Empowering Electroless Plating to Produce Silver Nanoparticle Films

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Empowering Electroless Plating to Produce Silver Nanoparticle Films for DNA Biosensing Using Localized Surface Plasmon Resonance Spectroscopy Falk Muench, Aleksei Solomonov, Tatyana Bendikov, Leopoldo Molina-Luna, Israel Rubinstein, and Alexander Vaskevich ACS Appl. Bio Mater., Just Accepted Manuscript • DOI: 10.1021/acsabm.8b00702 • Publication Date (Web): 23 Jan 2019 Downloaded from http://pubs.acs.org on January 26, 2019

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ACS Applied Bio Materials

Empowering Electroless Plating to Produce Silver Nanoparticle Films for DNA Biosensing Using Localized Surface Plasmon Resonance Spectroscopy Falk Muench,†,‡,┴,* Aleksei Solomonov,†,‡ Tatyana Bendikov,§ Leopoldo Molina-Luna,⸸ Israel Rubinstein,† ,‖ Alexander Vaskevich†,* † Weizmann Institute of Science, Department of Materials and Interfaces, 7610001 Rehovot, Israel. § Weizmann Institute of Science, Department of Chemical Research Support, 7610001 Rehovot, Israel. ⸸ Technische Universität Darmstadt, Department of Materials and Earth Sciences, 64287 Darmstadt, Germany. ‖ This work is dedicated to the memory of Prof. Israel Rubinstein.

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ABSTRACT. To facilitate the implementation of biosensors based on the localized surface plasmon resonance (LSPR) of metal nanostructures, there is a great need for cost-efficient, flexible, and tunable methods for producing plasmonic coatings. Due to its simplicity and excellent conformity, electroless plating (EP) is well suited for this task. However, it is traditionally optimized to produce continuous metal films, which cannot be employed in LSPR sensors. Here, we outline the development of an EP strategy for depositing island-like silver nanoparticle (NP) films on glass with distinct LSPR bands. The fully wet-chemical process only employs standard chemicals, proceeds within minutes and at room temperature. The key step for producing spreadout NP films is an accelerated ripening of the silver seed layer in diluted hydrochloric acid, which reduces the nucleation density during plating. The reaction kinetics and mechanisms are investigated with scanning (transmission) electron microscopy (SEM / STEM), X-ray photoelectron spectroscopy (XPS), and UV-Vis spectroscopy, the latter enabling a convenient live monitoring of the deposition, allowing its termination at a stage of desired optical properties. The sensing capabilities of chemically deposited NP films as LSPR transducers are exemplified in DNA biosensing. To this end, a sensing interface is prepared using layer-by-layer (LbL) buildup of polyelectrolytes, followed by adsorption and covalent immobilization of ssDNA. The obtained LSPR transducers demonstrate robustness and selectivity in sensing experiments with binding complementary and unrelated DNA strands.

KEYWORDS. Seed-mediated nanoparticle growth; electroless silver plating; hydrochloric acid; localized surface plasmon resonance; layer-by-layer deposition; DNA biosensing.


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1. INTRODUCTION LSPR sensors, employing separated noble metal nanostructures, have attracted considerable attention as a new class of plasmonic nanosensors and arrays compatible with simple and lowcost optics.1–4 The electromagnetic field around such plasmonically excited nanostructures decays over some tens of nanometers, providing confined sensing volume which suit the size regime of medically important biomacromolecules, allowing both the detection of binding events and biomolecule transformations.2 Binding of the analyte alters the refractive index of the immediate metal nanostructure’s environment and the surface plasmon resonance conditions, resulting optically readable changes in the LSPR band position / shape / intensity.4–7 Aside the unique capabilities of LSPR-based biosensors, the interest in this field is fuelled by the opportunities provided by employing nanoplasmonic materials in miniaturized and integrated devices.8,9 For instance, combining plasmonic materials with microfluidic10 or fiber optic setups3,11 provides avenues for multiplexing and point-of-care diagnostics. Accordingly, the development of scalable and cost-efficient tools for LSPR transducer fabrication is subject of intensive research.3 EP represents an industrially established solution deposition technique for the conformal metallization of work pieces, which is traditionally employed to outfit substrates with protective or electrically conducting coatings.12 With increasing momentum, the nanotechnology community adapts EP to its specific needs, making its beneficial characteristics available for the production of functional nanomaterials. Contrary to competing metal deposition methods such as sputter coating or vacuum evaporation, EP is capable of creating homogeneous metal films on complex shaped substrates and interior surfaces,12–14 including challenging variants such as high aspect ratio submicron channels15 and frameworks with multilevel hierarchy.16 It is applicable to a wide range of substrate materials


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and only requires stock chemicals, allowing the production of defined nano-architectures in any laboratory with basic wet-chemical equipment.17,18 The flexibility regarding substrate material and shape renders EP well suited for introducing functional materials into flow setups19–22 and fiber optics.23 Recently, EP is being introduced to preparation of densely packed metal nanofilms for surfaceenhanced Raman spectroscopy18,24,25 or surface plasmon resonance biosensors.23 All these examples comprise broad surface plasmon absorption band and are not suitable for LSPR spectroscopy. The exceptional plasmonic properties of silver, in particular its low plasmonic damping,1,26 make the preparation of LSPR transducers using electroless silver plating particularly interesting. The lack of EP protocols for fabricating island-like NP films and LSPR biosensors is inherently linked to its deposition mechanism and its traditional optimization towards closed coatings. In conventional EP, substrates are sensitized with Sn2+, which has reducing qualities. When brought into contact with noble metal ion solution, such sensitized substrates get covered with a dense NP layer.27 These NPs act as seeds and initiate metal deposition, resulting in the formation of percolating films.28,29 In order to create island-like NP films with distinct LSPR bands, the obtained metal films can be subjected to thermal dewetting,28 which however adds to the complexity of the process and subjects the sample / supporting device to thermal stress / damage. Alternative seeding strategies, such as anchoring colloidal NPs via silane chemistry, are more involved regarding the time need and the number of reaction steps (silane self-assembly, baking of silane layer, NP synthesis, purification and attachment), and also produce comparably dense films upon EP.24,30,31 In this study, we show how the solely solution-based electroless seeding and deposition approach can be expanded toward the target morphology of spread-out silver NP films. We


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demonstrate the applicability of the newly developed LSPR transducers based on EP films in DNA biosensing using oligonucleotides as the analyte. DNA biosensors and gene chips are of major interest due to their great potential for obtaining sequence-specific information in a faster, simpler and cheaper manner compared to the traditional hybridization.32 Our LSPR transducer is based on silver NPs deposited on glass, onto which polyelectrolyte multilayer (PEM) is assembled by the LbL method,33 which in turn are used to immobilize DNA recognition elements. Hence, each surface modification step (polyelectrolyte adsorption, DNA probe immobilization, target DNA adsorption) can be optically monitored. Additional labelling of the employed oligo-DNA strands with dyes allowed us to verify the DNA probe immobilization, to investigate the binding selectivity, and to assess the stability of the sensing interface.

2. EXPERIMENTAL SECTION 2.1 Materials and Reagents 4-dimethylaminopyridine (≥ 99.0%, Fluka); ammonia solution (25%, Merck, p.a.); cover slips (Menzel Gläser); DNase- and RNase-free ultra-pure water (Biological Industries); ethanol (Gadot, absolute AR); formaldehyde solution 36.5%, stabilized with methanol (puriss. p.a., Fluka); glutaraldehyde (50% in H2O, Sigma-Aldrich, grade I); HEPES buffer solution (1 M, pH = 7.3, Biological Industries); hydrochloric acid (32%, Bio-Lab, AR); nitric acid (65%, Merck, p.a.), magnesium chloride solution (1.00 M ± 0.01 M, Sigma-Aldrich, for molecular biology); polyallylamine hydrochloride (56 kDa, Sigma-Aldrich); polystyrene sulfonate sodium salt (70


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kDa, Polysciences Inc.); potassium sodium tartrate tetrahydrate (Sigma-Aldrich, ReagentPlus ≥ 99%); sodium chloride (Sigma-Aldrich, AR grade); silver nitrate (Fluka, Ph. Eur. puriss. p.a.); sodium cyanoborohydride (NaBH3CN 95%, Sigma-Aldrich, reagent grade); sodium tetrachloroaurate(III) dihydrate (99%, Sigma-Aldrich); tin dichloride dihydrate (Aldrich, 99.995+ %) were used as received. The oligo-DNA samples synthesized by standard phosphoramidite chemistry were obtained from IDT-Syntezza, Israel. They contain the following 23-base nucleotide sequences: (i) ssDNA 155-Cy5_AmMO, which is complementary to miRNA-155: 5'-/Cy5/ TTA ATG CTA ATC GTG ATA GGG GT /AmMO/-3' with Cy5 dye on 5'-prome and amine (AmMO) modifier on 3'-prime; (ii) ssDNA 155c-D2, which is complementary to 155-Cy5_AmMO: 5'-/D2/ ACC CCT ATC ACG ATT AGC ATT AA-3' ssDNA with WellRED D2 dye on 5'-prime; (iii) ssDNA 191-D2, which is complementary to miRNA-191 and unrelated to ssDNA 155-Cy5_AmMO: 5'-/D2/ AAA CAG AAT ACC AAG AGC AGC TG-3' with WellRED D2 dye on 5'-prime. Triply distilled water and ultra-pure water were used in preparation on transducers and sensing experiments with DNA experiments, respectively. Stock solutions of ssDNA (100 µM) were prepared from purchased DNA and stored at –20 ºC. For sensing experiments the DNA stock solutions were diluted to 10 µM with ultrapure water following further addition of hybridization buffer (HB) to obtain 5 µM solutions. The HB solution (100 mM HEPES, 100 mM NaCl, 15 mM MgCl2, pH = 7.3) used in all biochemical experiments was prepared from 1 M stock solutions. The nitrogen used for slide drying was produced in-house from liquid N2. The transducer fabrication and DNA biosensing was carried out in a climate-controlled laboratory (temperature: 22.5±1.5 °C; relative humidity: 45±7 %).


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2.2 Nanoparticle Deposition Silver seeds were introduced using the classical substrate sensitization / activation strategy of EP.15,21,27–29,34 Cover slips were cut into 9 mm × 22 mm strips and cleaned with fresh piranha solution (concentrated H2SO4 + 30% H2O2 solution, 3:1 volume ratio). After rinsing with water, the glass strips were immersed in a solution of SnCl2 (20 mM) and HCl (55 mM) in a 1:1 mixture of ethanol and water for 1 min. To remove supernatant sensitization solution, the slips were washed with a 1:1 ethanol-water mixture, followed by water, and submerged in an aqueous solution of AgNO3 (20 mM) and NH3 (80 mM) for 1 min. After seeding, the glass strips were washed with water, and either transferred to the silver plating bath directly, or after a treatment with HCl (3 M) for 30 s, followed by rinsing with water. The silver plating bath contained AgNO3 (20 mM), ammonia (100 mM), potassium sodium tartrate (120 mM), and nitric acid (6 mM). After the desired deposition time, the samples were taken out of the plating bath and rinsed with water. Gold plating was performed according to a previously published protocol,35 which was adapted to yield spherical NPs instead of gold nanowires. Briefly, the deposition bath was operated at 70 °C, and prepared by complexing NaAuCl4 (7.5 mM) with 4dimethylaminopyridine (30 mM) before adding the reducing agent formaldehyde (400 mM). 2.3 Transducer Fabrication Immediately after preparation, the silver coated slips were transferred from water to HB for equilibration (45 min). Polyelectrolyte solutions were prepared by dissolving either polyallylamine hydrochloride (PAH) or polystyrene sulfonate sodium salt (PSS) in HB solution (1 mM with respect to the monomer, corresponding to 0.093 mg mL-1 PAH and 0.206 mg mL-1 PSS). The slides were alternatingly immersed in vials containing the PE solutions for 5 min each,


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starting with PAH. After each adsorption step, the slide was rinsed with water, dipped into HB, and immersed in the other polyelectrolyte solution to obtain a modified transducer with a stack of three layers. To determine the decay length of the transducer, the accumulation of the PEM was continued following the same procedure till the saturation of its optical response. We recently improved previously employed protocols for preparing DNA-terminated PEMs36– 38

by covalently binding the ssDNA to the PEM. This prevents desorption of oligo-DNA receptor

during further exposure of the transducer to the analyte. To obtain a monolayer of DNA-receptor a 30 µL drop of 5 µM ssDNA Cy5-155_AmMO was placed on the polyelectrolyte-modified surface and incubated in a closed Petri dish at a room temperature for 30 min, followed by washing with HB solution to remove unbound DNA. The samples were then immediately subjected to a cross-link procedure, without drying. First, a 50 µL drop of 10% glutaraldehyde was spread onto the polyelectrolyte-modified slide, followed by incubation in a humidity chamber for 2 h. The samples then were washed with water to remove the excess of cross-linker, covered with a 50 μL drop of NaBH3CN solution (10 mM) and left for 1 h in a humidity chamber, followed by washing with water, and drying under a nitrogen stream. A more detailed study of the cross-linked DNA-PEM interface will be published elsewhere. 2.4 DNA Biosensing A 30 µL drop containing 5 μM of complementary ssDNA 155c-D2 or unrelated ssDNA 191D2 strands was placed on a crosslinked DNA-polyelectrolyte modified silver transducer surface and left for 1 h for incubation in a humidity chamber. Next, the samples were washed with HB and water to remove excess DNA, dried under a nitrogen stream, and spectra were measured ex situ.


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2.5 Characterization Transmission UV–vis spectra were obtained with Varian CARY 50 and CARY 300 spectrophotometers. The spectra were recorded in the range of 400–900 nm with wavelength resolution 1 nm and acquisition time 1 sec per point. Baseline was taken in air, HB or silver plating solution in corresponding measurements. Samples measured ex situ were dried from solvent under a nitrogen stream and placed in a special holder which enables interrogation of the same spot on the slide during all stages of the experiment. In situ measurements were performed in quartz cuvettes filled with HB in the case of polyelectrolyte-coated silver transducers, and with plating solution in the case of the of silver deposition. Measurements of the UV-vis spectra after seeding steps (sensitization, activation, ripening in HCl) were performed in water. Surface plasmon wavelength and extinction intensity was determined using a custom Matlab routine for a high-degree polynomial fit of experimental data.39 SEM measurements were performed with a Zeiss Sigma 500 microscope in secondary electron detection mode. STEM imaging was performed in combination with energy-dispersive X-ray spectroscopy (EDX) using an aberration-corrected JEOL JEM ARM-200F (scanning) transmission electron microscope, equipped with a “Schottky” field-emission gun. The microscope was operated at 120 kV to reduce beam damage. XPS measurements were carried out using a monochromatic Al Kα X-ray source (hυ = 1486.6 eV) at 75W and detection pass energies ranging between 20 and 80 eV. A low-energy electron flood gun was applied for charge neutralization. To define the binding energies (BE) of silver and Sn, the C 1s line at 284.8 eV was taken as a reference. Curve fitting analysis was based on


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linear, Shirley or Tougaard background subtraction and application of Gaussian-Lorenzian line shapes.

3. RESULTS AND DISCUSSION 3.1 Electroless Plating of Silver Nanoparticle Films. The NP film deposition proceeds via seeding of the glass substrate, followed by seed amplification with EP (Figure 1). While the experiments shown below were performed using microscopy glass as substrate for the NP film, the procedure can be applied to other substrate materials as well, such as quartz glass or polycarbonate (Supporting Information, section 1).

Figure 1. Scheme of the silver NP film syntheses, including the two-step sensitization / activation procedure for introducing the silver seeds, which are optionally treated with HCl, and


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finally amplified via EP. The SEM images show the seeded and plated states of the pristine and ripened sample variants.

Seeding was performed by immersing the substrate in a SnCl2 solution, resulting in the adsorption of Sn2+ (Figure 1, step 1). After removing excess Sn2+ by washing, the substrate is immersed in an ammoniac AgNO3 solution, in which the superficially bound Sn2+ reduces Ag+ to elemental metal (eq. 1), resulting in the formation of silver NPs of typically < 10 nm size (Figure 1, step 2).27 Sn2+ads + 2[Ag(NH3)2]+ → Sn4+ads + 2Ag↓ + 4NH3

(1)

In the corresponding SEM image, the high NP density of the as-seeded substrate is apparent. We chose the seeds to match the metal applied later during EP, in order to avoid the intricacies associated with bimetallic systems (such as the formation of core-shell structures, alloying, and galvanic replacement), which affect the optical response of the resulting materials in a complex manner.40 Nevertheless, the general methodology is compatible with the seeded growth of other nanoparticulate metal films, such as the important case of gold,31 as was demonstrated in preliminary experiments (Supporting Information, section 2). Consequently, the substrates are subjected to silver plating. The bath employed here is designed to combine a high stability with a balanced reaction rate, so that nanoscale silver deposits are produced in a convenient timeframe of minutes, but not so fast that the end point is difficult to control. It is solely based on readily available chemicals (AgNO3 as the metal source, NH3 as the ligand, and sodium potassium tartrate as the reducing agent). HNO3 is added for adjusting the bath reactivity, which is lowered in less alkaline media.41 Applying EP to the


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seeded glass substrates resulted in the quick formation of percolating NP films (Figure 1, step 2B), such as observed in previous studies.28,29 We overcome this issue by applying an accelerated ripening protocol to our seed layer, resulting in a decimation of seeds. In turn this allows the realization of an island-like morphology in the final silver NP films. By immersing the seeded glass substrate in 3 M HCl for just 30 s, we find a clearly separated appearance of the NPs both before (Figure 1, step 3) and after the plating step (Figure 1, step 3B). We attribute this to the promoting effect of halide ions on the oxidation and morphological transformation of silver nanostructures, which are exceptionally prone to dissolution in the presence of hydrochloric acid.42 Due to halide-assisted silver dissolution and redeposition, silver atom mobility is greatly enhanced, which was previously employed for the low-temperature welding of silver nanowires.43,44 In our nanoparticulate system, we expect these two aspects to manifest in boosted particle dissolution and Ostwald ripening,45 which is in agreement with the observed structural changes (reduced nucleation site density, resulting in the formation of spread-out NPs during EP). By changing the HCl concentration and exposition time, the degree of seed transformation – and subsequently, the structure of the plated NP film – can be adjusted (Supporting Information, section 3). The seed layer ripening proceeds within two distinct time regimes: Upon immersion of silver-seeded substrates in hydrochloric acid, an almost immediate color change from brown to yellowish is observed, which corresponds to the blue-shift of the silver plasmon band in Figure 2A. This process is superimposed by a much slower, continuous decrease of the plasmon band intensity during storage of the seeded substrate in acid, which can be related to ongoing silver oxidation (Supporting Information, Figure S6).


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Figure 2. (A) UV-Vis spectra of glass substrates after sensitization, activation and ripening, corresponding to the synthesis steps 1–3 in Figure 1. (B) In situ UV-Vis spectra monitoring the silver deposition during EP. (C-E) SEM images of the silver NP films after plating times of (C) 6 min, (D) 10 min, and (E) 20 min.

Due to the plasmonic properties of silver, the processes occurring during seeding and NP amplification was conveniently monitored by UV-Vis spectroscopy (Figure 2A, B). Following the adsorption of Sn2+ (Figure 2A, blue dotted curve), the deposition of the silver seeds is indicated by the appearance of a plasmon peak with a maximum at ~430 nm (Figure 2A, bordeaux red curve). Upon the HCl treatment, this peak is drastically narrowed, and blue-shifted to ~395 nm (Figure 2A, orange curve). We can exclude the formation of a silver chloride shell during treatment with HCl because in such a case, a red-shift of the plasmon is expected due to the presence of a high refractive index medium around the metallic core.1 The blue-shift and


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peak-narrowing are probably caused by NP reshaping during partial dissolution, and reflect the formation of an ensemble of small symmetrical silver NPs with narrow size distribution.46 Ongoing autocatalytic deposition of silver on the NPs results in their continuous growth (Figure 2C-E). At 6, 10 and 20 min of plating, the silver NPs reach effective projected diameter of 33 ± 17, 48 ± 13 and 64 ± 19 nm, respectively (Figure 2C-E). Correspondingly, the plasmon peak intensity gradually increases, while its maximum shifts red to ~450 nm (Figure 2B), which is characteristic for larger silver NPs of sizes of about 40–80 nm.47,48 The development of a shoulder at ca. 650 nm (Figure 2B) is related to an increased asymmetry of the NPs due to overlap between growing crystals, which can be clearly seen after 20 min of EP (Figure 2E). Silver deposition can be furthered to obtain particles of several hundreds of nm in size (Supporting Information, section 4). XPS was employed to gain further insights into the NP film formation (Figure 3), analyzing each synthesis step along the path to sample type 3B as shown in Figure 1. Chlorine was present at trace levels in the pristine glass substrates, and its content did not change significantly during synthesis (< 0.09 at% in all cases). Notably, the HCl treatment did not result in increasing chloride levels, corroborating the absence of solid AgCl as a silver corrosion product, as suggested by the UV-Vis spectra. Although the employed charge neutralization, in conjunction with the distinguishability issues of the relevant silver and tin species (Ag0 / Ag+, Sn2+ / Sn4+)49,50 complicate the speciation analysis, several important points can be made: The mean Ag 3d5/2 peak energy of 368.1 eV supports the interpretation as metallic silver,49 whereas the tailing at the higher binding energy flank (clearly exceeding the natural asymmetry


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found for elemental bulk Ag)49 indicates the presence of oxidized silver at the NP surface. The mean Sn 3d5/2 energy of 486.9 eV corresponds to oxidized tin species (Sn2+ / Sn4+).50

Figure 3. XPS spectra of the Sn 3d and Ag 3d binding energy regions of 0) pristine glass substrates, 1) after sensitization, 2) after activation, 3) after seed ripening, and 3B) after 2 min of electroless silver plating, including the evaluation of the tin and silver content in atomic percent (at%) in the surface layer at each stage.

During the different synthetic stages, the elemental ratios of silver and tin in the XPS spectra change considerably (Figure 3). While the inhomogeneous structure of the film (most importantly, the granular nature of the Ag deposit) in conjunction with the limited penetration depth of the characterization method restrict the precision of our quantification, the observed trends clearly confirm the mechanistic interpretation of our system based on the previously shown SEM results (Figure 1): Sensitization and activation lead to the introduction of tin and silver at comparably high concentrations. The HCl treatment results in an almost complete


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removal of tin (-90%) and a pronounced reduction of the silver content (-80%). While the former is known from the activity enhancement of palladium seed layers via removal of inert tin oxides,51,52 the latter nicely illustrates the loss and concentration of silver in the form of fewer, less distributed particles during HCl aging in our case. Upon EP, the silver content increases again, corresponding to the NP growth. An in-depth study of the dynamic nanoscale transformations during the HCl treatment with in situ liquid cell TEM experiments is out of the scope of present paper. To elucidate the nanostructural and compositional changes occurring in our system at higher resolution, we performed ex situ STEM imaging in combination with EDX. For this purpose, TEM grids covered with ultrathin “holey” carbon films were slued in the reaction and washing solutions as outlined above, to allow for a direct investigation of the seed layers deposited onto them (Figure 4). While the difference in substrate material must be taken into account, the nanostructure of both the as-deposited and the ripened seed layer resemble those of their counterparts on glass. After tin sensitization and silver activation (Figure 1, step 2), a grainy / fibrous, comparably dense film is obtained, in which the silver content is highly scattered and mixed with tin (Figure 4A, B). Aging in hydrochloric acid (Figure 1, step 3) removes much of the material, and causes the silver to concentrate in the form of distinct, roundish, and mostly enlarged particles (Figure 4C, D), providing direct experimental evidence for the accelerated silver migration.


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Figure 4. Bright field STEM images of the seed layer in the as-deposited state (A, B) and after treatment with hydrochloric acid (C, D). The insets show representative EDX point spectra which were recorded at contrast-rich areas (Sn-Ag sample) and large, solitary particles (Sn-AgHCl sample), respectively.

3.2 Optical Properties of LSPR Transducers based on Silver Nanoparticle Films. To evaluate the analytical volume of the Ag NP films, we measured the optical response of films obtained after 2 and 6 min of EP to the build-up of PEMs (Figure 5A, B), similar to our previous study on Au island films.6 In the current work, UV-vis spectra were measured in situ, avoiding morphological changes in the plasmonic system which may influence the optical response.


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Figure 5. UV-Vis spectra of silver NP films obtained after (A) 2 min and (B) 6 min of electroless silver plating, recorded in the pristine state and during the sequential adsorption of PEM. Right to the spectra, SEM images of the NP films covered with 8 bilayers are shown.

The build-up of PEMs increases the refractive index of the particle-surrounding medium, resulting in a red-shift and intensity increase of the LSPR bands with increasing shell thickness (Figure 5A, B). We evaluated the thickness of the PEMs using XPS data on the intensity attenuation of the underlying substrate due to PEM accumulation.53 We found that the thickness of the PAH-PSS bilayer obtained with HEPES buffer is ca. 2.5 nm, compared to 2.1 nm previously obtained for PEMs accumulated with NaCl electrolytes.54 Aside the differing electrolytes, the difference might also be caused by the PEM preparation procedure. In the current study, PEM accumulation was performed without drying of the sample, while intermittent PEM drying was employed in the previous work.54


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Figure 6 summarizes the experimental data on the plasmon peak shift obtained during PEM build-up combined with fitting to the exponential dependence of the optical response commonly used for the description of plasmonic systems1,55 Δλ = m Δη [1-e(-2d/l)]

(2)

where Δλ is the wavelength shift, m is the bulk refractive index sensitivity (RIS), Δη = nPEMnaq is the change in refractive index (nPEM, naq) of the surrounding medium effected by the adsorbate, d is the PEM thickness, and l is the plasmon decay length.

Figure 6. Dependence of the shift of surface plasmon band on thickness of PEM for silver films plated for different time as in Figure 4. Solid lines present the fitting of the experimental data to the exponential decay of the optical response.

As seen in Figure 6, the exponential fit to eq. 2 perfectly describes the experimental data. Assuming the commonly used values of the dielectric constants of the PEM and the solution of


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1.5 and 1.33, respectively,1,54 resulting in Δη = 0.17, we calculated the RIS and the decay length of the silver LSPR transducers, which are presented in Table 1.

Table 1. Quantification of the optical response of the silver NP films. Film type

Refractive index sensitivity, RIU-1

Decay length, nm

2 min deposition

90.8 ± 0.6

10 ± 0.2

6 min deposition

165.3 ± 1.2

13.6 ± 0.4

The values of the bulk refractive index sensitivity of our EP silver films are well comparable to those of silver island films prepared using nanoscale lithography.56,57 The decay length of the exemplified Ag NP films is comparable to the typical dimensions of biomolecules and can be tuned by the deposition conditions, allowing the optimization of the analytical volume of the LSPR transducer.6 The sensing parameters of both silver NP films are well suited for typical biosensing applications. In further demonstration of the sensing capabilities of our LSPR transducers, we focused on the preparation of a robust recognition interface. The optimization of the transducers to specific analytes (in our case, oligo-DNA), in conjunction with a thorough analysis of their selectivity and detection threshold, was out of the scope of the current study. A detailed investigation of the biosensing aspects of our interface (e.g., DNA hybridization efficiency, DNA melting and single- / double-base mismatch behavior) is underway and will be reported separately.58


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To demonstrate the sensing capabilities of the Ag NP films, we chose the 6 min deposited sample due to its higher RIS value. An extended deposition time of 10 min yielded results quite similar to the 6 min sample, with slightly increased refractive index sensitivity (see Supporting Information, section 5).

3.3 LSPR DNA Biosensing Using Plated Silver Nanoparticle Films. As previously mentioned, LbL adsorption of PEMs represents a convenient technique for surface functionalization. Initially developed for a sequence of oppositely charged polymer pairs, it was extended to polymers bound by hydrogen bonds including DNA.59 The ability of ssDNA immobilized on PEMs to hybridize with corresponding complementary strands60 opens a possibility of the use ssDNA-PEM interface in DNA sensing. However, opposed to long (up to 5 kilobases) DNA adsorbates which retain the ability for specific hybridization,61 shorter oligoDNA molecules (10-50 bases) electrostatically adsorbed on PEM are prone to desorption, obstructing the use of the method in biosensing applications.38 Stabilization of the recognition interface was achieved using covalent coupling of DNA to the PEM after adsorption. Immediately after the PEM deposition on the bare silver transducer (Figure 7, steps 1–3), amine-terminated ssDNA was adsorbed on the outermost PAH layer (Figure 7, step 4), followed by cross-linking with glutaraldehyde and reduction by sodium cyanoborohydride as described in earlier (Figure 7, step 5). Exposure of the LSPR transducer functionalized with ssDNA to complementary strands produces an optically readable plasmon shift corresponding to specific binding of complementary DNA strand (Figure 7, step 6).


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Figure 7. Scheme of the preparation and utilization of the DNA biosensing interface. In the steps 1–3, a polyelectrolyte trilayer is deposited on the silver transducer surface, onto which ssDNA is adsorbed in step 4, and covalently attached by cross-linking in step 5. In the presence of the complementary DNA strand, hybridization occurs (step 6).

After obtaining the PE interface, we checked the DNA interactions of the transducer. The adsorption of ssDNA 155-Cy5_AmMO on the PEM and sequential crosslinking leads to a strong red-shift of the plasmon band by ~19 nm, which is accompanied by an increase of the extinction intensity by ~0.01. (Figure 8). Simultaneously, the absorption peak of the Cy5 dye appears at ~650 nm in the spectrum, providing direct proof of the surface immobilization of oligo-DNA (Figure 8).


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Figure 8. Transmission UV-Vis spectra of LSPR transducer prepared by silver plating for 6 min, taken after the following steps: (i) Adsorption PE trilayer (PAH-PSS-PAH (see Figure 7, step 3), (ii) Adsorption of ssDNA 155-Cy5_AmMO and sequential crosslinking (see Figure 7, step 5), (iii) exposed to either complementary (A) or unrelated (B) DNA strands.

Next, the transducers are exposed either to complementary or unrelated DNA oligonucleotides, both labelled with WellRED D2 dye. Exposure to complementary DNA strands in solution results in an altered optical response of LSPR transducer, which is characteristic for binding: The plasmon band is red-shifted by ~2.8 nm, the extinction intensity increases by ~0.024, and simultaneously, the WellRED D2 peak appears at ~770 nm (Figure 8A). On the contrary, exposure to a solution containing unrelated DNA strands doesn’t show any binding, as evidenced by the complete absence of both a LSPR band change and a WellRED D2 peak (Figure 8B). During exposure to the unrelated strand, no changes in the Cy5 region and the


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plasmon peak are observed, indicating that no DNA desorption occurred (Figure 8B), demonstrating the high stability of our sensing interface. Marking the DNA strands with two orthogonal dyes (Cy5 and D2) allowed us to distinguish the absorption bands of the immobilized molecules and to validate the label-free optical response of LSPR spectroscopy.

4. CONCLUSIONS In summary, we complemented the classical sensitization, activation and deposition chemistries of EP with an accelerated seed aging step to produce island-like silver NP films, establishing this approach as a fast, fully wet-chemical, low-temperature method for preparing LSPR biosensors. Mechanistically, the seed layer transformation is characterized by instantaneous silver NP reshaping and tin oxide dissolution, which is accompanied by continuous silver oxidation. For DNA detection, we devised a robust, PEM-based interface, utilizing covalent attachment to firmly tie a DNA oligonucleotide biorecognition unit in the active sensing volume surrounding the plasmonic NPs. Our approach yields transducers with optical properties comparable to those obtained with, e.g., lithographic methods. Furthermore, it provides straightforward means for tuning the NP density and size, is compatible with curved substrates and different substrate materials, and can be employed for depositing different metals.


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In future work, it will be interesting to exploit the favourable characteristics of the EP technique for preparing advanced transducer designs. For instance, microfluidic channels can be coated by simply flushing them with EP solutions,14,19,22 predestining this approach for lab-on-achip applications, while its conformity makes it compatible with fiber optics.23 EP also is capable of depositing gold23 and anisotropic NPs,19,35 which are valued in plasmonic applications due to their extraordinary chemical stability and altered optical properties,62 respectively, making the development and optimization of analogous synthesis routes towards spread-out Au NP films and complex NP morphologies worthwhile. ASSOCIATED CONTENT Supporting Information. Substrate variation; fabrication of gold nanoparticle films; variation and dynamics of the HCl treatment; continued silver particle growth; transducer fabrication using extended silver plating. AUTHOR INFORMATION Corresponding Author * Falk Muench: [email protected] * Alexander Vaskevich: [email protected] Present Addresses ┴

Technische Universität Darmstadt, Department of Materials and Earth Sciences, 64287

Darmstadt, Germany. Author Contributions


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The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. ‡ F. M. and A. S. contributed equally. Notes The authors declare no competing financial interest. ACKNOWLEDGMENT This work was supported by the German Research Foundation (F. M., Research Fellowship, MU 4125/1-1), the Minerva Foundation with funding from the German Ministry of Education and Research (A. V., I. R.), and by the Scholarship Program for Research Students from The Ministry of Aliyah and Integration of the State Israel (A. S.). F.M. thanks M. Diefenbach (Technische Universität Darmstadt) for providing access to the Cary spectrophotometer of the Biesalski group for additional UV-Vis measurements. ABBREVIATIONS EP, electroless plating; HB, hybridization buffer; LbL, layer-by-layer; LSPR, localized surface plasmon resonance; NP, nanoparticle; PAH, polyallylamine hydrochloride; PE, polyelectrolyte; PEM, polyelectrolyte multilayer; PSS, polystyrene sulfonate sodium salt; SEM, scanning electron microscopy; XPS, x-ray photoelectron spectroscopy.

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TOC GRAPHIC.


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Scheme of the silver NP film syntheses, including the two-step sensitization / activation procedure for introducing the silver seeds, which are optionally treated with HCl, and finally amplified via EP. The SEM images show the seeded and plated states of the pristine and ripened sample variants.



(A) UV-Vis spectra of glass substrates after sensitization, activation and ripening, corresponding to the synthesis steps 1–3 in Figure 1. (B) In situ UV-Vis spectra monitoring the silver deposition during EP. (C-E) SEM images of the silver NP films after plating times of (C) 6 min, (D) 10 min, and (E) 20 min.


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XPS spectra of the Sn 3d and Ag 3d binding energy regions of 0) pristine glass substrates, 1) after sensitization, 2) after activation, 3) after seed ripening, and 3B) after 2 min of electroless silver plating, including the estimated tin and silver content in atomic percent (at%) at each stage.



New figure 4 85x85mm (299 x 299 DPI)


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UV-Vis spectra of silver NP films obtained after (A) 2 min and (B) 6 min of electroless silver plating, recorded in the pristine state and during the sequential adsorption of PEM. Right to the spectra, SEM images of the NP films covered with 8 bilayers are shown.



Dependence of the shift of surface plasmon band on thickness of PEM for silver films plated for different time as in Figure 4. Solid lines present the fitting of the experimental data to the exponential decay of the optical response. 84x59mm (300 x 300 DPI)


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Scheme of the preparation and utilization of the biosensing interface. In the steps 1–3, a polyelectrolyte trilayer is deposited on the silver transducer surface, onto which ssDNA is adsorbed in step 4, and covalently attached by cross-linking in step 5. In the presence of the complementary DNA strand, hybridization occurs (step 6).



Transmission UV-Vis spectra of LSPR transducer prepared by silver plating for 6 min. taken after the following steps: (i) Adsorption PE trilayer (PAH-PSS-PAH (see Figure 6, step 3), (ii) Adsorption of ssDNA 155-Cy5_AmMO and sequential crosslinking (see Figure 6, step 5), (iii) exposed to either complementary (A) or unrelated (B) DNA strands.


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83x59mm (300 x 300 DPI)


Empowering Electroless Plating to Produce Silver Nanoparticle Films

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