Fabrication of
Plasmonic
Substrates and
Study of their
Performance for
Sensitive Detection
by Surface
Enhanced Raman
Spectroscopy
Cláudia Gomes Espinha
Medical physics
Department of Physics and Astronomy 2018
Supervisor
Joaquim Agostinho Gomes Moreira, Assistant Professor Faculty of Sciences of the University of Porto
Todas as correções determinadas pelo júri, e só essas, foram efetuadas.
O Presidente do Júri,
Acknowledgments
Firstly, I would like to thank my supervisor, Professor Joaquim Agostinho Moreira his guidance, enormous dedication and constant availability during all the dissertation. Also, my co-supervisor professor Eulalia Pereira for all the assistance during my research.
I am also very grateful to Miguel Peixoto for always being available to and answer my questions and for helping me to give my first sets in a chemical lab.
I thank Professor Pedro Jorge for providing the commercial substrate used in this work and to Professor Abílio Almeida for is help with the SERS equipment.
I am very grateful to the other members of "ClauGuDiAnAlex" for all these years of fellowship, to Mariana and Rafaela for all the support even from a far and to Norton for the friendship.
A special thanks to "Orfeão Universitário do Porto" for all the new experiences and new friends that gave me.
A profound thanks to my family for the constant affection, support and patience. Finally, I am especially thankful to my sistahh for all the "sister times" that help me to maintain the optimist during all these months of work.
Resumo
A necessidade de expansão de diagnósticos não-invasivos e da análise não-destrutiva de estruturas levou a grandes progressos na aplicação da espetroscopia de Raman aumentada pela superfície (Surface Enhanced Raman Spectroscopy, SERS) na área das ciências biomédicas. Usando SERS é possível melhorar o limite de deteção de bio-marcadores que permitem a deteção precoce de doenças. Este estudo tem como objetivo final a produção de substratos com características plasmónicas e desenvolver técnicas que um dia possam ser usadas com fins biomédicos.
Este trabalho teve dois objetivos: i) Análise e adaptação de diferentes técnicas de deposição de nanoestrelas de prata em vidro e em papel para substratos reprodutíveis e homogéneos. ii) Uso de SERS para analisar o estado reduzido de glutationa, medindo a proporção de uma sonda (5,5-dithio-bis-(2-nitrobenzoic acid), DTNB) em amostras.
Foram testados dois métodos diferentes de deposição, a deposição direta por gota e o método de sedimentação por centrifugação. Com o primeiro método o aumento foi de 102, para o segundo método o aumento foi da ordem dos 104. Além disso, um substrato comercial foi utilizado para comparação dos resultados obtidos com os nossos substratos. Com estes substrato funcionalizado com DTNB consegui-se um aumento do sinal de 104.
Na segunda parte deste trabalho, as diferenças dos espectros de SERS do DTNB e do TNB são promissoras. No entanto, são necessários mais estudos para que este processo possa ser aplicado a amostras clinicas.
Palavras chave: Nanoparticle Deposition, Surface Enhanced Raman Spectroscopy, Surface Plasmon Resonance, SERS biosensors
Abstrat
The growing need for noninvasive diagnostics and nondestructive structural analysis led to great progress in SERS medical applications. By using SERS one can improve the detection limit of biomark-ers that allow the detection of some diseases. This work has as final objective the production of plasmonic substrates that can be used in biomedical applications.
This study has 2 goals: 1) Analyze and adaptation of different techniques to deposit silver nanostars in glass and paper to form reproducible and homogeneous substrates; 2) Using SERS to analyze the redox state of the thiols measuring DTNB/TNB proportion in the samples.
Two different deposition methods were testes, direct drop deposition and centrifuge sedimentation method. In the first method an enhancement of 102 was achieved and for the second one the
en-hancement was of 104 order. Furthermore, a commercial substrate, with an enhancement of 104, was functionalized with DTNB and analyzed by SERS to serve as comparison.
On the second part of this work the SERS results of the DTNB and TNB are promising, because of the differences shown in both spectra. Nevertheless, more studies are still need before the process could be applied to clinical samples.
Key words: Nanoparticle Deposition, Surface Enhanced Raman Spectroscopy, Surface Plasmon Resonance, SERS biosensors
Contents
1 Introduction 11
2 Surface Enhanced Raman Spectroscopy (SERS): theory and applications 12
2.1 Theoretical background . . . 12
2.1.1 Raman scattering spectroscopy . . . 12
2.1.2 Surface Localized Plasmon Resonance (SLPR) . . . 16
2.1.3 Surface Enhanced Raman scattering . . . 19
2.2 SERS based biosensors for medicine . . . 22
2.2.1 Literature review . . . 22
2.3 Objectives . . . 27
3 Experimental Details 28 3.1 Synthesis of silver nanostars . . . 28
3.2 Morphological and Optical characterization of Ag nanostars . . . 28
3.3 Substrate preparation . . . 29
3.3.1 AgNSs substrates obtained by direct drop deposition method . . . 29
3.3.2 AgNSs deposition using the centrifuge sedimentation method . . . 31
3.4 Deposition of the SERS probe . . . 34
3.5 SERS instrumentation and result analysis . . . 35
4 Results and Discussion 37 4.1 Optical and Morphological characterization of the silver nanostars . . . 37
4.2 Raman and SERS spectra of DTNB . . . 40
4.3 SERS performance of the fabricated substrates . . . 42
4.3.1 AgNSs substrates direct drop deposition (DD) method . . . 42
4.3.2 Glass-based substrates . . . 42
4.3.3 Paper-based substrate . . . 48
4.3.4 AgNSs deposition using a centrifuge sedimentation (CD) method . . . 51
4.4 Commercial substrate . . . 56
4.5 Redox state study: . . . 59
4.5.1 SERS study of the DTNB reduction . . . 59
4.5.2 SERS study of DTNB reduction with GSH . . . 64
List of Figures
1 Schematic of the elastic and inelastic scattering processes and typical values. Where a is the typical dimension of the scatterin system; λ is the wavelength of the incident light; ν0 is the frequencie of the incident light; I0 is the intensity of the incident light;
Isis the intensity of the scattered light. . . 13
2 Energy levels and scheme of the elastic and inelastic light scattering. . . 15
3 Exemple of Raman spectrum. Spectrum of crocoite excited with the Ar+514.5nm [1]. . 15
4 Metallic sphere of radius a, placed in space with a relative electric permittivity rmand where there is a uniform electric field. . . 16
5 Module and phase of the complex polarizability of metallic nanoparticle as a function of the energy, from ref. [2]. . . 17
6 Induced fundamental plasmon. Two nanoparticles subject to an electric field. . . 18
7 a) Spherical Ag nanoparticle with the electric field distribution; b) Elliptical nanoparticle in resonance and the associated with a longitudinal electric field distribution; c) Elliptical nanoparticle in resonance and the associated with a transverse electric field distribution [3]. . . 19
8 Examples of variations of the local field intensity measure in: a) the surface of a glass sphere; b) the surface of a silver sphere; c) gap between two silver spheres. MLOCit’s the electric field enhancement factor. The electric field enhancement factor was calculated for a molecule in air at 1nm from the surface at point A [4]. . . 20
9 Examples of different types of hotspots; a) hotspot formed between two closely placed nanoparticles resulting from a coupled plasmon resonance; a) hotspot formed at a sharp geometrical feature. [5] . . . 22
10 Schematic representation of the sequence of steps followed to fabricate the biosensor. Glass substrates, gold nanostars, EpCAM aptamer, mercaptohexanol, EpCAM protein, and 4-aminothiophenol not drown to scale [6]. . . 23
11 SERS spectra collected on the substrates after exposure to increasing concentration of EpCAM [6]. . . 24
12 Cross section normalized diffusion with relation to the particle radius, for particles of silver and gold in media with different permittivities from ref [2]. . . 24
13 Scanning electron micrograph of silver film over nanosphere (AgFON), a widely used substrate for SERS. [7] . . . 25
14 a) Fabrication of SERS active substrates with gold nanostars using Electrostatically as-sisted APTES-Functionalized Surface-Assembly Method; b)SEM images of the resulting substrate [8]. . . 26
15 Schematic diagram of the direct deposition fabrication process, step by step. . . 29
16 a) Preparation of syringe and centrifuge tube; b) Experimental assembly: the syringe was placed inside the centrifuge tube. . . 31
17 Schematic diagram to illustrate the centrifuge deposition process [9]. . . 32
18 Reaction scheme of the DTNB reduction with GSH to form TNB. . . 35
19 Scheme of the T64000 system in subtractive mode. [10] . . . 36
20 Excitation spectra of the AgNSs: a) spectrum from the as-processed nanostars in this work; b) spectrum of AgNSs from Garcia et al. [11]. . . 37
21 Distribution of AgNSs size by NTA. a) Averaged nanostar concentration per hydrody-namic diameter; b) Intensity per hydrodyhydrody-namic diameter. The different color spots correspond to individual particles detected in 5 different videos by NTA software. . . . 38
22 SEM images of the a prepared substrate by the centrifuge deposition method; a) SEM image with a magnification of 2µm;b) SEM image with a magnification of 1µm. . . 39 23 SERS and Raman spectra of DTNB obtained by Akmaral [12]. The asterisk marks the
signal corresponding to the solvent used (methanol). . . 40 24 a) Raman spectrum of DTBN molecule deposited in glass substrate; b) Example of the
best fit of Eq. 15 to the experimental spectrum. The Residuals, green spots in b) are the difference between the experimental and simulated spectra. . . 41 25 a) Raman spectrum of DTBN deposited in paper substrate. (b) Example of the best
fit of Eq. 15 to the experimental spectrum. The Residuals, green spots in b) are the difference between the experimental and simulated spectra. . . 42 26 Results from the analysis of the sample G|DDep0.23: a) Normalized SERS spectra
relative to the acquisition time, in different zones of the sample; b) Microscopic picture of the sample in the Zone 4, zone with the larger density of the nanostars. The dark spots are nanostars, blue spot corresponds to the laser focus and the white are zones without nanostars. . . 43 27 Results from the analysis of the sample G|DDep2.52|rep2; a) Normalized SERS spectra
from several sample zones. b) Microscope picture of the substrate from zone 1; c) Microscope picture of the substrate from zone 3. The focus area is marked by a black dot in the picture, the darker areas correspond to nanostars and the white areas are zones without nanostars. . . 44 28 Results from the analysis of the sample G|DDep3*0.84|rep2; a) Normalized SERS
spec-tra from several sample zones. b) Microscope picture of the subsspec-trate from zone 1; c) Microscope picture of the substrate from zone 2. The focus area is marked by a black dot in the picture, the darker areas correspond to nanostars and the white areas are zones without nanostars. . . 46 29 Normalized SERS spectra of DTNB molecule, relative to the acquisition time, in
differ-ent zones of the samples. a) G|DDep2.52|rep1; b) G|DDep2.52|rep2. . . 47 30 Normalized SERS spectra of DTNB, relative to the acquisition time, deposited into two
different substrates and recorded in different zones. Rep 1 corresponds to the signals from sample G|DDep2.52|rep 1 and rep 2 corresponds to the signals from different zones of the sample G|DDep2.52|rep2. . . 48 31 Normalized Raman spectra, relative to the acquisition time in two different zones of a
paper sample, without nanostars. . . 49 32 Normalized SERS spectra, relative to the acquisition time, of different zones of a sample
with an AgNSs concentration of 0.46nM, P|DDep0.46. . . 50 33 Normalized SERS spectra, relative to the acquisition time, of different zones of a sample
with an AgNSs concentration of 1.84nM, P|DDep1.84. . . 51 34 Normalized SERS spectrum, relative to time of acquisition, of DTNB for two
differ-ent samples: a) Sample G|CDep0.2|rep2 with a base substrate of glass; b) Sample P|CDep0.2|rep2 with a base substrate of paper. . . 52 35 AgNSs substrates with increasing surface coverage. Picture of the samples: a) G|CDep1|rep2,
substrate fabricated without methanol and with an AgNSs suspension of 1nM; b) G|CDep0.7|rep2, fabricated with methanol and with a AgNSs suspension of 0.7; c) G|CDep1, fabricated with methanol and with a AgNSs suspension of 1; d) G|CDep1.4|rep1, fabricated with methanol and with a AgNSs suspension of 1.4. . . 52
36 Normalized Raman spectra relative to time of DTNB for two different samples with the same concentrations of nanoparticles deposited in a glass-based substrate: a) Substrate G|CDep1.4|rep1 ; b) Substrate G|CDep1.4|rep2. . . 53 37 SERS spectra of DTNB with AgNSs deposited by centrifuge sedimentation in paper.
AgNSs concentration of 1nM. . . 54 38 a) Raman spectrum of Hydroxylamine [13]; b) Raman spectrum of Sodium citrate [14]. 55 39 Normalized SERS spectrum, relative to time of acquisition, of DTNB (0.1M) deposited
in a substrate produce by centrifuge sedimentation deposition. . . 55 40 Normalized DTNB spectrum measured from the commercial substrate. DTNB
concen-tration of 0.02nM. a) Approximation and peak analysis. Raman shift ranging from 400 to 1800cm−1; Raman shift ranging from 400 to 3500cm−1. . . 57 41 Normalized DTNB SERS spectra from 4 different sample in comparison to the
commer-cial substrate. a) DTNB SERS signal of the sample G|DDep2.76; b) DTNB SERS signal of the sample P|DDep1.84; c)DTNB SERS spectra of the sample G|CDep1.40|rep1; d) DTNB SERS spectra of the sample P|Dep1. . . 58 42 UV-vis spectra of TNB (black) and DTNB (red). . . 60 43 Normalized Raman spectra of DTNB and TNB liquid solutions. a) Spectra from
400-1700cm−1; b)Spectra from 2500 to 3500 cm−1. . . 60 44 Normalized SERS spectra of DTNB and TNB, deposited in a silver substrate produced
by CD. . . 61 45 Schematic of the DTNB and TNB interaction with Ag nanoparticles [15]. . . 62 46 SERS spectra of DTNB:TNB proportions. The graph caption it’s related with the
amount of DTNB present in the sample, DTNB:Total solution. . . 63 47 Normalized SERS spectra of products of reaction of DTNB with GSH. . . 64
List of Tables
1 Chemicals used in this work. The putity and supplier are also shown. . . 28 2 Prepared glass substrates by direct deposition and the different fabrication condition:
nanostar suspension concentration, number of deposited droplets and DTNB concentra-tion. . . 30 3 Prepared paper substrates by direct deposition and the different fabrication conditions:
nanostar suspension concentration, number of deposit droplets and DTNB concentra-tion. . . 31 4 Glass substrates made by centrifuge sedimentation method and the different fabrication
conditions: AgNSs concentration, DTNB concentration, MeOH volume and centrifuge times. . . 33 5 Paper substrates made by centrifuge sedimentation method and the different fabrication
conditions: AgNSs concentration, DTNB concentration, MeOH volume and centrifuge times. . . 34 6 Vibration modes and frequencies for Raman and SERS spectra of DTNB [12]. . . 40 7 Characterists of the commercial substrate. . . 56 8 Enhancement values for each one of the substrates deposition techniques and comparison
of the enhancement performance to the commercial substrate of enhancement 1.6 × 104. 59
9 Vibration modes and SERS Raman shifts for DTNB, TNB and DTNB:TNB proportions. 63 10 Concentration of the reagents employed in DTNB/GSH reaction studied. The names of
the samples are related to the dilution factor of GSH, where GSH_0 corresponds to a sample without DTNB. . . 64 11 Vibration modes and SERS Raman shifts for the different GSH concentrations. . . 65
List of Abbreviations
.
SERS - Surface Enhanced Raman Spectroscopy AgNSs - Silver nanostars
NP - nanoparticles
LSPR – localized Surface Plasmon Resonance EF – enhancement factor
AEF – analytical enhancement factor UV – ultraviolet
ITO - Indium Tin Oxide
DTNB - 5’5- dithiobis-bis-2-nitrobenzoic acid TNB - 1,3,5-Trinitrobenzene
HA – hydroxylamin GSH – glutathione
NTA - nanoparticle tracking analysis SEM - Scanning Electron Microscopy Introduction
1
Introduction
Early diagnosis is the first step for a successful treatment, this leads to more likelihood of cure or longer survival [16]. So, early diagnosis techniques have attracted a lot of attention from scientists and there are already many techniques for early diagnosis of diseases. Periodic screening examinations for detecting specific chronic disease, for instance, helps to detect cancer, diabetes and cardiovascular diseases [16]. The aim of recent investigations is to find noninvasive and easier methods with low impact in the patient that allow early diagnosis and follow-up testing of diseases.
Detection and measurement of biomarkers is of vital importance in disease detection and recog-nition, management and monitoring of therapeutic efficiency. Early diagnosis and management of a disease relies heavily in the detection of low concentration of biomarkers, present in biological fluids, like blood and urine. These biomarkers appear in the human fluids in very low concentrations and that is why techniques that can detect these low concentrations of biomolecules are essential.
Plasmonic nanostructures have received great attention for the detection of analyte molecules in low concentration. Some metal structures, also known as plasmonic nanoparticles, have optical properties that allow for the enhancement of the electric field, with consequent amplification of the spectroscopic signal and an increase of the limit of detection. With the help of plasmonic nanoparticles, Surface Enhanced Raman Spectroscopy (SERS) can, detect the presence of even one single molecule of an analyte localized at the surface or at a very short distance from the surface of the metallic substrate [17].
As a result of its high sensitivity, due to the electromagnetic field enhancement by metal surfaces that increases the fingerprint signal of the biomolecules, SERS has been proven to be a powerful method in biomedical application [18]. To use SERS in the analysis of clinical samples, metal-based substrates need to be prepared. These substrates must be reproducible, homogenous and stable to allow reliable results. Furthermore, to allow for the current use of this technique in medical applications, the substrates should be low-cost and have an appropriate enhancement for the biomolecules to be detected. In this work, two different methods of nanoparticles deposition were studied in order to fabricate substrates that could be used for the analysis of clinical samples. These substrates were made with silver nanostars and tested with a probe molecule, 5,5-dithio-bis-(2-nitrobenzoic acid), DTNB. To test the efficiency of our substrates they were compared with a commercial substrate, in terms of quality of DTNB profile and enhancement. Lastly, the substrates with the best results were used to test a method to measure the thiol concentration. Glutathione was used along with DTNB for this tests.
This work is divided in 5 parts. Chapter 1 presents an introduction to the work, emphasizing the current importance of the subject; chapter 2 is a theoretical contextualization of the subject and a bibliographic revision of the state of art. In this chapter Raman spectroscopy, Surface localized plasmon resonance and the basis of surface enhanced Raman Spectroscopy (SERS) are discussed. Chapter 3 describes the experimental procedures. In chapter 4 the experimental results are presented and discussed. Finally, chapter 5 presents the conclusions of the work as well as some future perspectives.
2
Surface Enhanced Raman Spectroscopy (SERS): theory and
applications
2.1
Theoretical background
Surface Enhanced Raman Spectroscopy (SERS) is a powerful vibrational spectroscopy technique that allows for highly sensitive structural detection of low concentration analytes, through the amplification of electromagnetic fields generated by the excitation of localized surface plasmons [17]. The main ingredients of SERS are Raman spectroscopy and plasmonic resonances; mixing them together makes SERS possible.
2.1.1 Raman scattering spectroscopy
When light impinges a material, several processes occur, among which radiation scattering. From now on, we will consider the scattering processes of light with wavelength much larger than the typical dimensions of the scattering system (λ ∼ 400 − 800nm). This enables to simplify the formal approach of the scattering process. The light is scattered by two main mechanisms depending the relation between the energy of the incident and scattered photons:
• Elastic scattering, where the incident and scattered photons have the same energy. This process is often referred to as Rayleigh scattering.
• Inelastic scattering, where the scattered photon has a different energy than the one of the incident photon. One of the most important forms of inelastic scattering is the Raman scattering by optical vibrations of the system.
From know on, we shall consider just non-resonant Raman scattering, when the frequency of the incident light is small enough to prevet diatomic transitions.
Figure 1 schematically summarizes the main features of the two types of scattering processes here discussed.
Figure 1: Schematic of the elastic and inelastic scattering processes and typical values. Where a is the typical dimension of the scatterin system; λ is the wavelength of the incident light; ν0is the frequencie
of the incident light; I0 is the intensity of the incident light; Isis the intensity of the scattered light.
Rayleigh scattering is the predominant process of light scattering by particles or molecules with dimensions much smaller than the wavelength of the radiation. As the energy of the both incident and scattered photons are equal, the Rayleigh scattering does not change the state of the material. The intensity of the Rayleigh scattering is typically 10−3 of the incident intensity.
In the inelastic light scattering process, the initial and final state of the scattering system differ from each other. This is due to the loss/gain of energy of the incident/scattered photon. Light scattering can be view as a re-radiating process by electric dipoles by the incident electric field. In this sense, Raman scattering appears as a modulation of the linear optical polarizability due to the presence of lattice or molecular vibrations. The intensity of the Raman scattering light is about 10−6 to 10−9 of the incident intensity, thus being a very weak process.
The Raman scattering can be easily understood in the framework of a classical model. The classical model considers the atomic system as a collection of independent oscillators, described by normal modes. The dimensions of the scattering molecules are very small compared with the wavelength of the incident light. The wavelength of the radiation commonly used in Raman scattering experiments is of the order of 103 to 105 nm. This value is several orders of magnitudes larger than the typical lengths
and distances between atoms and molecules of the scattering medium. In this approximation, the electric field of the incoming light can be considered uniform over the space occupied by the molecule and it is only necessary to consider its temporal dependence. According to these assumptions, the electric field of the incident light can be described, in the region occupied by the scattering molecule, by: ~ Einc(t) = −→ E0· e−i(w0t− − → k·−→r) + c.c (1)
where |−→k | = 2πλ is the propagation vector modulus, w0 = kc is the angular frequency of the
electro-magnetic wave, and −E→0 is the complex amplitude vector which defines the polarization state of the
−→
E0has complex components, but without loss of generality, it is possible to disregard the imaginary
components assuming a linear wave polarization. When the electric field of the incident wave interacts with the sample, it induces an electric dipole, whose components, in a linear approach, are given by:
pi=
X
j
αijEj(r, t) , i, j = 1, 2, 3 (2)
In Eq. (2) αij are the components of the electrical polarizability tensor and i,j are the cartesian
coordinates. Considering that the normal vibrations have small amplitudes, it is possible to expand the elements of the polarizability tensor in a series of powers of normal modes, Qk:
αij= α0ij+ X k ∂αij ∂Qk 0 Qk+ 1 2 X k,1 ∂2αij ∂Qk∂Q1 0 QkQ1+ ... (3) Here, α0
ij is the element of the polarizability tensor calculated in the equilibrium configuration; Qk
is the kthnormal vibration mode of the system; all the derivations are calculated in the equilibrium
configurations [10].
In the harmonic lattice or harmonic vibration approach, each normal mode can be express as follows:
Qk(t) = Q0ke−i(Ωkt+Φk), (4)
where Q0k is the amplitude of the kthnormal mode, Ωk is the angular frequency and Φk is the phase
of the vibration. So, the i component of the induced dipolar moment, in a first order approximation, is given by: pi(t) = X j α0ijEj· e−iw0t+ X k ∂αij ∂Qk 0 EjQk0· e−i[(w0+Ωk)t+φk]+ e−i[(w0−Ωk)t+φk]+ c.c (5)
The first term of Eq. (5) corresponds to the Rayleigh scattering, as the induced electric dipole oscillates at the same frequency of the incident electric field. The second term corresponds to the inelastic scattering. The induced dipolar moments oscillate at frequencies w0± Ωk. These frequencies
are the frequencies of the Stokes (w0− Ωk) and the Anti-stokes (w0+ Ωk), spectral components due
to the Raman scattering.
Although the classical model can explain the existence of Raman scattering, it cannot describe the intensity relation between the Stokes and the anti-Stokes bands. The quantum theory approach of Raman scattering considers lattice vibrations as quasi-particles - phonons - interacting with photons of the incoming beam. The intensity of the Raman bands is related with the population in each phonon energy level in thermal equilibrium. Assuming that the probability of a phonon level being occupied is given by the Boltzmann distribution (e−kTU ), the low energy levels are the most occupied in
thermal equilibrium. Figure 2 shows a diagram to explain the relation between Stokes and anti-Stokes intensities:
• Stokes radiation happens if the scattered photon has less energy than the incident photon, then the molecule is excited to a higher-energy vibration level, and the scattered photon frequency is w0− Ωk.
• Anti-Stokes radiation happens if the scattered photon has more energy than the incident photon. The molecule has relaxed from an excited vibrational state. The frequency of the anti-Stokes radiation is w0+ Ωk.
Figure 2: Energy levels and scheme of the elastic and inelastic light scattering.
The difference w ± Ωk is called Raman-shift and it is commonly expressed in wave number (cm−1).
A vibrational mode gives origin to two bands, symmetrically located relatively to the excitation wave number/frequency w0. The Raman shifts are measured relatively to the excitation line, it is usual
to assume the origin of the wave number as the excitation wave number. In the case of the thermal equilibrium at temperature T, the intensity of the Stokes bands is higher than the anti-Stoke ones. This is why just the Stokes range of the scattering spectra is usually studied. The main features of a typical Raman spectrum are depicted in figure 3. As can be seen, at room temperature, the anti-stokes scattering intensity is weaker than the Stokes, due to the population difference between the ground state and excitation levels.
The Raman shift of a peak is equal to the vibrational frequency of the corresponding mode. There-fore, it can be said that the Raman spectroscopy provides a fingerprint of the molecular vibrations. As the intensity of a Raman band is proportional to the population of the vibrational level, the study of a particular Raman line intensity provide information regarding the concentration of a molecule in a sample and a quantitative analysis can be performed.
Summarizing, Raman spectroscopy induces inelastic scattering of the light by molecules, with the scattered light providing a unique vibrational fingerprint of each molecule [19].
The main limitations of the Raman spectroscopy are related to molecules with very low Raman efficiency and very low concentrations of the probe molecule. All these factors make the intensity of the Raman signal very low and contribute to the need to enhance the signal for medical applications. 2.1.2 Surface Localized Plasmon Resonance (SLPR)
Metals with SERS effect appear in the form of metal nanostructures that enhance the Raman signal through plasmonic resonance. Surface plasmons are collective oscillations of conduction electron gas excited by the electromagnetic field of light. In the case of metal nanoparticles, the electron oscillations induce an electric field around the nanoparticles (NP) that can be much larger than the one from the incident light [20].
In the following, a simple classical approach of the free electron contribution to the electric polar-izability of the nanoparticle is presented. In this approach, the size of the metal particle is very small compared with the wavelength of the electromagnetic field (a << λ). In this case, we can neglect the spatial deformation of the electric field over the particle and assume it as uniform. This is the so-called quasi-static approach. Considering this, the problem becomes the case of a spherical particle in a uniform electric field,−E→0= E0−u→zsurrounded by a homogenous medium with electric permittivity
rm (see figure 4).
Figure 4: Metallic sphere of radius a, placed in space with a relative electric permittivity rm and
where there is a uniform electric field.
Now, we assume that the relative electric permittivity of the dielectric medium rmis a real quantity
and frequency independent. The relative electric permittivity (r(w)) of the metal particle is described
by the Drude model [2]. In this case, the electrical potential as a function of position is: φin(−→r ) = −
3rm
r+ 2rm
E0r cos θ (6)
φout(−→r ) = −E0r cos θ +
− →p · −→r 4π0rmr3
where 0 is the electric permittivity of the free space; and −→p is the induced dipole moment in the
sphere, given by:
− →p = 4π 0a3 r− rm r+ 2rm −→ E0= 0rmα −→ E0 (8)
α is the electric polarizability of the sphere in the quasi-static approach: α = 4πa3r− rm
r+ rm
(9) The polarizability depends of the nanoparticle size, a, and of the relative permittivity of the sur-rounding medium. Figure 5 shows the module and argument of complex polarizability as a function of the energy.
Figure 5: Module and phase of the complex polarizability of metallic nanoparticle as a function of the energy, from ref. [2].
From Eq. (9), a maximum of the magnitude of the electric polarizability can be found at a frequency for which the following condition holds: |r+ 2rm| is minimum. If the frequency of the
electromagnetic wave is larger than the plasma frequency of the metal, we can discard the imaginary part of the polarizability. In this condition, resonance occurs at the frequency for which:
Re [r] = −2rm (10)
This relation is known as the Frohlich condition. For a Drude metal, this criterion is satisfied at the frequency w:
2rm= −1 +
w2 p
w2 (11)
So, the resonant frequency depends on the relative electric permittivity of the surrounding medium. If the dielectric medium is air (rm= 1), then:
w = √wp
3 (12)
Suspensions of metal nanoparticles are, therefore, ideal for the manufacture of optical sensors for refractive index changes [2]. Metals, such as gold (Au), silver (Ag) and copper (Cu), have interesting optical properties in the visible spectrum due to the contribution of the free electrons to the electric
permittivity, the complex electric permittivity of the metals is an important feature of the plasmon behavior [17].
A polarizability enhancement at resonance conditions implies an enhancement in the electric field. This is because the field in the surroundings of the particle has a dipolar origin. So, when the polariz-ability is in resonance conditions, the field for an electric dipole is maximum. In the quasi-static field calculation an ideal dipole was consider. Nevertheless, the solution is valid for time dependent fields, discarding the delay on the particle. For a plane and monochromatic wave, the electric field induces an oscillation in the electric dipole, which in a linear approximation, is:
− →p =
0rmα
−→
E0e−iwt (13)
In a region near the particle, the metal nanoparticle acts like an optical antenna, converting the incident electrical energy into a strongly confined field in the neighborhood of the particle. This is possible because of the excitation of the localized plasmon. Figure 6, shows the induced fundamental plasmon in two non-interacting spherical nanoparticles subject to an electric field and the consequent charge distribution inside the nanoparticles. If the frequency of the electric field of the electromagnetic wave is in resonance condition with one plasmonic frequency of the nanoparticles, the polarizability will increase, also increasing the electric dipole and the electric field. The electric field is enhanced by localized and non-interacting plasmonic modes. The frequency calculated from Eq.11 corresponds to the lowest frequency plasmon. Other plasmon can be also excited, but the calculation of their frequencies are beyond the scope of this work.
Figure 6: Induced fundamental plasmon. Two nanoparticles subject to an electric field. The Localized Surface Plasmon Resonance (LSPR) depends on the size, shape, interparticle spacing and dielectric properties of the metal. For non-spherical nanoparticles, the polarizability is anisotropic. As shown in figure 7, changes in the shape of the nanoparticles alters the field distribution near the nanoparticles and consequently the resonance positions:
• A decrease in the nanoparticle radius of curvature will induce an increase of surface density charge and consequently a more confined field ("hotspots").
• Stretched nanoparticles will present strong anisotropic optical answer, with a strong dependence of the incident field direction, as shown in figure 7 (b) and (c).
Figure 7: a) Spherical Ag nanoparticle with the electric field distribution; b) Elliptical nanoparticle in resonance and the associated with a longitudinal electric field distribution; c) Elliptical nanoparticle in resonance and the associated with a transverse electric field distribution [3].
As the distance between nanoparticles decrease, interactions between plasmonic modes becomes more intense, and couple plasmonic modes must be considered. The interaction between plasmons also creates changes in resonance frequency. If the distance between the particles, d λ, decreases a strong field confinement can be observed between the adjacent particles, in unidimensional chains. This confinement of the field in the space between the particles is of great interest, especially in SERS. The LSPR extinction maximum (λmax), is the resonance maximum of the nanoparticle arrays
and can be measured with UV-visible extinction spectroscopy. For silver and gold nanoparticles, the resonance conditions are met with visible light, making these materials suitable for numerous applications and of special interest in SERS. Since the LSPR of the nanoparticles is highly dependent on the local environment, the extinction maximum will shift when the adsorbates bind to the nanoparticles [21].
2.1.3 Surface Enhanced Raman scattering
Surface Enhanced Raman Spectroscopy originates from light scattering by the vibrations of molecules that are in close proximity to nanostructured metal surfaces that are capable of supporting plasmon resonances in the spectral region where Raman scattering is excited [22]. The SERS intensity for a given vibrational mode of a given analyte should also be proportional to the laser intensity and to the normal Raman cross-section but it is also affected by an enhancement factor (EF) [4].
SERS enhancement are traditionally separated into two main multiplicative contributions [4]: • Chemical enhancement (CE) factor: the chemical mechanism contributes to enhancement through
chemisorption of the molecule to the noble metal surface, allowing the electrons from the molecule to interact with the electrons from the metal surface. These interactions lead to an enhancement of signal up to 102.
• The electromagnetic enhancement (EM) factor: the electromagnetic enhancement is a wavelength-dependent effect arising from the excitation of the localized surface plasmon resonance. This collective oscillation of conduction electrons can occur in noble metal nanoparticles (NPs), sharp
metal tips, or roughened metal surfaces, and enhances the incident electric field intensity 104 times in the vicinity of the metal surface.
Due to the interaction between the molecules and the metallic particle, alteration of the Raman activity are observed [4]:
• Alterations in the electromagnetic field at the molecule position due to presence of metallic objects, this results in a possible local field enhancement.
• Variations in the radiation properties of the Raman dipole pi, that results in a possible radiation
enhancement.
• Alterations in the Raman polarizability tensor may be modified. Such modification would typi-cally be classified as the chemical enhancement.
As it was referred to above, the electromagnetic field is strongly modified in the vicinity of the metallic objects, especially when the frequency is close to the plasmonic resonances of the system. These alterations can be both in magnitude and in orientation, and are associated with the localized surface plasmon resonances created by the metal surfaces. Figure 8 shows examples of modifications of the local field intensity measured by the field enhancement factor MLOC, in the presence of metal nanoparticles
and the wavelength dependence. MLOCis the ratio between local electric field and the incident electric
field [4].
Figure 8: Examples of variations of the local field intensity measure in: a) the surface of a glass sphere; b) the surface of a silver sphere; c) gap between two silver spheres. MLOC it’s the electric field
enhancement factor. The electric field enhancement factor was calculated for a molecule in air at 1nm from the surface at point A [4].
For the glass sphere (figure 8a), there is a small enhancement of the field, hardly dependent on the wavelength. For a silver sphere (see figure 8(b)) the electric field enhancement factor shows maxima values for specific values of the wave length, assigned to resonances of the localized surface plasmon of the silver sphere (λ = 360nm). Such effects are even more striking in the case of a molecule localized between two silver spheres separated by a 2nm gap, presented in figure 8(c). Figure 8(c) shows the effect of the interacting plasmons in two Ag spheres. In this case, more than one resonance are observed, assigned to different coupled mode of the system.
In SERS, the radiation properties of the Raman scattering process are modified. This phenomenon is known as modified spontaneous emissions. The environment alters the radiation pattern and the total power radiated by the dipole. These effects will depend on several factors including the substrate geometry and optical properties, the dipole position, orientation, and its emission frequency.
In addition to the physical changes between Raman and SERS, the spectra of the samples will also suffer alterations. Most molecules exhibit a SERS spectrum that is very similar to their normal Raman spectrum (at the same excitation wavelength), and most of the fingerprint Raman peaks are easily identifiable. However, some minor differences may arise. The Raman spectrum under SERS conditions can be affected because the plasmon resonances, producing the enhancement, are typically wavelength dependent. As a consequence, different parts of the spectrum can be amplified differently, depending on the dispersion of the underlying resonance producing the enhancement. Furthermore, the molecule may change its "identity" upon adsorption and become a surface complex. This may result in small shifts and/or broadening of Raman peaks [4].
The intensity of the SERS signal is compared with the intensity of the Raman signal by the analytical enhancement factor (AEF). The analytical enhancement factor is obtained by comparison of the SERS spectrum with what would be the Raman spectrum under non-SERS conditions for the same molecule [4]:
AEF =ISERS/cSERS IRS/cRS
, (14)
where ISERS and cSERs are the SERS signal and concentration respectively and IRsand cRS are the
same measurements in a Raman spectrum [4]. SERS enhancement factors ranging from 106 to 108
have been observed with several substrates [7].
There are some reports of enhancements in the order of 1015 , due to the existence of hotspots
in a sample [5]. Hotspots are regions of intense field enhancement due to LSPR. There can be two types of hotspots: those formed between two closely spaced nanoparticles and those formed on sharp features of the nanoparticles [12]. The first ones, shown in figure 9(a), benefit from an aggregation of the metal colloidal suspension, since a huge intensification of the field can be induced in interparticle gaps or on the surface of large aggregates. Nevertheless, aggregation decreases the reproducibility of the substrates. To achieve such high amplifications, the molecules that are under study must be adsorbed to nanoparticles. The hotspots due to the sharpness of the nanoparticle, presented in figure 9(b), happen because the restoring force for SP is related to the charge accumulated at the particles surfaces, and there will be a bigger charge accumulation in the sharp edges of the nanoparticles and consequently a higher enhancement in these regions [20].
Figure 9: Examples of different types of hotspots; a) hotspot formed between two closely placed nanoparticles resulting from a coupled plasmon resonance; a) hotspot formed at a sharp geometrical feature. [5]
The SERS process depends on a long list of parameters, including characteristics of the laser excitation (wavelength, polarization, angle of incidence...), detection setup, intrinsic properties of the angle (Raman polarization tensors of the modes), analyte adsorptions properties and morphological features of the substrate.
In conclusion, good SERS substrates are, in simple terms, those that support the "strongest" plasmon resonances, in other words, those that provide the largest enhancement or amplification. In this regard, one should in addition distinguish between those that provide a relatively uniform enhancement on the surface and those with large variation. Moreover, because the SERS enhancements arise from a resonant response of the substrate, they are typically strongly wavelength dependent. A given SERS substrate will, therefore, typically exhibit good enhancements in a limited excitation wavelength range. Most SERS substrates are designed to operate with visible/near-infrared excitation (∼400-1000nm), which is the typical range of interest for molecular Raman scattering experiments. Furthermore, to have a good SERS signal, the molecule must have two important characteristics: intrinsic Raman properties and strong molecule/metal interaction.
2.2
SERS based biosensors for medicine
2.2.1 Literature review
In recent years, SERS based biosensors have received much attention, especially for the detection and quantification of biomarkers of diseases. These biosensors can be applied in the medical field, in diagnostic and monitoring of therapeutic efficiency. For example, a sensor for monitoring glucose levels was developed [17]. In cancer detection, Bhamidipati et al. [6] developed a new biosensor for early detection of cancer cells. Other studies have developed the use of thiols as biomarkers. Thiols are good biosensors, because there is a link between the thiol concentration and the status of proliferative and degenerative diseases[23]. Banne et al. showed that thiols can be used to determine individual diseases status by comparing the levels of thiols in healthy subjects with an active disease holder [24]. Some reliable methods of measuring thiols using UV-Vis spectroscopy already exist, but are quite laborious and have a high limit of detection of thiols compounds. So, in recent years, SERS based sensors have also been studied with the purpose of measuring the concentration of thiols in the human blood [24]. SERS has several advantages in the detection of biomarkers namely a strong enhancement of the Raman signal of the molecules [18]. This strong enhancement is particularly interesting for biomarkers that are present at low concentration in biological fluids like blood or urine. Other features to be
considered in SERS-biosensor are reproducibility, specificity, sensibility, stability and, in order to facilitate the access during the progression of the disease, it must be inexpensive [25]. Nowadays, an intensive research on new types of SERS-based biosensor is enabling the improvement of those important attributes.
In cancer detection, Bhamidipati et al. [6] developed a new biosensor to detect the "cancer biomarker epithelial cell adhesion molecule" (EpCAM) and to quantify its expression at the single cell level. EpCAM is one of the most commonly expressed markers on a variety of cancer cells [6]. It has been suggested that the reduction of its expression levels could be associated to the epithelial to mesenchymal transition (EMT), and thus to the onset of metastasis. Therefore, monitoring vari-ations in expression levels of this membrane biomarker would improve the ability to monitor cancer progression.
Figure 10: Schematic representation of the sequence of steps followed to fabricate the biosen-sor. Glass substrates, gold nanostars, EpCAM aptamer, mercaptohexanol, EpCAM protein, and 4-aminothiophenol not drown to scale [6].
Figure 10 shows the fabrication scheme of the biosensor used in cancer detection. [6] After silaniza-tion, the glass slides were incubated with a 3nM solution of gold nanostars, synthesized according to a modified version of surfactant-free nanostar synthesis. This concentration ensures a uniform distribu-tion of the nanoparticles on the glass surface, without substantial clustering. The second step of the substrate preparation is the in situ functionalization with the suitable aptamer, the 48 bp SYL3C. The aptamer sequence is designed to possess a thiol group, to bind to the surface of the gold nanostars. In order to optimize aptamer-protein recognition, while avoiding non-specific binding, the substrate is backfilled with 6-mercaptohexanol (MCH). The substrates are later incubated with varying con-centration of EpCAM protein that is prepared in the binding buffer specific for this aptamer-target pair. The final step in the substrate preparation is the addition of SERS tags to identify and localize aptamer protein binding events. These substrates are loaded with cancer cells of different types and their limit of detection was determined. Figure 11 show a SERS spectra for increasing concentration of the EpCAM biomarker.
Figure 11: SERS spectra collected on the substrates after exposure to increasing concentration of EpCAM [6].
From the analyses of the spectra of the figure 11, it was possible to conclude that there was an intensity variation of the 1076cm−1 peak with EpCAM concentration. This change was used in additional studies to measure, with efficiency, the number of cancer cells deposited on the substrate. It was possible to detect EpCAM within limits of detection of 10pM. [6]
SERS-based sensors can have many forms and they can be produced using many processes, ac-cording to the purpose. Usually, the SERS-based sensor uses solid substrates. In this case, the first parameter to consider is the choice of enhancing substrates. The substrates can vary in material, shape and coating. The most used materials are silver nanoparticles (AgNPs) and gold nanoparticles (AuNPs), because they are air stable materials and show LSPR that covers most of the visible and near infrared wavelength range [17]. Among them, silver nanoparticles are expected to have a higher enhancement factor at UV-vis wavelength because of the dielectric functions of silver are very high, as can be seen in figure 12.
Figure 12: Cross section normalized diffusion with relation to the particle radius, for particles of silver and gold in media with different permittivities from ref [2].
Furthermore, highly faceted particles are a better choice for SERS, because of the strategic presence of edges and tips that can concentrate the electromagnetic field onto specific regions of the nanoparticle surface [26]. Hereafter, we will call substrate to the physical support covered with nanoparticles.
The deposition technique of the nanoparticles is another important factor to consider in the fab-rication of SERS-based sensors. Many experimental techniques can be used to fabricate SERS-based substrates. Among them, Pulse laser Deposition (PLD) has been intensively used [27, 28]. Other technique is alumina-modified AgFON substrates fabricated by atomic layer deposition as described by Zhang et. al. [29].
Figure 13: Scanning electron micrograph of silver film over nanosphere (AgFON), a widely used substrate for SERS. [7]
Figure 13 shows a scanning electron micrograph of AgFON. Xiaoyu Zhang demonstrated that AgFON substrates show temporal stability exceeding 9 months, and they can be prepared by coating Ag surfaces with a sub-1mn alumina overlayer [7]. In the fabrication process, 2µL of the nanosphere suspension was drop-casted into glass and allowed to dry in ambient conditions. The metal films were deposited by a vapor deposition system with a pressure of 10−6Torr. Alumina films were fabricated on the AgFON substrates by atomic layer deposition [29]. The coating of the substrates serves to stabilize the Ag surfaces of the AgFON substrates, which allows for a higher chemical stability, that is major concern for Ag-based substrates. Although this substrate has all the essential characteristics of a good SERS substrate, the use of expensive deposition techniques and machines during fabrication, makes it little recommended for medical applications. For medical use, it is necessary to find a simpler and inexpensive technique of nanoparticle deposition that will allow for disposable use of the substrates throughout the treatment process, without losing properties required to a biosensor.
Some less expensive fabrication techniques have already been studied. Most of these techniques use a colloidal nanostar suspension in the deposition. The major disadvantage in this case is the possible occurrence of aggregation that lowers the reproducibility of the substrate. Nevertheless, to increase the SERS performance of these substrates, an aggregation of the colloidal suspensions must be carried out, since a huge intensification of the field can be induced in the interparticle gaps or on the surface of large aggregates.[11] Qianqian Su et al. [8], describes a SERS substrate, using an ITO (Indium Tin Oxide) glass base, fabricated by electrostatically assisted APTES-functionalized ((3-Aminopropyl)triethoxysilane) surface assembly of gold nanostars. This method was an attempt to provide a solution to the fabrication of low cost, large scale and reproducible SERS active substrates.
Figure 14: a) Fabrication of SERS active substrates with gold nanostars using Electrostatically assisted APTES-Functionalized Surface-Assembly Method; b)SEM images of the resulting substrate [8].
Figure 14 shows the process of production of the SERS active substrates with gold nanostars using electrostatically assisted APTES-functionalized surface-assembly. The glass slides were cleaned with a mixture of water and ethanol and then were immersed in ethanol for two hours. After drying, the glass slides were vertically dipped overnight into a stirred colloidal suspension of gold nanostars for the purpose of preparing Au nanostar layers. The homogeneity of the SERS signal and the large area of the resulting substrates is the greatest advantage of this method (up to several square centimeters). Furthermore, for probe substances like Nile blue (NBA) and Rhodamine 6G (R6G), the detection limits are 5×10−11M and 1 × 10−9M respectively, when using a 785nm excitation source. The estimated AEF is 5 × 10−6for NBA and 2 × 105 for R6G.
Polacarapu el al. [30] highlighted other important factor, the base of the substrates. Materials like paper, free-standing nanofibers, elastomers, plastics, carbon nanotubes and graphene should be used to produce low-cost and flexible nanoplasmonic devices. Flexible substrates display several advantages. They can be used in packaging (can be wrapped around underlying non-planar substrates) or used as swabs to collect samples. Moreover, the sensing methods that can be applied with such substrates are non-invasive and the majority of these materials are inexpensive. Paper has been widely used as a flexible supporting material because it is one of the cheaper materials and also is ecofriendly and biodegradable. The techniques of deposition reported are very different. Vo-Dinh et al.[31] used a two-step method, where polystyrene latex particles were first spin-coated on the surface of filter paper, followed by thermal evaporation of the silver nanoparticles. There are also reports of easier and less expensive methods like printing technology. This technique can also be applied to fabricate flexible plasmonic devices by printing nanoparticle inks on paper substrates. Another simple solution process is the “dip-coating” of metal nanoparticles on paper substrates [30]. The disadvantage of paper is the presence of cellulose which has a Raman signal, unlike glass. This can sometimes change the Raman spectrum of the sample being tested.
“SERS has progressed from studies of model system on roughened electrodes to highly sophisticated studies, such as single molecules spectroscopy” [17]. Its many applications, not only in the medicine field, but in other areas like geography or agriculture will continue to be study, what makes SERS a pertinent area of studied in these days.
2.3
Objectives
The objective of this work is to develop an easy and inexpensive way to produce substrates that allow a strong enhancement in SERS analysis. The substrate must be reproducible and homogeneous in order to obtain consistent measurements independently of the measurement point or the substrate. Furthermore, the enhancement achieved with this substrates must be high enough to allow the SERS analysis of analytes with very low concentrations, because biomarkers appear in very low concentrations in human fluids.
In conclusion, the aim of this study is: i) produce SERS substrates that can, in future works, be used and further developed for biomedical field applications like diagnostics and monitoring diseases; ii) use the produced substrates to test a method of measuring the redox state of clinical samples. The aim is that in the future this procedure, based in SERS for the analysis of biomolecules, can be adapted to disposable devices and replace the current methods that use UV-vis spectrometry. This would allow an increase in the detection limit of the analyte.
3
Experimental Details
In this chapter we will present a description of the experimental techniques used in this work to prepare the substrates and characterize their SERS performance.
In the next table all the chemicals used in this work, as well the corresponding purity and the supplier, are presented.
Table 1: Chemicals used in this work. The putity and supplier are also shown.
Chemicals Supplier Purity % silver nitrate Aldrich 99.999% hydroxylamine (HA) Aldrich 50wt. % solution
sodium hydroxide Fisher 98.7% tri-sodium citrate dihydrate Panreac PA-ACS
methanol Sigma-Aldrich ≥99.9% 5’5- dithiobis-bis-2-nitrobenzoic acid (DTNB) AcrosOrganics 99%
glutathione (reduced, GSH) AcrosOrganics 98% sodium borohydride Fulka ≥ 99%
hydrogen peroxide Fisher >30% w/v
3.1
Synthesis of silver nanostars
The synthesis of silver nanostars (AgNSs) was performed as described by Garcia-Leis et al [11] with modifications proposed by Suleimenova, A.[12]. Colloidal suspensions of AgNSs were prepared by chemical reduction of Ag+ in a two-step process: i) formation of silver spherical nuclei by reduction
with neutral hydroxylamine (HA); ii) formation of the star tips by further reduction and capping with citrate.
AgNSs were prepared as follows: (i) 2.5mL of 6 × 10−2M solution of HA was mixed with 2.5 mL of 5 × 10−2M of sodium hydroxide at moderate stirring (900 rpm), at room temperature; (ii) then, 44.5 mL of silver nitrate solution was injected using a syringe pump (flow 44 mL/min), under stirring; (iii) the solution became dark grey and after 90 seconds 500 mL of 1.5% (m/V) trisodium citrate was added to the mixture still being stirred; (iv) after 3 hours, the suspensions became grey and were centrifuged for twelve minutes at 1600g. At the end, the supernatant was discarded and the AgNSs suspension was stored in the dark for further studies. The suspensions obtained were used directly in the preparations of the SERS substrates.
All the glass ware used were previously cleaned with Aqua-Regia1 and Milli-Q water.
3.2
Morphological and Optical characterization of Ag nanostars
The concentration of AgNSs in solution was measured by nanoparticle tracking analysis (NTA) using a Malvern Panalytical Nanosight NS300 with a dilution of 1:100 or 1:200 with Milli-Q water, depending of the initial concentration of the solution.
The surface morphology of the samples was checked using a Scanning Electron Microscopy (SEM) Schottky - FEG-ESEM/EDS/EBSD FEI Quanta 400 FEG ESEM/EDAX Genesis X4M - with 15.00 kV in secondary electrons mode.
1Aqua-regia is extremely corrosive and it should be handled with extreme care in the fumehood and using appropriate
The optical characterization was made using UV-vis spectroscopy. The extinction spectra of the nanostars was obtained using a Varian Cary 50 Bio UV-Vis spectrophotometer in the wavelength range 250 nm to 800nm. Nanostars suspensions were diluted 1:30 with Milli-Q.
3.3
Substrate preparation
In this work, we have studied the SERS performance of two types of substrates, based on glass and paper, respectively. In each case, the as-prepared nanostars were deposited using the direct drop deposition or centrifuge deposition techniques, which will be described in the following sections. 3.3.1 AgNSs substrates obtained by direct drop deposition method
In this section, it will be firstly described the procedure used for glass-based substrates, followed by the necessary changes for a paper-based substrates.
Glass slides (26×76mm, Labbox) were firstly washed with Milli-Q water and then with methanol (MeOH). A sticky plastic band (d-c-fix) with 5 circular holes, with a 5.5mm diameter, was used to completely cover each glass slide (see figure 15). This allows to have the same area for all the nanostars deposition and prevents the mixing of the different zones. A 10 µL volume drop of AgNSs was deposited in each one of the holes and left to dry over 12h. The substrates were then stored (in the dark) for later use. Different substrates have been prepared, with variations of the AgNSs concentration and volume. Table 2 presents the data concerning the substrate, concentration of the AgNSs in suspension, and volume deposited. When more than one deposition was made, the interval between depositions was 30 minutes.
Table 2: Prepared glass substrates by direct deposition and the different fabrication condition: nanostar suspension concentration, number of deposited droplets and DTNB concentration.
Sample AgNSs concentration (nM) DTNB concetration (mM) Number of DTNB depositions G|DDep0.23 0.23 10 1 G|DDep0.12 0.12 10 1 G|DDep0.06 0.06 10 1 G|DDep0.04 0.04 10 1 G|DDep0.03 0.03 10 1 G|DDep0.02 0.02 10 1 G|DDep0.46 0.46 10 1 G|DDep0.92 0.92 10 1 G|DDep1.38 1.38 10 1 G|DDep1.84 1.84 10 1 G|DDep2.30 2.30 10 1 G|DDep2.76 2.76 10 1 G|DDep0.84|rep1 0.84 10 1 G|DDep0.84|rep2 0.84 10 1 G|DDep0.84|rep3 0.84 10 1 G|DDep3*0.84|rep1 0.84 10 3 G|DDep3*0.84|rep2 0.84 10 3 G|DDep3*0.84|rep3 0.84 10 3 G|DDep2.52|rep1 2.52 10 1 G|DDep2.52|rep2 2.52 10 1 G|DDep2.52|rep3 2.52 10 1 G|DDep3*2.52|rep1 2.52 10 3 G|DDep3*2.52|rep2 2.52 10 3 G|DDep3*2.52|rep3 2.52 10 3
Paper-based substrates were fabricated by the same assembly used for glass, but the circular holes were covered with paper. The paper used, unlike the glass, didn’t suffer any type of cleaning process. A square piece of paper (6mm side) was placed between the plastic band and the glass slide in each hole. The paper was cut slightly larger than the circles in the plastic in order to prevent the AgNSs suspension to spread out of the paper. As the solution spreads throughout the paper, there was the need to keep it small. Table 3 summarizes the data regarding the paper substrates.
Table 3: Prepared paper substrates by direct deposition and the different fabrication conditions: nanostar suspension concentration, number of deposit droplets and DTNB concentration.
Sample AgNSs concentrations (nM) DTNB concentration (mM) Number of DTNB depositions Paper 0 0 0 P|DTNB 0 10 1 P|DDep1.84 1.84 0 0 P|DDep0.23 0.23 10 1 P|DDep0.46 0.46 10 1 P|DDep0.92 0.92 10 1 P|DDep1.38 1.38 10 1 P|DDep1.84 1.84 10 1
3.3.2 AgNSs deposition using the centrifuge sedimentation method
For this deposition technique, a syringe and a centrifuge tube were used (see figure 16(a)). A glass (squares of 0.8 × 0.8cm) were placed on the bottom of 5 mL syringes (of 1.2 cm internal diameter, 6cm of length and 0.1cm of wall thickness), with the tip covered with hot glue and the free extremity cut off. The ensemble was placed inside the centrifuge tube of 15 mL, as shown in figure 16(b).
Figure 16: a) Preparation of syringe and centrifuge tube; b) Experimental assembly: the syringe was placed inside the centrifuge tube.
Following the method described by Markelonis et al.[9], the glass slides were washed with Aqua-Regia, Milli-Q water and finally with methanol, before being placed in the bottom of the syringes. 400L of AgNSs (1.5nM) and 1mL of MeOH were added to the syringes. The samples were sonicated for 1 minute and then centrifuged in a RS-720G swinging bucket rotor of a Kubota Centrifuge, 2 times for 8 minutes at 3200 rpm (1147g), figure 17 shows a schematic diagram of the centrifuge deposition process. At the end, the remaining liquid supernatant was discarded and the substrates were stored in open Eppendorf tubes to dry for 24 hours, for later use.
Figure 17: Schematic diagram to illustrate the centrifuge deposition process [9].
To find the best deposition conditions, shown in the procedure presented above, some variations were tested. Table 4 presents all the substrates of glass made and the changes tested during the process.
Table 4: Glass substrates made by centrifuge sedimentation method and the different fabrication conditions: AgNSs concentration, DTNB concentration, MeOH volume and centrifuge times.
Sample Label AgNSs Concentration (nM) DTNB Concentration (mM) MeOH Volume (mL) Centrifuge Times G|CDep0.2|rep1 0.20 0.02 0 1 G|CDep0.2|rep2 0.20 0.02 0 1 G|CDep0.2|rep3 0.20 0.02 0 1 G|CDep0.2|rep4 0.20 0.02 0 1 G|CDep0.2|rep5 0.20 0.02 0 2 G|CDep0.2|rep6 0.20 0.02 0 2 G|CDep1|rep1 1.00 0.02 0 1 G|CDep1|rep2 1.00 0.02 0 2 G|CDep1.73|rep1 1.73 0.02 0 1 G|CDep1.73|rep2 1.73 0.02 0 2 G|CDep1.40|rep1 1.40 0.02 1 2 G|CDep1.40|rep2 1.40 0.02 1 2 G|CDep0.7|rep1 0.70 0.02 1 2 G|CDep0.7|rep2 0.70 0.02 1 2 G|CDep0.19 0.19 0.02 1 2 G|CDep1 1.00 0.02 1 2 G|CDep0.71 0.71 0.02 1 2 G|CDep1.5|rep1 1.50 100 1 1 G|CDep1.5|rep2 1.50 0.02 1 1 G|CDep1.5|rep3 1.50 DTNB/GSH 1 2 G|CDep1.5|rep4 1.50 DTNB/GSH 1 2 G|CDep1.5|rep5 1.50 DTNB/GSH 1 2 G|CDep1.5|rep6 1.50 DTNB/GSH 1 2 G|CDep1.5|rep7 1.50 DTNB/GSH 1 2 G|CDep1.5|rep8 1.50 DTNB/GSH 1 2 G|CDep1.5|rep9 1.50 DTNB/GSH 1 2 G|CDep1.5|rep10 1.50 DTNB/GSH 1 2 G|CDep1.5|rep11 1.50 DTNB/GSH 1 2 G|CDep1.5|rep12 1.50 DTNB/GSH 1 2
Table 5: Paper substrates made by centrifuge sedimentation method and the different fabrication conditions: AgNSs concentration, DTNB concentration, MeOH volume and centrifuge times.
Sample label AgNSs concentration (nM) DTNB concentration (mM) MeOH volume (mL) Centrifuge times P|CDep0.2|rep1 0.20 0.02 0 1 P|CDep0.2|rep2 0.20 0.02 0 1 P|CDep0.2|rep3 0.20 0.02 0 1 P|CDep0.2|rep4 0.20 0.02 0 1 P|CDep1 1.00 0.02 1 2
3.4
Deposition of the SERS probe
The study of the performance of substrates was done using a probe molecule, DTNB diluted in methanol. In the direct deposition, one drop of 10µL of 10mM DTNB solution was deposited in each of the AgNSs substrates. After 10 min, the glass slide was washed with MeOH. For the centrifu-gation method, DTNB with a concentration of 2mM was prepared and then diluted 100 times. Then 10µL were dropped in the center of the substrate and left to dry for 30 minutes. To test the SERS efficiency of the substrates, only DTNB was used.
The redox state of the DTNB molecule was studied only using the substrates deposited by the cen-trifuge method with a nanostar suspension of 1.5nM. Two different studies were made to test the redox state of the samples. First, the reduced state of DTNB, TNB, was studied and its Raman spectrum compared with the one of the original molecule. For this study, TNB (0.1M) was prepared by reduction of DTNB with sodium borohydride (NaBH4). For the oxidized form, complete oxidation of DTNB
(0.1M) was performed by reaction with hydrogen peroxide (H2O2). To verify the oxidation/reduction
of DTNB, the sample was previously analyzed by UV-vis. The samples were prepared by drop-casting 10µL of solution with different molar ratios of DTNB/TNB in the substrates, in substrates prepared with AgNSs (1.5nM) by the centrifuge sedimentation method.
A similar study was made using the thiol, glutathione (GSH). In this case, the GSH served as the reducing agent of the DTNB, as shown in figure 18. The aim was to find a way to measure the concentration of GSH deposited, by watching the states of the DTNB reduction process in the SERS spectrum. For the sample preparation, 15µL of GSH solution with different concentration were drop-casted in a substrate already functionalized with 15µL of DTNB (c=0.02mM).
Figure 18: Reaction scheme of the DTNB reduction with GSH to form TNB.
3.5
SERS instrumentation and result analysis
The spectra were recorded using a T64000 Jobin-Yvon spectrometer at Instituto de Física dos Materiais
de Universidade do Porto (IFIMUP). This spectrometer operates in the triple subtractive mode, and
is equipped with a cooled liquid nitrogen to cool the charge-coupled device (CCD). The T694000 was used in a microanalysis mode, suitable for the detailed analysis of surfaces. The microanalysis system consists of a microscope equipped with a view screen that allows for the choice of the spots of the sample to be analyzed.[10] The excitation source was a linear polarized argon laser emitting at 514.53nm in a continuous mode. The laser was also working in a power mode in order to ensure 10mW at the focus of the substrate sample. This power has suitable for all the Raman and SERS experiments, avoiding both sample heating and photodegradation. In this work, a 50x lens was used to focus the laser beam and to collect the scattered signal from the sample.
Figure 19: Scheme of the T64000 system in subtractive mode. [10]
For the liquid samples the solution was placed in a quartz cell and the Raman spectra were recorded in a 90o geometry, using a macrosampling mode.
A wide-range of acquisition times was used for the different samples. All the spectra were divided by the acquisition time. Thus, the intensity of the graphics presented does not depend on the acquisition time. Furthermore, the intensity of the laser and the lens were also kept constant in all the acquisitions. The Raman-scattering spectra were analyzed using the Origin Pro and the Igor Pro programs. The first permits the band analysis. The last simulates the Raman spectra by using a sum of damped oscillators according to the following equation [32]:
I(w, T ) = [1 + n(w, T )] N X j=1 Aoj wΩ2ojΓoj (Ω2 oj− w2)2+ w2Γoj , (15)
where n(w,T) is the Bose-Einstein factor, Aojis the strength, Ωojis the wavenumber and Γojis the
4
Results and Discussion
This chapter will present and discuss the results obtained from the characterization of the as-produced AgNSs, and SERS performance of the prepared substrates. We will compared the deposition technique with the substrate performance, regarding the SERS signal recorded at different substrate surface positions, in order to evaluate the reproducibility of the recorded signal. The results obtained using the best substrates here produced were then compared with those obtained with a commercial one. Finally, the analysis of the results obtained in the study of the redox state of the sample will be presented and discussed.
4.1
Optical and Morphological characterization of the silver nanostars
The optical and morphological characterization of the as-prepared nanostars were performed in order to assess the quality of the produced particles for our purposes. Figure 20(a) shows a representative UV-vis spectrum of a suspension of the as-prepared AgNSs. For comparison, figure 20(b) depicts the UV-vis spectrum of the nanostars prepared by Garcia et al. [11]. In both cases, a peak is observed at approximately 380 nm, assigned to a plasmon resonance, likely from the spherical core of the AgNSs [11]. No other peaks, corresponding to plasmonic modes in the tips of the AgNSs, were detected in the spectrum of figure 20 (a), but a strong extinction background was detected in the wavelength range 450-1000 nm. The excitation background at lower wavelengths is due to the absorption and scattering emissions produced by the different morphologies of all the nanoparticles, with different number of arms or length [11].
Figure 20: Excitation spectra of the AgNSs: a) spectrum from the as-processed nanostars in this work; b) spectrum of AgNSs from Garcia et al. [11].