Paoli, V.; Rosenzweig, N.; Rosenzweig, Z. J. Am. Chem. Soc. 2006, 128, 10378–10379. Gill, R.; Willner, I.; Shweky, I.; Banin, U. J. Phys. Chem. B 2005, 109, 23715–23719. DOI: 10.1021/nl102083j | Nano Lett. 2010, 10, 3805-–3815
PERSPECTIVE (31) Freeman, R.; Li, Y.; Tel-Vered, R.; Sharon, E.; Elbaz, J.; Willner, I. Analyst 2009, 134, 653–656. (32) Freeman, R.; Finder, T.; Willner, I. Angew. Chem., Int. Ed. 2009, 48, 7818–7821. (33) (a) Clever, G. H.; Kaul, C.; Carell, T. Angew. Chem., Int. Ed. 2007, 46, 6226–6236. (b) Tanaka, K.; Clever, G. H.; Takezawa, Y.; Yamada, Y.; Kaul, C.; Shionoyama, M.; Carell, T. Nat. Nanotechnol. 2006, 1, 190–194. (34) Freeman, R.; Gill, R.; Shweky, I.; Kotler, M.; Banin, U.; Willner, I. Angew. Chem., Int. Ed. 2009, 48, 309–313. (35) Braun, E.; Eichen, Y.; Sivan, U.; Ben-Yoseph, G. Nature 1998, 391, 775–778. (36) (a) Gu, Q.; Cheng, C.; Gonela, R.; Suryanarayanan, S.; Anabathula, S.; Dai, K.; Haynie, D. T. Nanotechnology 2006, 17, R14–R25. (b) Richter, J.; Mertig, M.; Pompe, W.; Monch, I.; Schackert, H. K. Appl. Phys. Lett. 2001, 78, 526–538. (c) Mertig, M.; Ciacchi, L. C.; Seidel, R.; Pompe, W.; De Vita, A. Nano Lett. 2002, 2, 841–848. (d) Monson, C. F.; Woolley, A. T. Nano Lett. 2003, 3, 359–363. (37) (a) Reches, M.; Gazit, E. Science 2003, 300, 625–627. (b) Scheibel, T.; Parthasarathy, R.; Sawicki, G.; Lin, X. M.; Jaeger, H.; Lindquist, S. L. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 4527–4532. (38) (a) Cheglakov, Z.; Weizmann, Y.; Braunschweig, A. B.; Wilner, O. I.; Willner, I. Angew. Chem., Int. Ed. 2008, 47, 126–130. (b) Weizmann, Y.; Braunschweig, A. B.; Wilner, O. I.; Cheglakov, Z.; Willner, I. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 5289–5294. (c) Li, H.; Park, S. H.; Reif, J. H.; LaBean, T. H.; Yan, H. J. Am. Chem. Soc. 2004, 126, 418–419. (39) (a) Ding, B.; Sha, R.; Seeman, N. C. J. Am. Soc. 2004, 126, 10230– 10231. (b) Yan, H.; Park, S. H.; Finkelstein, G.; Reif, J. H.; LaBean, T. H. Science 2003, 301, 1882–1884. (c) Cohen, J. D.; Sadowski, J. P.; Dervan, P. B. Angew. Chem., Int. Ed. 2007, 46, 7956–7959. (d) Rinker, S.; Ke, Y.; Liu, Y.; Chhabra, R.; Yan, H. Nat. Nanotechnol. 2008, 3, 418–422.
© 2010 American Chemical Society
(40) (a) Chen, J.; Seeman, N. C. Nature 1991, 350, 631–633. (b) Aldaye, F. A.; Lo, P. K.; Karam, P.; McLaughlin, C. K.; Cosa, G.; Sleiman, H. F. Nat. Nanotechol. 2009, 4, 349–352. (c) Douglas, S. M.; Dietz, H.; Liedl, T.; Hogberg, B.; Graf, F.; Shih, W. M. Nature 2009, 459, 414–418. (d) Andersen, E. S.; Dong, M.; Nielsen, M. M.; Jahn, K.; Subramani, R.; Mamdouh, W.; Golas, M. M.; Sander, B.; Stark, H.; Oliveira, C. L. P.; Pedersen, J. S.; Birkedal, V.; Besenbacher, F.; Gothelf, K. V.; Kjems, J. Nature 2009, 459, 73–76. (41) Le, J. D.; Pinto, Y.; Seeman, N. C.; Musier-Forsyth, K.; Taton, T. A.; Kiehl, R. H. Nano Lett. 2004, 4, 2343–2347. (42) Sharma, J.; Chhabra, R.; Cheng, A.; Brownell, J.; Liu, Y.; Yan, Y. Science 2009, 323, 112–116. (43) Lo, P. K.; Karam, P.; Aldaye, F. A.; McLaughlin., C. K.; Hamblin, G. D.; Cosa, G.; Sleiman, H. F. Nature Chem. 2010, 2, 319–328. (44) Willner, I.; Baron, R.; Willner, B. Adv. Mater. 2006, 18, 1109– 1120. (45) Basnar, B.; Weizmann, Y.; Cheglakov, Z.; Willner, I. Adv. Mater. 2006, 18, 713–718. (46) Wilner, O. I.; Shimron, S.; Weizmann, Y.; Wang, Z. -G.; Willner, I. Nano Lett. 2009, 9, 2040–2043. (47) Patolsky, F.; Weizmann, Y.; Willner, I. Nat. Mater. 2004, 3, 692– 695. (48) Keren, K.; Berman, R. S.; Buchstab, E.; Sivan, U.; Braun, E. Science 2003, 302, 1380–1382. (49) Yan, Y. -M.; Yeheskeli, O.; Willner, I. Chem.sEur. J. 2007, 13, 10168–10175. (50) (a) Haes, A.; Stuart, D. A.; Nie, S.; Van Duyne, R. P. J. Fluoresc. 2004, 14, 355–367. (b) Odom, T. W.; Nehl, C. I. ACS Nano 2008, 2, 612–616. (c) Jain, P. K.; Huang, X.; El-Sayed, I. H.; El-Sayed, M. A. Plasmonics 2007, 2, 107–118. (51) Gu, H.; Chao, J.; Xiao, S. -J.; Seeman, N. C. Nature 2010, 465, 202– 205.
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