Immobilization of Oligonucleotides on Metal-Dielectric Nanostructures

Subscriber access provided by CORNELL UNIVERSITY LIBRARY

Article

Immobilization of oligonucleotides on metaldielectric nanostructures for miRNA detection Alessandro Chiado', Chiara Novara, Andrea Lamberti, Francesco Geobaldo, Fabrizio Giorgis, and Paola Rivolo Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.6b02186 • Publication Date (Web): 07 Sep 2016 Downloaded from http://pubs.acs.org on September 7, 2016

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Analytical Chemistry is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 29

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Analytical Chemistry

Immobilization of oligonucleotides on metaldielectric nanostructures for miRNA detection Alessandro Chiadò*†, Chiara Novara†, Andrea Lamberti†, Francesco Geobaldo†, Fabrizio Giorgis† and Paola Rivolo*† †

Department of Applied Science and Technology, Politecnico di Torino, C.so Duca degli Abruzzi 24

10129, Torino, Italy. * Corresponding authors: Alessandro Chiadò (+390110907370; [email protected]), Paola Rivolo (+390110907383; [email protected]), fax (+390110907399); ABSTRACT: The development of nanostructured metal-dielectric materials, suitable for biodetection

based on Surface Plasmon Resonance and Surface Enhanced Raman Scattering (SERS), requires the refinement of proper biological protocols for their effective exploitation. In this work, the immobilization of DNA probes on nanostructured metal-dielectric/semiconductor substrates has been optimized, in order to develop a bioassay for the detection of miRNA. To ensure a broad relevance, the proposed biological protocol was applied to different silver-decorated functional supports: porous silicon (pSi), TiO2 nanotube arrays and polydimethylsiloxane (PDMS). The efficiency and the stability of the substrates was carefully analyzed by Raman spectroscopy and electron microscopy after the incubation in buffers with the appropriate combination of pH, ionic strength and surfactant content. The customized protocol, initially developed on multi-well plates, was transferred and refined on the nanostructured substrates. The nonspecific interaction of the biological species with the surface was evaluated and reduced thanks to a tailored surface pretreatment. SERS analysis was applied to check

1 ACS Paragon Plus Environment


Page 2 of 29

the immobilization of DNA probes on pretreated samples. Silvered PDMS-supported pSi membranes, 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

the most promising substrates in terms of stability, were subjected to further optimizations. Concentrations, volume and duration of incubations were finely adapted with respect to the surface probe density and to the corresponding hybridization of the complementary miRNA. The optimized ELISA-like assay shows sensitivities comparable to commercial plates for the detection of miRNA222 (LOD: 485 pM), paving the way for the application of the developed protocol on metaldielectric/semiconductor nanostructures for ultrasensitive SERS biosensing applications. KEYWORDS: probe immobilization, ELISA, Surface Enhanced Raman Scattering, miRNA, biosensor, metal-dielectric nanostructures, porous silicon. INTRODUCTION In the last decades, many sensing platforms based on silicon, graphene, metal oxides and elastomers have been studied and devoted to the fabrication of several kinds of biosensors, owing to their attractive ability to combine biological systems with platforms aimed to optical analysis1–3. In order to take advantage of their peculiar physicochemical properties, these materials are more and more nanostructured and/or combined each other4. In particular, noble metals are often combined with semiconductors (e.g. doped Si, InAs/GaAs)5, transition metal-oxides (e.g. ZnO, TiO2, CuO)6,7 and dielectric materials (e.g. glass or polymers) to fabricate metal-dielectric substrates suitable to conduct highly performing analytical techniques, such as Localized Surface Plasmon Resonance (LSPR) and Surface Enhanced Raman Scattering (SERS) spectroscopies. The physical phenomenon common to these techniques is the resonant excitation of surface plasmons in metal nanostructures. In particular for the SERS effect, the huge enhancement and spatial localization of electromagnetic field, provided by the resonant excitation of Localized Surface Plasmons supported on metal nanoparticles, is applied to enhance the Raman scattering of molecules close to their surface. The resonance frequency of the metal nanostructures is strongly dependent on the dielectric environment and can be finely tuned over the entire visible and NIR spectrum, by varying their size, shape, composition and spatial arrangement8,9.


Page 3 of 29


Thanks to the high sensitivity shown by such techniques for a broad number of analytes, metal1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

dielectric substrates became a choice for the analysis of biological systems10 providing efficient signal transducers in optical biosensors8. Recently, the development of new ultra-sensitive metal-dielectric/semiconductor substrates for SERS analysis was reported.4,6,9,11 These substrates, in optimized configurations, can even approach the single molecule detection. The excellent sensing performances of these SERS substrates make them suitable as biosensing platform of challenging biological analytes, such as microRNA (miRNA), a large family of non-coding small RNAs12. These short sequences (20-22 nts) of RNA are able to control gene expression through post-transcriptional regulation via either translational repression or mRNA turnover and are involved in a variety of biological processes including cell proliferation, differentiation, apoptosis, and development13–15. Thus, the aberrant expression of miRNAs has been associated with a number of diseases, genetic disorders, and oncogenesis16. For this reason, they represent one of the most studied classes of biomarkers of the last years16. Nowadays, the large majority of miRNA detection methods involves the use of fluorescent reporters, which require costly chemicals and complex equipment17. For instance, the microarray technologies, the most widely exploited techniques for miRNA profiling, are based on the immobilization of an antisense oligo-probe on flat surfaces and the hybridization of the complementary target sequence is revealed by fluorescence measurement of the labelled miRNAs17–19. Usually oligo-probes are chemically grafted on flat surfaces, such as functionalized glasses or gold thin films, by means of amine or thiol terminations to form self-assembled monolayers (SAM)20–23. Although miRNA detection has been quickly developed, the small size and low abundance of these analytes, whose expression level varies over 4 orders of magnitude, still demands for new specific and sensitive analysis characterized by a large dynamic range16. SERS has been recently regarded as a possible strategy for miRNA profiling of biological samples because it is effective over a large range of



concentration, even in multiplexing analysis, reaching a low limit of detection (LOD) in the label-free 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

configuration18,24. Unfortunately, this option has not been fully investigated, and most of the applications reported in literature are related to the analysis of colloidal systems, to simply physisorbed molecules, or to highly complex systems24–28. Moreover, the application of colloidal systems to biosensing results in poorly reproducible data, mostly because of scarcely controlled aggregation states, and problems concerning with the blinking of the Raman signal due to the nanoparticles Brownian motion29. Instead, it is known that the immobilization of the biorecognition element is critical in biosensing in order to obtain reproducible and accurate results combined to extremely low LOD, especially for a label-free detection. In this work, highly sensitive metal-dielectric/semiconductor substrates used for SERS analysis are investigated for their application to the detection of miRNA. These substrates are composed by different dielectric supports decorated with silver NPs, showing nanostructured surfaces, intense Raman enhancements (ranging from 107 up to 1010 in case of resonant electronic excitation of the analyte) and good uniformity of the SERS signal intensity6,30. Moreover, they can be synthesized on large area, differently from substrates produced by the expensive and time-consuming top-down nanolithography techniques such as electron-beam lithography (EBL) or focus ion beam lithography (FIB). It is worth to emphasize that not only sensitivity, but also stability and reproducibility are key factors for SERS substrates. This is particularly critical for biosensing of complex molecules such as miRNA, whose SERS signal depends significantly on the molecular conformation, orientation and binding specificity to the substrate surface31. Moreover, the hybridization on solid supports is considerably more complex than in solution because of additional interactions that arise from the high probe density and from the support itself32. Finally, the surface probe density is a key parameter for an optimal miRNA hybridization: a low surface coverage yields a low hybridization signal, but an impaired


Page 4 of 29

Page 5 of 29


hybridization can be also obtained with a too high coverage due to steric hindrance33. For these reasons, 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

optimized surface functionalizations and biological protocols designed for the discussed metaldielectric nanostructures are strongly demanded in order to develop biosensing tools for routine miRNA profiling6,34. First of all, a biological protocol for the immobilization of thiol-capped oligo probes and complementary miRNA detection was designed on the basis of more established methods applied to microarray expression analysis35,36 and alkane-thiol SAM formation on metal thin film31,33,37–39. The protocol was then tested on commercial ELISA plates. The miRNA222 was chosen as a model miRNA because of its implication in numerous neoplastic diseases, such as brain, lung, liver, renal, prostatic and pancreatic cancer40–43. The hybridization event was assessed by incubation of a biotinylated miRNA sequence at different concentrations. In order to maximize the hybridization of the miRNA, the effect of ionic strength was evaluated. Subsequently, the biological assay was transferred to the substrates used for SERS analysis, by developing an ELISA-like bioassay on solid samples. The compatibility of the biological protocol with the tested substrates was assessed by incubating the SERS substrates in different buffer solutions. The stability of the substrates was evaluated by means of Field Emission Scanning Electron Microscopy (FESEM), whereas their residual SERS efficiency was measured by comparing the enhancement of the Raman scattering before and after the treatment. Subsequently, the nonspecific signal of the SERS substrates was measured and reduced. Among the investigated substrates, silver-decorated PDMS-supported pSi membranes were subjected to the entire ELISA-like protocol and selected to further optimize the bioassay. In particular, concentrations, volume and duration of incubations, were finely adjusted, with a special attention to the surface probe density, which determines the hybridization efficiency of the complementary miRNA. Thanks to an ELISA calibration curve, the density of molecules, immobilized or hybridized on the surface, was evaluated.



During each experiment, the surface probe density was compared and used to determine the optimal 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

conditions. At the same time, SERS analysis was employed to further confirm probe immobilization. EXPERIMENTAL SECTION Materials and chemicals. DL-dithiothreitol (98.0% DTT), acetic acid (99.0%), acetone (99.5%), ammonium fluoride (98.0%), Bovine Serum Albumin (BSA, IgG free), dimethyl sulfoxide (BioUltra, > 99.5%, DMSO), ethylene glycol (> 99.0%), polyethylene glycol sorbitan monolaurate (Tween-20™), sodium acetate (99.0%), sodium chloride (99.5%), tris(hydroxymethyl)aminomethane (TRIS), sodium phosphate monobasic (98%), sodium phosphate dibasic (98.5%), ethanol (> 99.8%), ethylenediaminetetraacetic acid (EDTA, 99.4%), Rhodamine 6G (99.0%, R6G), saline sodium citrate (SSC, sterile stock 20x, 300 mM trisodium citrate, 3 M NaCl), sodium dodecyl sulfate solution (BioUltra, 10%, SDS), 3,3',5,5'-tetramethylbenzidine (TMB), and all the DNA and RNA oligos were from Sigma Aldrich, Milan, IT. HF was from Carlo Erba, Milan, IT. PDMS pre-polymer and curing agent (Sylgard 184) were from Dow Corning. Ti foil (70 µm thick, 99.6%) and Pt foil (250 µm thick, 99.9%) were from Goodfellow. Kapton® tape was from RS (Milan, IT). Horseradish Peroxidase conjugated Streptavidin (Str-HRP) was purchased from Jackson ImmunoResearch Europe Ltd. (Suffolk, UK). Illustra MicroSpin G-25 columns were from GE Healthcare (Fisher Scientific, Illkirk, FR). Water used during each step was Milli-Q™ dispensed from a DirectQ-3UV (Merck-Millipore, Milan, IT) and sterilized. Fabrication of SERS substrates. In this work, three pSi substrates were used: immersion-plated (IP), ink-jet printed (printed pSi or PPS) and PDMS-supported pSi (PSD). The pSi was prepared by anodization of highly boron-doped silicon wafers (34-40 mΩ-cm resistivity) at room temperature, in HF solution (20:20:60 HF/H2O/CH3CH2OH) with a current density of 125 mA/cm2 for 30 s. Further details are reported elsewhere11. Silver NPs were synthesized in situ from silver nitrate, either by inkjet printing or by immersion plating. Concerning the first method, a piezoelectric Jetlab 4-XL printer from


Page 6 of 29

Page 7 of 29


MicroFab Technologies Inc. (Plano, TX, USA) equipped with a 60-µm nozzle diameter MJ-AT-01 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

dispenser was used to print 2.5 10-2 M AgNO3 water/DMSO/ethanol solutions (68/22/10 v/v%) on the freshly etched mesoporous silicon. Six printhead passes and 1000 dpi spatial resolution were selected as optimized printing parameters. IP substrates were silver-decorated through immersion plating, by dipping the porous samples in an aqueous 10-2 M AgNO3 solution, at 50 °C. PSD membranes were produced by electrochemical detachment of pSi from the original Si substrate, anodizing in a low concentrated ethanolic HF electrolyte (4%), at a current density of 4 mA/cm2 for 95 s. Meanwhile, a partially cross-linked PDMS substrate was obtained by casting PDMS oligomer and curing agent mixed at a 10:1 weight ratio, degassing for 20 minutes and curing in an oven at 80 °C for 10 minutes. The PDMS substrate was placed in contact with the pSi membrane, which was immobilized on the polymer after Si stripping. Silver NPs were synthesized by dipping the pSi/PDMS membranes in an aqueous 10-2 M AgNO3 solution supplemented with ethanol 2.5% v/v and HF 0.0006% v/v for 2 minutes at 50 °C. PDMS-Ag was prepared starting from PDMS slices obtained by mixing pre-polymer and curing agent (Sylgard 184, Dow Corning) with a 20:1 weight ratio, degassing for 1 hour, casting the mixture into a PMMA mold (fabricated by milling machine) and finally curing in a convection oven for 1 h at 70 °C. Ag NPs were deposited on the PDMS membranes by DC sputtering in Ar atmosphere at 10−4 bar (Q150T-ES, Quorun Technologies) using a sputtering current of 40 mA for 5 s; the PDMS membranes were at 5 cm from the Ag target. TiO2 nanotube arrays (TiO2-NTs) were grown by anodic oxidation of a Ti foil in an electrolytic solution consisting of 0.5 w/v% NH4F and 2.5 v/v% Milli-Q™ water in ethylene glycol. Ti foils were cleaned by ultrasonication in acetone, rinsed in ethanol and etched for 1 minute in 1% HF aqueous solution. The backside of the Ti foil was protected by Kapton® tape and used as anode while a Pt foil was employed as a cathode in a two-electrode configuration Teflon cell. The electrochemical process



was conducted applying an anodization potential of 60 V using a DC power supply (GW Instek SPD1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

3606) for 15 minutes. After the anodization, the oxidized Ti foils were abundantly rinsed in Milli-Q™ to remove electrolytic solution trapped in the NTs during the growth. More details on NTs growth and characterizations can be found elsewhere44. The NT arrays were subsequently annealed at 450 °C in air to crystallize them into anatase phase. Ag NPs were deposited at room temperature by DC sputtering in an Ar atmosphere, as reported above, but at 40 mA for 30 s. This process was optimized in order to maximize the SERS phenomenon, as previously reported6. Probe reduction and SERS substrates pretreatment. A 5’-alkylthiol-capped DNA probe (probe222, 5’-C6SH-ACCCAGTAGCCAGATGTAGCT-3’), corresponding to the antisense sequence of the miRNA222, was used to detect the complementary RNA sequence. The probe was reduced with DTT, purified with Illustra MicroSpin G-25 columns and quantified by means of a Cary5000 UV-VisNIR spectrophotometer (Agilent Technologies Italia S.p.A., Milan, Italy) equipped with a TrayCell (Hellma GmbH & Co., Müllheim, Germany). Then, the freshly reduced probe was diluted to the working concentration (10-25 µM) in the proper buffer. All SERS substrates were pretreated by one-hour incubation in 1% BSA in 50 mM sodium acetate buffer pH 4.0 (NaAc) to reduce the nonspecific binding. Finally, they were washed in TE buffer (10 mM Tris, 1 mM EDTA, pH 7.5) to remove the protein excess. Bioassay development on ELISA plates. An ELISA protocol was developed for optimization purposes. With this aim, a 3’-biotinylated probe222 (5’-C6SH-ACCCAGTAGCCAGATGTAGCTbiotin-3’) was used to check the proper immobilization of the probe. At first, the whole protocol was tested in maleimide-functionalized 96-well plates (Sulfhydryl-BIND™, Corning) and then scaled-down to SERS substrates. The wells were washed with TE and then 30 µL/well of freshly reduced probe222 in TE-NaCl (TE, 1M NaCl, pH 7.5) were incubated overnight at room temperature; then, the wells were washed trice with TE-t (TE, 0.05 % Tween20™, pH 7.5) and blocked with 1 % BSA in TE.


Page 8 of 29

Page 9 of 29


Afterwards, the plate was incubated with 50 µL of 0.5 µg/mL Str-HRP in TE-t, washed trice with TE-t 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

and developed. Briefly, the TMB substrate solution was added onto the samples to initiate the colorimetric reaction, which was stopped after 2 minutes adding H2SO4 (0.5 M), with a 1:1 TMB:H2SO4 ratio. The Optical Density (OD) of the solution was immediately measured at 450 nm and at 630 nm by means of a 2100-C microplate reader (Ivymen Optic System). In order to indirectly quantify the amount of probe immobilized on the surface, an in-liquid titration curve with Str-HRP, serial diluted in Milli-Q™, was prepared and the related OD was measured. The OD recorded for the different wells was normalized and transformed in surface density (molecules/cm2) by comparison with the in-liquid titration curve, as previously reported45,46. Since the measured signal corresponds to the molecules of Str-HRP, the probe density was estimated by multiply that value by a factor of 50, corresponding to the Str/probe surface ratio. The surface occupied by one molecule of probe and one of Str-HRP was calculated by means of Marvin Sketch (6.1.7, 2014, ChemAxon http://www.chemaxon.com) and UCSF Chimera47, by applying the methods of Palmara et al.46. Several replicates for each experimental condition were analyzed and the error bars reported on the histogram diagrams concerning with colorimetric ELISA-like measurements represent the standard deviations (SD). To find the optimal conditions for the miRNA hybridization, a non-biotinylated probe222 was used to detect a 5’-biotinylated miRNA222 (5’-biotin-AGCUACAUCUGGCUACUGGGU-3’). The same protocol reported above was used, unless that 50 µL of biotinylated miRNA222 in SSC 4x (from stock 20x, 0.1% SDS, pH 7.5) was incubated for 1 hour, after the blocking step. Then, the plate was washed trice in SSC 1x (diluted SSC 4x), incubated with 50 µL of 0.5 µg/mL Str-HRP in SSC 1x, washed trice with SSC 1x and developed as reported above. The LOD was calculated considering the slope of the linear portion of the calibration curve in the range 0–1 nM (y = 1.10×1011 x + 3.21×109, r2 = 0.984). Such a linear fit is extrapolated towards the mean of the blank samples plus three times its standard deviation, as reported in the literature40.



SERS substrates compatibility to bioassay. All SERS substrates were tested for compatibility to 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

the bioassay by overnight incubation in the buffers used during the whole procedure. The buffers used were NaAc, TE, TE-t, TE-NaCl, TE-NaCl-t (TE-NaCl, 0.05 % Tween20™, pH 7.5), SSC 4x and SSC 1x. The samples (two replicates for each buffer) were previously tested for their SERS efficiency, by measuring the minimum detectable concentration of the R6G (see below); 0.2 µL/mm2 of R6G ethanolic solution was spotted onto the substrates, starting from 10-14 M concentration and let dry before the Raman measurements. After the overnight incubation, the samples were washed in MilliQ™ water, dried under a stream of nitrogen and tested again for SERS efficiency by means of R6G. Afterwards, all the samples were analyzed by FESEM to check any morphological change. FESEM images were acquired as secondary electron contrast images with 5 keV electrons using an in-lens detector of a Zeiss SUPRA 40 (Zeiss SMT, Oberkochen, Germany) microscope. PSD samples were covered with a copper grid anchored to the FESEM stub to dissipate charge loading. Afterwards, the nonspecific signal of the Str-HRP, diluted in the above-reported buffers and incubated on pretreated and not pretreated samples was carefully evaluated by TMB colorimetric reaction (see above). ELISA-like bioassay on metal-dielectric substrates. The ELISA protocol developed in 96-well plates was transferred and refined on the metal-dielectric substrates. During these tests, the 3’biotinylated probe222 was used to check its proper immobilization on silver NPs, whereas, the 5’biotinylated miRNA222 was used to find the optimal conditions for the miRNA hybridization. All the SERS substrates were cut in 5x5 mm2 pieces and pretreated as reported above. The immobilization of the probe222 on the silver NPs was obtained by overnight incubation at room temperature of 5 µL of freshly reduced probe222 diluted in TE-NaCl, unless differently stated. Negative controls were as well incubated overnight in TE-NaCl without the probe (no-probe control). After overnight incubation, the samples were washed thrice in TE-t to remove nonspecific binding, and blocked with 1 % BSA in TE. For the washing and blocking steps the samples were put in 48-well plates and a working volume of 300 µL was applied. Afterwards, the same protocol reported for 96-well plates was applied to SERS


Page 10 of 29

Page 11 of 29


substrates (see above). As well, the LOD was calculated as reported above by the calibration curve in 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

the range 0–5 nM (y = 4.29×109 x + 1.50×109, r2 = 0.998). 2.7 SERS analyses on metal-dielectric substrates. The same protocol finely tuned to immobilize the probe222 on the SERS substrates was used for Raman analysis, unless that after the overnight incubation, the samples were washed once in TE-t and once with Milli-Q™. Thus, the substrates were not incubated with miRNA or Str-HRP, but dried under a nitrogen stream and analyzed with a Renishaw InVia Reflex micro-Raman spectrometer (Renishaw plc, Wotton-under-Edge, UK) with a 514.5 nm laser excitation in backscattering light collection under a 50x objective. The same measurement conditions were employed in the SERS efficiency tests with R6G. RESULT AND DISCUSSION Bioassay development in ELISA plates. Starting from various protocols related to microarray expression analysis selected from the literature, an ELISA bioassay was developed in 96-well plates for optimization purposes. Commercial maleimide-activated plates were used because they ensure the covalent binding of protein and oligonucleotide through amine and thiol groups. Owing to the presence of silver or other noble metals on SERS substrates, thiol-capped DNA probes (probe 222) were used. To covalently bind these oligos to the surface, they were reduced with DTT to ensure the presence of free –SH. Then, buffers commonly used with DNA and based on Tris and EDTA (TE, TE-t, TE-NaCl, TE-NaCl-t) were tested for their binding capabilities. As reported in the literature, in order to maximize the amount of well-packed probes on a surface, a certain amount of salt is required to shield the negative charges carried by the phosphate groups of the backbone39,48. At the same time, surfactants (e.g. Tween-20™, Triton™ X-100) are added to reduce the nonspecific binding due to the hydrophobic interaction with materials such as pSi or PDMS. The effect of these buffers on the surface density of immobilized probe222 and the corresponding miRNA222 hybridization has been carefully analyzed. This parameter is of paramount importance for an optimal miRNA hybridization: the probe density



Page 12 of 29

should be in a range that would give rise to the maximum detectable signal, but avoiding steric 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

hindrance due to tightly packed probe molecules33. The results show that only the TE-NaCl buffer is able to promote an effective immobilization of the probe (Figure S1A), which allows observing the hybridization of the complementary miRNA sequence (Figure S1B). In the other buffers (e.g. in TE) the probe molecules randomly adsorb on the surface or the surfactant masks any possible interaction. For this reason, the signal recorded during the following hybridization of miRNA is comparable to the experimental background. In order to better understand the effect of salt concentration, the probe222 was incubated in TE supplemented with increasing concentration of NaCl and the surface probe density and the corresponding miRNA222 hybridization were assessed (Figure S2A). The plot highlights that the surface probe density increases along with the increasing amount of salt, up to 1 M NaCl. Once this salt concentration is reached, the hybridization of miRNA does not change anymore, probably because of the surface saturation, or due to other effects concerning the stability of the densely packed grafted probes and DNA-RNA hybrids at relatively high salt concentration. Furthermore, the hybridization efficiency was determined during these experiments by calculating the ratio of the miRNA molecules annealed to the probes on the surface. Even if this efficiency is higher at 250 mM NaCl, the optimal salt concentration is confirmed to be 1M NaCl, which ensures to double the signal of miRNA hybridization. Finally, the ELISA measurements performed in 96-well plates, concerning the labelled miRNA incubated at different concentrations (Figure S2B), show that the developed bioassay is suitable for the detection between 0.2 and 50 nM, corresponding to a physiological range of concentrations49. Moreover, a LOD of 177 pM was calculated by taking into account the signal obtained from blank samples (see Experimental section).


Page 13 of 29


SERS substrates compatibility to bioassay. The bioassay protocol developed in maleimide1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

activated plates was transferred and refined on metal-dielectric/semiconductor substrates in an ELISAlike assay. Three solid supports were tested: a porous silicon matrix (pSi), an elastomeric material (PDMS) and a nanostructured semiconductor (TiO2-NTs). Each dielectric material decorated by silver NPs is able to confer new interesting properties to the SERS substrate. In detail, the substrates can show a high surface area and high refractive index for pSi2, a stretchability and therefore a tunability of the plasmonic properties for PDMS6, an additional chemical enhancement due to the semiconducting nature of TiO2-NTs4,6. Given that each surface is chemically and physically different from the others in terms of nanostructuration, chemical reactivity and wettability, the immobilization and the sensing protocols must be refined during the scale-down of the assay from 96-well plates to SERS substrates. In particular, the functionalized plates are flat and the immobilization of the probe occurs through the covalent bonding of the thiol groups to maleimide groups, whereas on the substrates the probe is grafted through a semi-covalent bond due to the interaction between the thiol groups and surface silver atoms of the NPs. Moreover, since these substrates are intended for biosensing applications, a crucial step to be investigated is their stability in aqueous buffer solutions. In order to exclude any morphological change (e.g. size and distribution of Ag NPs), the surfaces were analyzed by means of FESEM (Figures 1 and S3). Furthermore, the SERS efficiency of the substrates was investigated by evaluating their ability to enhance the Raman signal by SERS effect, using R6G spotted on their surface before and after the treatment in the buffers used for the biological sensing protocol (Figure 2).



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 14 of 29

Figure 1. FESEM images of SERS substrates before and after incubation in TE-NaCl buffer: A) inkjet printed Ag-pSi (PPS); B) silvered PDMS-supported pSi (PSD); C) immersion plated Ag-pSi (IP); D) silvered TiO2-NTs.

Figure 2. A) Minimum detectable R6G concentration on all the investigated SERS substrates before and after incubation in TE-NaCl; B) SERS spectra of R6G on PSD substrates at several concentrations. *The minimum detectable concentration is missing due to the marked heterogeneity of the distribution of Ag NPs after the buffer incubation as shown in Figure 1. The FESEM analyses of the inkjet printed Ag-pSi (PPS, Figure 1A, S3A) and silvered PDMSsupported pSi (PSD, Figure 1B, S3B) samples revealed that the morphology of these substrates remains


Page 15 of 29


almost unchanged after the incubation into the buffers. Tests on immersion plated Ag-pSi (IP, Figure 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1C) were also carried out, demonstrating that the main stability issues are related to the damage of the porous layer. This layer suffers from mechanical strains produced during the electrochemical etching and the subsequent NPs synthesis, thus the silver coating exfoliates, due to the extended dense particle coverage and surface oxidation. Porous silicon is also damaged (e.g. larger pores) by the buffers containing Tris (Figure S4), as already mentioned in the literature51. This could be observed also for pSi of PPS substrates, even if this did not compromise the NPs. Some surface cracks can be observed only outside of the printed region (Figure S5) or if the NPs are too much densely packed probably because of the higher applied surface strains51,52. All of these effects can be reduced by treating the substrates with DMSO/ethanol mixtures that quickly oxidize the pSi in the most strained regions53. On the other hand, PSD membranes did not display morphological changes, probably because of the reduced strains after transfer on the PDMS substrate, which provides also a higher mechanical stability. The images of NTs substrates after the incubation (Figure 1D, S3C) showed, instead, that the silver NPs formed small clusters and dispersed less homogenously on the surface in comparison to the untreated samples. This behavior is probably due to the different surface properties of the last kind of substrates and to the synthesis of the NPs. Our hypothesis is that the sputtered silver NPs interact very weakly with the surface of the TiO2-NTs, in comparison to those directly produced by a chemical reaction on pSi. Finally, a careful FESEM analysis of treated and untreated PDMS-Ag substrates was not possible because of strong charge-up effects. Raman spectroscopy performed on R6G adsorbed on the metal-dielectric substrates in aqueous solutions shows experimental data in agreement with the above discussed morphological analyses. Figure 2 reports the minimum detectable concentrations before and after the buffer treatment for the four SERS substrates (data are shown for the TE-NaCl buffer). It can be noticed that the SERS efficiency is preserved for printed and PDMS-supported pSi. On the other side, IP samples showed a noticeable change of the silvered surface and it was not possible to evaluate this parameter on the



Page 16 of 29

treated specimens because of the strong spatial heterogeneity of the Ag NPs reflected in the fluctuation 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

of the SERS signal intensities. Finally, three orders of magnitude in the minimum detectable R6G concentration are lost for the sputtered TiO2-NTs and PDMS after the buffer incubation due to the partial detachment of very weakly interacting silver NPs in polar, salt-containing environments. Nevertheless, despite the decreased SERS enhancement, PDMS-Ag and TiO2-NTs still show good sensitivities, displaying detectable concentrations of 10-9 M. At the same time, results of Figure 2 show that the highest enhancement of the Raman signal was obtained with the PSD samples, which allowed a R6G detection at a concentration as low as 10-14 M, one of the lowest values reported in the literature (Figure 2B). Bioassay optimization: nonspecific binding reduction. Once the buffers compatibility was attained, the nonspecific binding of the Str-HRP (essential for the colorimetric ELISA-like development) on the SERS substrates was evaluated and reduced to reach a good signal to noise ratio (S/N). The high reactivity towards biological species of some inorganic materials such as porous silicon can be useful for biosensing application (e.g. easier immobilization), but also detrimental in terms of S/N and often additional pretreatment steps are required to reduce nonspecific binding1,54. On the other hand, the efficiency of probe immobilization on the silver NPs should be preserved while keeping low the background signal in the following Raman measurements. BSA is typically applied as blocking agent to prevent the nonspecific binding in immunoassays55 and it seems to meets the listed requirements. The effect of a pretreatment step with 1% BSA and the subsequent incubation in the buffers selected in 96-well plates was then evaluated during the same experiment. It is well known that the adsorption of BSA on silica is strongly dependent on pH1,56,57. In order to maximize the adsorption of the BSA molecules on the pSi surface, the pretreatment was performed in NaAc at pH 4.0, a pH value just below the isoelectric point of the protein. The same approach was applied to the TiO2-NTs samples. The Str-HRP was diluted in different buffers and incubated for one hour on BSA-treated or untreated substrates. Afterwards, the substrates were developed as reported above (see Experimental


Page 17 of 29


section). A signal significantly similar to the background was considered not affected by nonspecific 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

adsorption of Str-HRP. Figure 3 shows the results of this test for PPS, TiO2-NTs, and PSD samples.

Figure 3. Evaluation of the nonspecific binding of the Str-HRP incubated in different buffers (a: H2O; b: TE; c: TE-t; d: TE-NaCl; e: TE-NaCl-t; f: SSC4x; g: SSC1x;) on the SERS substrates previously pretreated or not in BSA in NaAc. The signal obtained was transformed in number of molecules/cm2 (see Experimental section). The plots in Figure 3 clearly demonstrate that the pretreatment step is necessary to reduce the nonspecific adsorption of the Str-HRP on the analyzed SERS substrates. The untreated substrates, especially the silvered TiO2-NTs, showed an extremely variable nonspecific signal, whereas, all the BSA-blocked samples, except for those incubated in water, showed a reduced signal if compared to the freshly synthetized samples. It is feasible that the BSA molecules physisorb on the surface of the substrates (pSi, TiO2, PDMS and Ag), as observed by the higher wettability of the pretreated samples in comparison to the untreated ones, preventing the nonspecific binding of Str-HRP. Moreover, the adsorbed BSA molecules seem to be stably attached to the surface, as they cannot be removed by means of the subsequent rinsing steps, even if surfactants are present or pH is changed. These findings confirm that the behavior already observed on bare pSi1 is also obtained on TiO2-NTs and the other



Page 18 of 29

nanostructured materials. At the same time, the presence of surfactants, such as Tween-20™ or SDS, 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

further reduced the adsorption of Str-HRP on the surface. This can be noticed comparing the signal obtained by incubating the Str-HRP in buffers containing surfactants with the one obtained by incubating the samples in water (or TE). This effect is particularly evident on untreated TiO2-NTs: SDS only is able to reduce the Str-HRP adsorption on these kind of samples. Although these experiments confirm that the bioassay optimized on 96-well plates can be applied to pretreated SERS substrates, the binding of the BSA on silver NPs could be detrimental for the immobilization of the probe. In fact, the BSA can adsorb on the surface of the NPs through hydrophobic effect and through its cysteine residues. In order to check if the BSA pretreatment prevents the probe immobilization, the probe222 was incubated overnight on the SERS substrates with or without BSA-blocking and its binding was assessed by means of SERS analysis. In Figure 4A the Raman spectra of the probe grafted on PSD substrates are reported. This plot highlights that it is still possible to graft the probe on BSA-pretreated PSD samples. Indeed, it can be noticed that the two spectra show similar features, except for a few contributions around 1000 cm-1 and 1660 cm-1 which are only observed on the pretreated sample. These bands can be assigned to the aromatic aminoacids ring breathing and to the amide I mode of BSA.58 Typical vibrational peaks of the nucleobases can be instead detected on both samples at 730 cm-1 (adenine breathing), 790 cm-1 (cytosine and thymine breathing), 1270 cm-1, 1325 cm-1 (C-N stretching of cytosine and adenine ring respectively), 1480 cm-1 (guanine C-N stretching and N-H bending)59, and 1570 cm-1 (adenine and guanine amino groups scissoring).60 The evaluation of the probe222 binding in the presence of different buffers was also verified by ELISA assay on the surface of pretreated PSD samples, as reported in Figure 4B. This plot clearly shows that, even if the samples are pretreated, the probe incubated in TE-NaCl is grafted on the surface


Page 19 of 29


with an efficiency higher than the one obtained with the other buffers. These results remark the 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

behavior obtained on 96-well plates, thus strengthening the whole procedure.

Figure 4. A) Raman analysis of the probe222 grafted on the surface of PSD samples treated or not with BSA. Stars mark the BSA peaks. B) Evaluation of the surface density of probe222 incubated on PSD samples in different buffers. Scale-down of the bioassay to PSD SERS substrates: optimization of volumes, concentration and incubation times. The bioassay protocol developed in maleimide-activated plates was then transferred to the SERS-substrates, optimized and applied for miRNA detection. In order to scale-down the protocol from 96-well plates to the substrates, the amount of incubated probe per area (~2 µg/cm2) was kept at a fixed ratio. Taking into account the area of the well, incubated with a certain volume, and the amount of probe in this volume, the corresponding volume and concentration was calculated for the samples. The application and optimization of the protocol to the PSD samples is reported, as example. This kind of substrate is the newest one for what concerns the synthesis procedure and it presented the lowest variability among all the tested samples. Moreover, the PSD membranes are the most promising substrates for SERS analysis, due to the negligible background signal, the low intra-substrate signal fluctuation, the high SERS efficiency and intensity uniformity (SERS signal intensity RSD ~ 12% were calculated by Raman mapping; see above and Figure S6).



Page 20 of 29

At first, the concentration of incubated probe222 (10 and 25 µM at a fixed volume of 5 µL) and the 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

volume of the drop-incubations (10-25-50 µL) during the following steps (miRNA and Str-HRP incubations) were investigated, as reported in Figure 5A. The plot represents the signal of the biotinylated miRNA222 in the different tested conditions. This test demonstrates that the optimal volume for the drop-incubations of miRNA222 and Str-HRP during the bioassay is 25 µL. A volume of 50 µL results in a higher variability, whereas a lower volume (10 µL) limits the detection of miRNA. At the same time, by considering a volume of 10 and 25 µL, in both cases the detection of miRNA is a little reduced (~6%) when the concentration of probe222 incubated on the samples is 10 µM, thus indicating that the surface saturation is approaching at 25 µM of probe. This result was confirmed by the incubation of biotinylated probe222 solutions at 10 and 25 µM concentration, respectively. The obtained surface probe densities were 1.30 ± 0.12 × 1012 and 1.44 ± 0.07 × 1012 molecules/cm2, respectively, and then correspondingly to the estimated hybridization efficiencies of 85.4% and 88.2%. Therefore, the concentration of probe and the volume of drop-incubations were fixed to 25 µM and 25 µL for the following experiments.

Figure 5. Optimization of the ELISA-like bioassay on PSD samples: A) evaluation of the concentration of incubated probe222 and volumes of incubation of miRNA222 and Str-HRP; B) effect of the duration of incubation of miRNA222 and Str-HRP.


Page 21 of 29


The duration of incubation of miRNA222 (10, 20, 40 and 60 minutes) and Str-HRP (30 and 60 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

minutes) was investigated, as reported in Figure 5B. The plot shows a monotonic trend of the detected signal vs. the incubation time, with the highest signal obtained by incubating the miRNA222 for 60 minutes on the functionalized PSD samples. This behavior is attained with both the tested times of incubation of the Str-HRP; on the other hand, an incubation time of 60 minutes yields a signal intensity four times higher than the one obtained for 30 minutes of incubation. The final protocol was then applied to the detection of different concentrations of miRNA222, as reported in Figure 6. This test clearly demonstrated that the completely optimized protocol allows to specifically detect this biomarker in a physiological range of concentrations (0.5-50 nM), with a LOD of 485 pM. This LOD is probably the result of a very low background and variability of the blank samples. Finally, in comparison with the same test performed on commercial 96-well plates (Figure S2B), the ELISA-like performed on the properly functionalized PSD substrates revealed a similar dynamic range and a comparable LOD.

Figure 6. Detection of different concentration of miRNA by means of functionalized PSD samples.

In the light of the discussed results, the optimized biological protocol is suitable and ready to be applied for SERS-based biosensing platforms, further boosting the sensitivity checked in the discussed



Page 22 of 29

ELISA-like measurements. In particular, these findings open the way for the application of 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

nanostructured metal-dielectric/semiconductor substrates to ultrasensitive miRNA profiling. CONCLUSION In this work, a protocol optimized for the biosensing application of ultra-sensitive metal-dielectric nanostructures has been developed. This protocol was elaborated on commercial ELISA plates and then transferred and refined on different substrates made of silver-decorated pSi, TiO2-NTs and PDMS. The efficiency of such substrates and their stability were assessed step by step by FESEM and SERS analyses. In particular, an essential pretreatment was applied to reduce nonspecific signal due to the adsorption of biomolecules on the supporting surfaces. SERS analysis was exploited to confirm probe immobilization on pretreated substrates. Afterwards, silvered PDMS-supported pSi membranes were subjected to the whole protocol and selected to optimize the bioassay. Crucial parameters such as concentrations/volume/duration concerning with the incubation steps were carefully refined, taking into account the surface probe density and the corresponding hybridization of the complementary miRNA. During the experiments, the surface probe density, obtained by ELISA calibration, was compared and used to determine the optimal conditions. In such a way, a model sequence (miRNA-222) was detected in a physiological range of concentrations (0.5-50 nM), showing a LOD of 485 pM. This study opens the way for label-free ultrasensitive SERS-based miRNA profiling. ASSOCIATED CONTENT Supporting Information. The optimization of the bioassay in 96-well plates, the FESEM images related to the stability of SERS substrates and a representative analysis of the uniformity of the SERS signal on the substrates are reported in the supporting information. This material is available free of charge via the Internet at http://pubs.acs.org. AUTHOR INFORMATION


Page 23 of 29


Corresponding Authors 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

*E-mail: [email protected], [email protected] Notes The authors declare no competing financial interest. ACKNOWLEDGEMENTS Financial support from Projects NANOMAX (Italian Flagship Project MIUR PNR, 2011−2013) and NEWTON (Italian MIUR FIRB, RBAP11BYNP, 2011−2014) is gratefully acknowledged.



Page 24 of 29

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(1)

Pacholski, C.; Sartor, M.; Sailor, M. J.; Cunin, F.; Miskelly, G. M. J. Am. Chem. Soc. 2005, 127, 11636–11645.

(2)

Virga, A.; Rivolo, P.; Descrovi, E.; Chiolerio, A.; Digregorio, G.; Frascella, F.; Soster, M.; Bussolino, F.; Marchiò, S.; Geobaldo, F.; Giorgis, F. J. Raman Spectrosc. 2012, 43, 730–736.

(3)

Frascella, F.; Ricciardi, S.; Pasquardini, L.; Potrich, C.; Angelini, A.; Chiadò, A.; Pederzolli, C.; Leo, N. De; Rivolo, P.; Pirri, C. F.; Descrovi, E. Analyst 2015, 140, 5459–5463.

(4)

Novara, C.; Lamberti, A.; Chiadò, A.; Virga, A.; Rivolo, P.; Geobaldo, F.; Giorgis, F. RSC Adv. 2016, 6, 21865–21870.

(5)

Quagliano, L. G. J. Am. Chem. Soc. 2004, 126, 7393–7398.

(6)

Lamberti, A.; Virga, A.; Chiadò, A.; Chiodoni, A.; Bejtka, K.; Rivolo, P.; Giorgis, F. J. Mater. Chem. C 2015, 3, 6868–6875.

(7)

Cong, S.; Yuan, Y.; Chen, Z.; Hou, J.; Yang, M.; Su, Y.; Zhang, Y.; Li, L.; Li, Q.; Geng, F.; Zhao, Z. Nat. Commun. 2015, 6, 7800.

(8)

Martinsson, E.; Otte, M. A.; Shahjamali, M. M.; Sepulveda, B.; Aili, D. J. Phys. Chem. C 2014, 118, 24680–24687.

(9)

Lamberti, A.; Virga, A.; Angelini, A.; Ricci, A.; Descrovi, E.; Cocuzza, M.; Giorgis, F. RSC Adv. 2015, 5, 4404–4410.

(10)

Citartan, M.; Gopinath, S. C. B.; Tominaga, J.; Tan, S.; Tang, T. Biosens. Bioelectron. 2012, 34, 1–11.

(11)

Novara, C.; Petracca, F.; Virga, A.; Rivolo, P.; Ferrero, S.; Chiolerio, A.; Geobaldo, F.; Porro,


Page 25 of 29


S.; Giorgis, F. Nanoscale Res. Lett. 2014, 9, 1–7. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(12)

Liu, J. Curr. Opin. Cell Biol. 2008, 20, 214–221.

(13)

Bartel, D. P. Cell 2004, 116, 281–297.

(14)

Cannell, I. G.; Kong, Y. W.; Bushell, M. Biochem. Soc. Trans. 2008, 36, 1224–1231.

(15)

Jackson, J. R.; Standart, N. Sci. STKE 2007, re1, 1–13.

(16)

Dong, H.; Lei, J.; Ding, L.; Wen, Y.; Ju, H.; Zhang, X. Chem. Rev. 2013, 113, 6207–6233.

(17)

Tian, T.; Wang, J.; Zhou, X. Org. Biomol. Chem. 2015, 13, 2226–2238.

(18)

Catuogno, S.; Esposito, C. L.; Quintavalle, C.; Cerchia, L.; Condorelli, G.; de Franciscis, V. Cancers (Basel). 2011, 3, 1877–1898.

(19)

Davis, S.; Lollo, B.; Freier, S.; Esau, C. Nucleic Acids Res. 2006, 34, 2294–2304.

(20)

Lange, S. a; Benes, V.; Kern, D. P.; Hörber, J. K. H.; Bernard, A. Anal. Chem. 2004, 76, 1641– 1647.

(21)

Elsholz, B.; Worl, R.; Blohm, L.; Albers, J.; Feucht, H.; Grunwald, T.; Jurgen, B.; Schweder, T.; Hintsche, R. Anal. Chem. 2006, 78, 4794–4802.

(22)

Jimenez-Monroy, K. L.; Kick, A.; Eersels, K.; van Grinsven, B.; Wagner, P.; Mertig, M. Phys. Status Solidi 2013, 925, 918–925.

(23)

Chrisey, L. a; Lee, G. U.; O’Ferrall, C. E. Nucleic Acids Res. 1996, 24, 3031–3039.

(24)

Driskell, J. D.; Seto, a. G.; Jones, L. P.; Jokela, S.; Dluhy, R. a.; Zhao, Y. P.; Tripp, R. a. Biosens. Bioelectron. 2008, 24, 917–922.



(25) 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Bi, L.; Rao, Y.; Tao, Q.; Dong, J.; Su, T.; Liu, F.; Qian, W. Biosens. Bioelectron. 2013, 43, 193– 199.

(26)

Prado, E.; Daugey, N.; Plumet, S.; Servant, L.; Lecomte, S. Chem. Commun. 2011, 47, 7425– 7427.

(27)

Wang, H.; Crawford, B. M.; Fales, A. M.; Bowie, M. L.; Seewaldt, V. L.; Vo-dinh, T. J. Phys. Chem. C 2016, Article ASAP.

(28)

Li, X.; Ye, S.; Luo, X. Chem. Commun. 2016, 52, 10269–10272.

(29)

Fan, M.; Andrade, G. F. S.; Brolo, A. G. Anal. Chim. Acta 2011, 693, 7–25.

(30)

Novara, C.; Marta, S. D.; Virga, A.; Lamberti, A.; Angelini, A.; Chiadò, A.; Rivolo, P.; Geobaldo, F.; Sergo, V.; Bonifacio, A.; Giorgis, F. J. Phys. Chem. C 2016, 120, 16946−16953.

(31)

Barhoumi, A.; Zhang, D.; Tam, F.; Halas, N. J. J. Am. Chem. Soc. 2008, 130, 5523–5529.

(32)

Liu, Y.; Irving, D.; Qiao, W.; Ge, D.; Levicky, R. J. Agric. Technol. 2011, 133, 11588–11596.

(33)

Peterson, A. W.; Heaton, R. J.; Georgiadis, R. M. Nucleic Acids Res. 2001, 29, 5163–5168.

(34)

Braun, G.; Lee, S. J.; Dante, M.; Nguyen, T.-Q.; Moskovits, M.; Reich, N. J. Am. Chem. Soc. 2007, 129, 6378–6379.

(35)

DeRisi, J. L.; Iyer, V. R.; Brown, P. O. Science (80-. ). 1997, 278, 680–686.

(36)

Lashkari, D. A.; DeRisi, J. L.; McCusker, J. H.; Namath, A. F.; Gentile, C.; Hwang, S. Y.; Brown, P. O.; Davis, R. W. Proc. Natl. Acad. Sci. U. S. A. 1997, 94, 13057–13062.

(37)

Page 26 of 29

Peterlinz, K. a.; Georgiadis, R. M.; Herne, T. M.; Tarlov, M. J. J. Am. Chem. Soc. 1997, 119, 3401–3402.


Page 27 of 29


(38) 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Petrovykh, D. Y.; Pérez-Dieste, V.; Opdahl, A.; Kimura-Suda, H.; Sullivan, J. M.; Tarlov, M. J.; Himpsel, F. J.; Whitman, L. J. J. Am. Chem. Soc. 2006, 128, 2–3.

(39)

Herne, T. M.; Tarlov, M. J. J. Am. Chem. Soc. 1997, 119, 8916–8920.

(40)

Bettazzi, F.; Hamid-Asl, E.; Esposito, C. L.; Quintavalle, C.; Formisano, N.; Laschi, S.; Catuogno, S.; Iaboni, M.; Marrazza, G.; Mascini, G.; Cerchia, G.; De Franciscis, V.; Condorelli, G.; Palchetti, I. Anal. Bioanal. Chem. 2012, 405, 1025–1034.

(41)

Teixeira, A. L.; Silva, J.; Ferreira, M.; Marques, I.; Gomes, M.; Mauricio, J.; Lobo, F.; Medeiros, R. Eur. J. Cancer 2012, 48, S216.

(42)

Fabris, L.; Ceder, Y.; Chinnaiyan, A. M.; Jenster, G. W.; Sorensen, K. D.; Tomlins, S.; Visakorpi, T.; Calin, G. A. Eur. Urol. 2016, 70, 312–322.

(43)

Zhao, Y.; Wang, Y.; Yang, Y.; Liu, J.; Song, Y.; Cao, Y.; Chen, X.; Yang, W.; Wang, F.; Gao, J.; Li, Z.; Yang, C. J. Cancer 2015, 6, 1230–1235.

(44)

Lamberti, A.; Garino, N.; Sacco, A.; Bianco, S.; Chiodoni, A.; Gerbaldi, C. Electrochim. Acta 2015, 151, 222–229.

(45)

Chiadò, A.; Palmara, G.; Ricciardi, S.; Frascella, F.; Castellino, M.; Tortello, M.; Ricciardi, C.; Rivolo, P. Colloids Surfaces B Biointerfaces 2016, 143, 252–259.

(46)

Palmara, G.; Chiadò, A.; Calmo, R.; Ricciardi, C. 2016, Analytical and Bioanalytical Chemistry. accepted – in press.

(47)

Pettersen, E. F.; Goddard, T. D.; Huang, C. C.; Couch, G. S.; Greenblatt, D. M.; Meng, E. C.; Ferrin, T. E. J. Comput. Chem. 2004, 25, 1605–1612.

(48)

Petrovykh, D. Y.; Kimura-Suda, H.; Whitman, L. J.; Tarlov, M. J. J. Am. Chem. Soc. 2003, 125,



Page 28 of 29

5219–5226. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(49)

Liang, Y.; Ridzon, D.; Wong, L.; Chen, C. BMC Genomics 2007, 8, 166.

(50)

Lamberti, A.; Virga, A.; Rivolo, P.; Angelini, A.; Giorgis, F. J. Phys. Chem. B 2015, 119, 8194– 8200.

(51)

Jarvis, K. L.; Barnes, T. J.; Prestidge, C. A. Langmuir 2008, 24, 14222–14226.

(52)

Bellet, D.; Canham, L. Adv. Mater. 1998, 10, 487–490.

(53)

Song, J. H.; Sailor, M. J. Inorg. Chem. 1998, 37, 3355–3360.

(54)

Janshoff, A.; Dancil, K. S.; Steinem, C.; Greiner, D. P.; Lin, V. S.; Gurtner, C.; Motesharei, K.; Sailor, M. J.; Ghadiri, M. R. J. Am. Chem. Soc. 1998, 120, 12108–12116.

(55)

Fu, R.; Wang, C.; Zhuang, J.; Yang, W. Colloids Surfaces A Physicochem. Eng. Asp. 2014, 444, 326–329.

(56)

Su, T. J.; Lu, J. R.; Thomas, R. K.; Cui, Z. F. J. Phys. Chem. B 1999, 103, 3727–3736.

(57)

Tarasevich, Y. I.; Monakhova, L. I. Colloid J. 2002, 64, 482–487.

(58)

David, C.; Guillot, N.; Shen, H.; Toury, T.; Lamy De La Chapelle, M. Nanotechnology 2010, 21, 475501.

(59)

Giese, B.; McNaughton, D. Phys. Chem. Chem. Phys. PCCP 2002, 4, 5171–5182.

(60)

Najjar, S.; Talaga, D.; Coffinier, Y.; Szunerits, S.; Boukherroub, R.; Servant, L.; Rodriguez, V.; Bonhommeau, S. J. Phys. Chem. C 2014, 118, 1174–1181.


Page 29 of 29


Table of Contents (TOC) 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Immobilization of Oligonucleotides on Metal-Dielectric Nanostructures

Recommend Documents