Applications of Raman Spectroscopy to Biology: From Basic Studies to Disease Diagnosis [1st ed.] 978-1-61499-020-8

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APPLICATIONS OF RAMAN SPECTROSCOPY TO BIOLOGY

Advances in Biomedical Spectroscopy Spectroscopic methods play an increasingly important role in studying the molecular details of complex biological systems in health and disease. However, no single spectroscopic method can provide all the desired information on aspects of molecular structure and function in a biological system. Choice of technique will depend on circumstance; some techniques can be carried out both in vivo and in vitro, others not, some have timescales of seconds and others of picoseconds, whilst some require use of a perturbing probe molecule while others do not. Each volume in this series will provide a state of the art account of an individual spectroscopic technique in detail. Theoretical and practical aspects of each technique, as applied to the characterisation of biological and biomedical systems, will be comprehensively covered so as to highlight advantages, disadvantages, practical limitations and future potential. The volumes will be intended for use by research workers in both academic and in applied research, and by graduate students working on biological or biomedical problems. Series Editor: Dr. Parvez I. Haris De Montfort University, Leicester, United Kingdom

Volume 5 Recently published in this series Vol. 4. Vol. 3. Vol. 2. Vol. 1.

A.B. Dahlin, Plasmonic Biosensors – An Integrated View of Refractometric Detection A.J. Dingley and S.M. Pascal (Eds.), Biomolecular NMR Spectroscopy A. Barth and P.I. Haris (Eds.), Biological and Biomedical Infrared Spectroscopy B.A. Wallace and R.W. Janes (Eds.), Modern Techniques for Circular Dichroism and Synchrotron Radiation Circular Dichroism Spectroscopy

ISSN 1875-0656 (print) ISSN 1879-811X (online)

App plicatio ons off Ramaan Speectrosccopy ogy to Biolo From Basic Studies to Disease Diagnosis

Edited by

Mahmoud Gh homi Groupe de Biophysique B Moléculaire,, UFR Santé-M Médecine-Biiologie Humaine, Univerrsité Paris 133

Amstterdam • Berrlin • Tokyo • Washington, DC

© 2012 The authors and IOS Press. All rights reserved. No part of this book may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, without prior written permission from the publisher. ISBN 978-1-60750-999-8 (print) ISBN 978-1-61499-020-8 (online) Library of Congress Control Number: 2011944964 Publisher IOS Press BV Nieuwe Hemweg 6B 1013 BG Amsterdam Netherlands fax: +31 20 687 0019 e-mail: [email protected] Distributor in the USA and Canada IOS Press, Inc. 4502 Rachael Manor Drive Fairfax, VA 22032 USA fax: +1 703 323 3668 e-mail: [email protected]

LEGAL NOTICE The publisher is not responsible for the use which might be made of the following information. PRINTED IN THE NETHERLANDS

Applications of Raman Spectroscopy to Biology M. Ghomi (Ed.) IOS Press, 2012 © 2012 The authors and IOS Press. All rights reserved.

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Series Editor’s Preface The sequence of the human genome and that of other organisms have provided a wealth of biological data that needs to be effectively utilised for our understanding of diverse biological processes. Translation of this information is vital for countering disease processes through the development of drugs and diagnostic tools. Progress in this area is much slower compared to successfully completing the genome sequence of yet another organism. Advances in drug discovery and development of diagnostic tools require a multidisciplinary research approach that brings together scientists with expertise ranging from physics and engineering to biochemistry and molecular biology. Improvements in existing technologies need to go hand-in-hand with the development of new techniques for understanding biological systems. The example of Raman spectroscopy can be used to show how an existing technique can be “updated” to address complex biological problems. The Indian scientist Chandrasekhar Venkata Raman was awarded the Nobel Prize in Physics (1930) for his role in the discovery, in 1928, of a physical phenomenon to be later called the Raman effect. Recent advances in technology (lasers, detectors, filters, and computers) have made possible the application of this relatively “old” technique to address problems in some of the most advanced fields of scientific research. This includes applications in both basic and applied sciences including nanotechnology, disease diagnosis and drug discovery. Mahmoud Ghomi has edited this current book covering the applications of Raman Spectroscopy to Biology: from basic studies to disease diagnosis. It nicely complements other books published in this series which focuses on the latest advances in different biophysical techniques that are used in biomedical research. Other books published in this series are as follows: Biological and Biomedical Infrared Spectroscopy (A. Barth and P.I. Haris); Modern Techniques for Circular Dichroism and Synchrotron Radiation Circular Dichroism Spectroscopy (B.A. Wallace and R.W. Janes); Biomolecular NMR Spectroscopy (A.J. Dingley and S.M. Pascal); Plasmonic Biosensors – An Integrated View of Refractometric Detection (A.B. Dahlin). The current and the existing books in this series have been possible thanks to the good will and co-operation of all the scientists who accepted my invitation to either edit/author a book or contribute Chapters. I cannot thank enough my good friend, Peter Brown, whose constant support and encouragement has been invaluable for both initiating and successfully completing the publication of each of the books in this series. Finally, Anne Marie de Rover and her colleagues at IOS Press are thanked for their support, diligence and hard work in organising the printing of the book. Parvez I. Haris Leicester, United Kingdom

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Preface From a fundamental point of view, Raman spectroscopy is based on the inelastic scattering of light by matter in one of its phases: gas, liquid and solid. The scattered light can have an energy (or a frequency) either higher, or lower than that of the incident light, giving rise to anti-Stokes or Stokes Raman effects, respectively. In the framework of quantum physics, the interaction of the light (photons) with matter, induces transitions between its vibrational states. Consequently, the bands observed on Raman spectra (representing the scattering intensity versus wavenumber, i.e. inverse of wavelength, routinely expressed in cm-1 units) reflect in fact these vibrational transitions. An important fact in the field of molecular Raman spectroscopy is that the Raman spectrum of a molecule and its conformational properties are closely related. This is of a fundamental interest in the case of organic molecules, in which rotations around the chemical bonds (torsions), generally lead to the generation of a variety of conformers. Many of these conformers may coexist at normal conditions (temperature and pressure), because of the small amount of energy separating them in the molecular energy landscapes. Conformational transitions may be induced by varying temperature, pressure, or other physicochemical parameters, and can be detected by Raman spectroscopy through the variation of Raman intensity and/or vibrational wavenumbers. This is the reason why we generally speak of Raman conformational markers, which recall the Raman bands considered as the fingerprints of different conformations. This spectroscopic tool can also be used to recognize the interaction sites of two molecular partners, through the changes observed in the intensity or wavenumbers of the molecular groups involved in the interactions. Simultaneously, one can also follow the conformational changes of interacting molecular partners by means of their Raman conformational markers. The application of Raman spectroscopy to biological molecules has rapidly grown up since the early 1970s, presumably because of the arrival of lasers (monochromatic and coherent sources of light) and modern dispersive spectrometers. The first period of this application was completely devoted to the constitution of a spectral database recorded from different important biological molecules (proteins, nucleic acids, phospholipids, etc) in aqueous solutions, i.e. natural medium of biological molecules and systems. Then the time arrived for studying more complex systems such as cells and tissues. This was especially made possible by the setup of microRaman spectrometers. Furthermore, the progress in microcomputer manufacturing together with the elaboration of rapid mathematical and statistical algorithms, could offer to Raman spectroscopy the possibility of acting as a powerful imaging tool. A Raman image, collected from a complex biological system, i.e. cell or tissue, can provide useful information on the localization or distribution of different constituting molecules. This is the point where global information (image) can also carry local (molecular) information. It was also natural to use Raman spectroscopy as a tool for diagnosis, for instance in analyzing metabolic effects in biological fluids, or alterations induced in tissues upon tumor formation, such as changes in DNA, protein and lipid conformations, or molecular degradation. Thanks to the continuing technical progress, portable miniature Raman spectrometers can be used in the future as a non-invasive tool for medical diagnosis.

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Raman spectroscopy is widely recognized as a label-free technique; the collected signal in a spectrometer directly arises from the vibrational motions of different chemical groups of the analyzed molecule. Moreover, the contribution of water to Raman signal is rather low compared to infrared absorption. The weakness of this technique arises from its low intensity: only one scattered photon out of one million incident ones, contributes to the Raman scattering process. As a consequence, the samples used for recording Raman spectra are generally much more concentrated than those needed for other optical spectroscopic methods such as fluorescence and circular dichroism. To overcome this difficulty, two different approaches have been developed during the last years. The first one is the resonance Raman spectroscopy (RRS), which is based on the excitation of a sample by means of a monochromatic light, of which the wavelength falls within its molecular absorption spectrum. This phenomenon leads to a considerable enhancement of Raman intensity, and allows the sample concentration to be decreased. However, the absorption of light by biological molecules corresponding to their absorption spectra, generally located in the ultra-violet range of electromagnetic radiation, can cause irreversible damages due to their degradation. The second possibility is to use the Raman signal enhancement effect by the so-called plasmon resonance, when the analyzed molecule is adsorbed on the surface of a noble metal, i.e. gold or silver, colloid or nanoparticle. This method, known as surface enhanced Raman spectroscopy (SERS) has found a large audience for analyzing the biological molecules. SERS signal was shown to be enhanced up to several orders of magnitude, depending on the molecular affinity for binding to the metal surface, as well as on the molecular group involved in the adsorption. This technique is gradually becoming a routine tool in pharmacology and molecular biology for analyzing molecular samples at very low concentrations. Mahmoud Ghomi Paris, 2011

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List of Contributors Guannan Chen

Rong Chen

Yves-Marie Coïc

Shangyuan Feng

Sheila E. Fisher

José Vicente García-Ramos

Mahmoud Ghomi

Catherine Gouyette Andrew T. Harris

Belén Hernández

Key Laboratory of Optoelectronic Science and Technology for Medicine, Ministry of Education and Fujian Provisional Key Laboratory for Photonics Technology, Fujian Normal University, Fuzhou 350007, China. Key Laboratory of Optoelectronic Science and Technology for Medicine, Ministry of Education and Fujian Provisional Key Laboratory for Photonics Technology, Fujian Normal University, Fuzhou 350007, China. Unité de Chimie des Biomolécules, URA 2128, Département de BSC, Institut Pasteur, 28 rue du Docteur Roux, 75724 Paris Cedex 15, France. Key Laboratory of Optoelectronic Science and Technology for Medicine, Ministry of Education and Fujian Provisional Key Laboratory for Photonics Technology, Fujian Normal University, Fuzhou 350007, China. PhD, MSc, FRCS, Clinical Research Fellow, Section of Experimental Therapeutics, Leeds Institute of Molecular Medicine, University of Leeds, St. James’s University Hospital, Beckett Street, Leeds, West Yorkshire, LS9 7JT, United-Kingdom. Hon Senior Research Fellow, School of Health Studies, University of Bradford. Instituto de Estructura de la Materia, Consejo Superior de Investigaciones Científicas, Serrano 121, 28006-Madrid, Spain. Groupe de Biophysique Moléculaire, UFR SantéMédecine-Biologie Humaine, Université Paris 13, 74 rue Marcel Cachin, 93017 Bobigny cedex, France. PF1 Génomique, Génopole, Institut Pasteur, 25/28 rue du Docteur Roux, 75724 Paris Cedex 15, France. PhD, MRCS, Registrar Otolaryngology–Head and Neck Surgery, Leeds Teaching Hospitals NHS Trust, Visiting Lecturer, Department of Oral Biology, Leeds. Dental Institute, University of Leeds. Groupe de Biophysique Moléculaire, UFR SantéMédecine-Biologie Humaine, Université Paris 13, 74 rue Marcel Cachin, 93017 Bobigny cedex, France.

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Zufang Huang

Daniel Jancura Gejza Lajos Yongzeng Li

Juqiang Lin

Raquel de Llanos

Pavol Miškovský Cees Otto

Marek Procházka

Kerstin Ramser

Santiago Sánchez-Cortés

Paz Sevilla

Josef Štĕpánek

Key Laboratory of Optoelectronic Science and Technology for Medicine, Ministry of Education and Fujian Provisional Key Laboratory for Photonics Technology, Fujian Normal University, Fuzhou 350007, China. Division of Biophysics, Safarik University, Jesenna 5, 41 54 Kosice, Slovakia. Division of Biophysics, Safarik University, Jesenna 5, 41 54 Kosice, Slovakia. Key Laboratory of Optoelectronic Science and Technology for Medicine, Ministry of Education and Fujian Provisional Key Laboratory for Photonics Technology, Fujian Normal University, Fuzhou 350007, China. Key Laboratory of Optoelectronic Science and Technology for Medicine, Ministry of Education and Fujian Provisional Key Laboratory for Photonics Technology, Fujian Normal University, Fuzhou 350007, China. Instituto de Estructura de la Materia. Consejo Superior de Investigaciones Científicas, Serrano 121. 28006-Madrid, Spain. Division of Biophysics, Safarik University, Jesenna 5, 41 54 Kosice, Slovakia. Medical Cell BioPhysics, MESA+Institute for Nanotechnology, MIRA Institute for Biomedical Technology and Technical Medicine, University of Twente, Enschede, P.O. Box 217, 7500 AE, The Netherlands. Charles University, Faculty of Mathematics and Physics, Institute of Physics, Ke Karlovu 5, Prague 2, CZ-121 16, Czech Republic. Department of Computer Science and Electrical Engineering, Luleå university of Technology, SE-971 87 Luleå, Sweden. Instituto de Estructura de la Materia, Consejo Superior de Investigaciones Científicas, Serrano 121, 28006-Madrid, Spain. Departamento de Química-Física II, Facultad de Farmacia, Universidad Complutense de Madrid, 28040-Madrid, Spain. Charles University, Faculty of Mathematics and Physics, Institute of Physics, Ke Karlovu 5, Prague 2, CZ-121 16, Czech Republic.

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Vishnu Vardhan Pully

Jing Wang

Haishan Zeng

Medical Cell BioPhysics, MESA+Institute for Nanotechnology, MIRA Institute for Biomedical Technology and Technical Medicine, University of Twente, Enschede, P.O. Box 217, 7500 AE, The Netherlands. Key Laboratory of Optoelectronic Science and Technology for Medicine, Ministry of Education and Fujian Provisional Key Laboratory for Photonics Technology, Fujian Normal University, Fuzhou 350007, China. Imaging Unit – Integrative Oncology Department, British Columbia Cancer Agency Research Centre, Vancouver, BC, Canada.

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Contents Series Editor’s Preface Parvez I. Haris

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Preface Mahmoud Ghomi

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List of Contributors

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Surface-Enhanced Raman Scattering (SERS) and Its Application to Biomolecular and Cellular Investigation Marek Procházka and Josef Štĕpánek

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Surface-Enhanced Fluorescence and Raman Spectroscopy (SEF and SERS) of Anthraquinone Anti Tumoral Drugs and Their Complexes with Biomolecules on Ag Nanoparticles Paz Sevilla, Gejza Lajos, Raquel de Llanos, Santiago Sánchez-Cortés, Daniel Jancura, Pavol Miškovský and José Vicente García-Ramos

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Probing the Interactions of Oligodeoxynucleotides with a Cationic Peptide by Raman Scattering Belén Hernández, Yves-Marie Coïc, Catherine Gouyette and Mahmoud Ghomi

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Applications of SERS Spectroscopy for Blood Analysis Rong Chen, Juqiang Lin, Shangyuan Feng, Zufang Huang, Guannan Chen, Jing Wang, Yongzeng Li and Haishan Zeng

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Raman Spectroscopy of Single Cells for Biomedical Applications Kerstin Ramser

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Hyperspectral Raman Microscopy of the Living Cell Cees Otto and Vishnu Vardhan Pully

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The Application of Raman Spectroscopy to Human Cancer Diagnostics Andrew T. Harris and Sheila E. Fisher

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Author Index

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Applications of Raman Spectroscopy to Biology M. Ghomi (Ed.) IOS Press, 2012 © 2012 The authors and IOS Press. All rights reserved. doi:10.3233/978-1-61499-020-8-1

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Surface-enhanced Raman Scattering (SERS) and its Application to Biomolecular and Cellular Investigation Marek Procházka1, Josef Štpánek Charles University, Faculty of Mathematics and Physics, Institute of Physics, Ke Karlovu 5, Prague 2, CZ-121 16, Czech Republic. Abstract. Surface-enhanced Raman scattering (SERS) is a specific technique of Raman spectroscopy when signal enhancement of several orders is achieved for numerous molecules placed in the closest vicinity of certain rough metal surfaces. Despite of some experimental difficulties and undesirable concomitant effects, this technique provides numerous promises for both highly specific and sensitive measurements. This chapter reviews the state-of-the-art in the field of SERS applications in biomolecular, cellular and biomedical studies. Prior to this, the reader is familiarized with the basics of SERS phenomenon and its experimental aspects including the most common or promising types of SERS-active substrates. Keywords. biomolecules, SERS substrates, SERS labels, immunoassays, biosensors, intracellular SERS, in vitro SERS, in vivo SERS

Introduction Raman spectroscopy embodies advantages of highly specific molecular spectral pattern and optical-scattering measuring optical scheme, which similarly as fluorescence spectroscopy can be easily combined with confocal microscopy and other techniques of optical imaging. A severe limitation for application of this spectroscopic technique consists in the low cross section of the Raman scattering (RS) process (between 10-30 and 10-25 cm2). The effect of giant Raman intensity enhancement obtainable for molecules in the close vicinity of certain rough metal surfaces can overcome this problem providing in some cases the cross section at the order of 10-16 cm2 per molecule, corresponding to 12-14 order of magnitude enhancement compared with standard RS. The unique sensitivity of Surface-Enhanced Raman Scattering (SERS) is though counterpoised by the necessity to prepare specific SERS-active substrate and to solve numerous added tasks, like a proper adsorption of the analyte at the enhancing surface or interpretation (and/or reduction) of the surface effect on the geometry, physicochemical properties, and Raman spectrum of the analyte. 1

Corresponding Author: Marek Procházka

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SERS was originally first of all considered as a very sensitive analytical technique capable to detect specific molecules in extremely low amounts. Due to the recent development of new SERS-active substrates, labeling and derivatization chemistry, as well as new instrumentations SERS became very promising tool for a wide field of applications, including bioanalytical, biophysical and biomedical studies. In this chapter we review contemporary knowledge and achievements in this field considering also limitations, possibilities, and prospects of this technique.

1. Basics of SERS SERS was discovered in seventies by the group of M. Fleischmann [1], who observed surprisingly very strong Raman signal of a thin layer of pyridine on an electrochemically roughened silver electrode. The phenomenon was soon confirmed and quantified by other two groups [2, 3], reporting the 105 – 106 enhancement for pyridine adsorbed on the roughened silver electrode. Nowadays, SERS is understood as a very large enhancement of Raman cross-section (basically 105 – 107, but even 1012 in some special cases) for molecules adsorbed on roughened metal surfaces. 1.1. SERS enhancement factor The SERS enhancement factor (EF) is a key characteristic of the SERS effect. Up to now, several types of SERS EFs have been proposed with the aim to find an optimal quantity enabling to compare experiments across different substrates and different conditions as well as theoretical calculations [4, 5]. It is thus worth to deal with its formulation prior to describe details of the SERS experiment. The widely used definition compares integral intensities of the strongest band in SERS (ISERS) and conventional Raman (IRS) spectrum, normalized to numbers of molecular scatters participating in respective cases, measured at the same set-up [6]: EF

I SERS / N SERS I RS / N RS

In the case of backscattering experiment, the value of NSERS/NRS is often estimated as a ratio of the effective surface density of adsorbed molecules in SERS measurement to the spatial molecular density in conventional Raman measurement multiplied by the effective high of the scattered volume [5, 7]. The analytical enhancement factor (AEF), used in SERS analytical applications, considers ratio of molecular concentrations in analyzed solutions [5]: AEF

I SERS / c SERS I RS / c RS

This parameter is suitable to display how the surface enhancement improves analytical capability of Raman method in particular cases, but depends not only on the

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SERS phenomenon but also on the type of the substrate and its coverage by the adsorbate. 1.2. SERS mechanisms First theories of the SERS mechanism have already appeared in the cited above early works of D.J. Jeanmaire and R.P. Van Duyne, and M.G. Albrecht and J.A. Creighton, but up to now they are further developed and refined (see e.g. [8-16]). Simple speculation that the induced dipole moment P, which characterizes the effect of optical radiation on a molecule, is given by a product of a molecular polarizability D and an electric vector E, P = D( shows two possible ways how to explain SERS effect: interaction of the molecule with a rough metal surface must enhance either D or E. There are really two considered basic mechanisms: chemical (or molecular) mechanism and electromagnetic mechanism. Electromagnetic mechanism is based on enhancement of the electromagnetic field due to resonance excitations of localized conduction-electron oscillations at the roughened metal surface, called surface plasmons (SPs). The coupled state of the photon and localized SP (polariton) is accompanied by sharply enhanced amplitude of the electromagnetic field in the closest vicinity of the roughened metal surface. The molecule adsorbed at the surface is thus subjected to much stronger E. If the molecule emits RS at a frequency still within the region of SP resonance, the same mechanism may enhance Raman signal [8, 17]. The electromagnetic enhancement factor EFem(Qs) can be described explicitly for a small metal sphere (its radius r is smaller than one twentieths of the incident light wavelength) as: EF em Q S #

H Q L  H 0 H Q L  2H 0

2

2

H Q S  H 0 § r · ¸ ¨ H Q S  2H 0 © r  d ¹

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where HQ is the complex frequency-dependent dielectric function of the metal, Ho is the dielectric constant of the bulk medium, d is the distance of the molecule from the surface. Index L stands for the incident and S for the scattered light. This equation well illustrates two very important characteristics of SERS produced by electromagnetic mechanism: 1. Large SERS enhancement is obtained if, at the wavelengths of the incident and/or scattered radiation, the real part of HQ is close to –2Ho and the imaginary (damping) part of HQ is small. In other words, matching of resonance conditions for SP excitation is required to achieve high enhancement. Only the free-electron-like (noble) metals (Ag, Au, Cu) or the alkali metals (group Ia) are therefore proper materials for surface-enhancing substrates (EF up to 105) when the common visible or near-infrared excitation is used. Transition metals (namely Pt, Ru, Rh, Pd, Fe, Co, and Ni) are also applicable [18, 19], but they are generally less enhancing (EF up to 101-104) [19]. On the other hand, they also provide SERS when excited by ultraviolet radiation [19, 20]. 2. The scattered molecule does not need to be in a direct contact with the metal surface; the intensity of RS falls though off very rapidly with the increasing distance d from the surface. It has been demonstrated that the Raman enhancing effect is detectable for d10 nm [21, 22].

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Due to an inhomogeneity of the enhanced electromagnetic field, the selection rules for SERS spectroscopy are not exactly the same as those of conventional Raman spectroscopy [23]. Three types of vibrational modes may be found in SERS spectrum: (1) those excited by the normal component of the field and resulting in an induced dipole with a strong component perpendicular to the surface, (2) those excited by the tangential component of the field with a strong induced dipole tangential to the surface and (3) the mixed cases. Analysis of the SERS spectrum may thus also reveal the adsorbate orientation with respect to the metal surface [24, 25]. The chemical (or molecular) mechanism lies in increased molecular polarizability D, and consequently Raman cross-section, as a result of the surface interaction [26]. The molecule is expected to be at the metal SERS-active site chemisorbed [6], i.e. bound with the adsorption enthalpy comparable to a chemical-bond energy (more negative than –40 kJmol-1). Chemisorption allows formation of a charge-transfer (CT) complex between the molecule and an optically-excited conductive electron at the metal surface, which expands the effective molecular polarizability [27, 28]. The chemical mechanism is generally weaker than the electromagnetic one, and contributes to the total enhancement by a factor of 101-103 [9, 29]. On the other hand, surface roughness and the presence of localized SPs are not required for the chemical mechanism as has been demonstrated by SERS measurements on smooth copper surface [30]. Measurement of the EF distance dependence is a good way how to estimate the proximity of the adsorbate functional groups to the surface as well as to identify the respective contribution from each of the two mechanisms, since the chemical one requires a “direct“ contact between the metal and the adsorbate while the electromagnetic not [22]. 1.3. Surface enhancement in real SERS experiments EF strongly depends on properties of the SERS-active system (metal substrate and analyte) and experimental conditions (excitation wavelength). Maximal EF provided by isolated gold and silver spheroids is 103-104 and 106-107, respectively [31]. SERSactive substrates often consist of a collection of aggregated nanoparticles (NPs) exhibiting fractal structure [32, 33]. In this case, the conditions for SP excitation are modified. Whereas the sphere resonates at a particular frequency, its random aggregate or fractal could resonate at different wavelengths within broad spectral region. Certain localized resonances in fractals are the source of extremely high enhanced local field and can produce SERS EF of 1012 or higher. For closely spaced but interacting NPs (dimers) with gap of 0.5-1 nm, EF up to 1010 was predicted [34-36]. Similar effect is observed for trimers [37], nanolenses [38] or interpenetrating (intergrowing and/or sintered) NPs [39]. All these structures contain SERS-active sites called “hot spots” with extremely high enhancement (up to 1011) [40]. It is necessary to note that not all surface sites provide SERS enhancement. Only one out of 100 to 1000 NPs is optically hot (EF ~ 102-103) and only one out of 10000 surface sites on a hot particle shows efficient EF (106-107) [40]. Number of “hot spots” is extremely low: for example, Aroca [13] estimated that less than 0.5% of sites are “hot spots”. Other studies show that only 3% of illuminated molecules (a total approximately of 5-15) are localized in “hot spots” [36]. Only selective excitation of such “hot spots” leads to EF ~ 1012 allowing spectral detection in single molecular

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levels (single-molecule SERS) [40-42]. In ordinary SERS systems, EF value of 105-106 is obtained as an average from a few highly enhancing “hot spots” and many weak enhancing sites. 1.4. Surface effect on other optical phenomena Interaction of electromagnetic field with nanostructured metal surface also strongly influences other optical phenomena, namely absorption and fluorescence. Absorption can be enhanced generally by EF 101-103. Surface-enhanced infrared absorption (SEIRA) is often used as a complementary technique to SERS [13, 43]. In contrast to that, surface effects on fluorescence intensity are diverse. Fluorescence of highly fluorescent species adsorbed on metal nanostructures is often completely quenched [44, 45], which is considered to be a big advantage of SERS over the conventional RS. On the other hand, in the case of molecules with low quantum yield (2 transmittance at 490 nm increases in the first milliseconds and after decreases until the final value (Figure 16). The first part corresponds to a fast binding, the second one to a slow binding. In concordance with the kinetics, absorbance at 490 nm decreases in the first process and increases in the second one. According to Figure 14 (left) the first process means a

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change of absorbance value from emodin to emodin-BSAf complex and the second from the emodin-BSAf complex to emodin-BSA complex. This indicates the first binding corresponds to emodin that situates in Sudlow´s site II and the second bonding to emodin situates in Sudlow´s site I. 4.2.

Hypericin: interaction with LDL. SERS and interaction model.

LDL particles are the main carriers of cholesterol in the human circulation and are thus key players in cholesterol transfer and metabolism. Both physical and physiological characteristics have led to a common convention for defining LDL particles as lipoprotein particles within the density limits of 1.019-1.063 g/ml. Therefore, LDL forms a heterogeneous group of particles varying greatly in size, composition and structure [57]. Current knowledge and understanding of the LDL particle structure is therefore far from trivial. Steim and co-workers gave convincing evidence in 1968 [58] for the micellar model of lipoprotein particles. In the mid 1970s the LDL surface was suggested to be a trilayer or a bilayer, the protein moiety existing in the core of the LDL particle [59, 60]. Piece by piece, however, the data have led to the development of the current picture of the spherical lipoprotein particle as an amphipathic monolayer surrounding a hydrophobic lipid core. Moreover, a biophysical model has recently been developed for micellar lipoprotein particles [60]. More specifically, LDL particles have an average diameter of 22 nm, the core consisting of about 170 triglyceride (TG) and 1600 cholesteryl ester (CE) molecules and the surface monolayer comprising about 700 phospholipid molecules and a single copy of apoB-100 [61] (Figure 17, left).

Figure 18. Left: Fluorescence and Raman spectrum of hypericin/LDL complexes at different concentration ratios in water a), and on AgHx colloid, prepared using hydroxylamine hydrochloride as reducing agent b). The concentration of LDL was maintained constant at 10-8 M. Spectrum were recorded on a Confocal Raman Microscope Renishaw RM2000, with macro objectives (f : 30mm) and using an excitation line provided by a Spectra Physics Model 165 argon ion laser with λexc=514,5 nm. Right: a) SERS spectrum of LDL and hypericin/LDL complexes at the following concentration ratios: 10:1, 50:1, 200:1. b) Comparison between the SERS spectrum of LDL alone (×3) and the hypericin/LDL complex at 10:1 ratio. The concentration of LDL was 10-8 M. AgHx colloid, prepared using hydroxylamine hydrochloride as reducing agent, has been used in SERS experiments. Spectrum were recorded on a Confocal Raman Microscope Renishaw RM2000, with macro objectives (f : 30mm) and using an excitation line provided by a Spectra Physics Model 165 argon ion laser with λexc=514,5 nm.

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In addition, the particles contain about 600 molecules of unesterified cholesterol (UC), of which about one-third is located in the core and two-thirds in the surface [62]. It should also be noted that a few percent of the TG and CE molecules penetrate toward the surface. The main phospholipid components are phosphatidylcholine (PCH) (about 450 molecules/LDL particle) and sphingomyelin (SM) (about 185 molecules/LDL particle). The LDL particles also contain lysophosphatidylcholine (lyso-PC) (about 80 molecules/LDL particle), phosphatidylethanolamine (PE) (about 10 molecules/LDL particle), diacylglycerol (DAG) (about 7 molecules/LDL particle), ceramide (CER) (about 2 molecules/LDL particle) and some phosphatidylinositol [63-65]. The particles are in a dynamic state, their structure and physical properties being dependent on their lipid composition as well as on the conformation of apoB-100. LDL could play a key role in the targeted delivery of hydrophobic and/or amphiphilic photosensitizers to tumor cells in PDT [66-74], due to the enhanced expression of specific LDL receptors (regulated by the cholesterol needs of the cell, usually higher in fast growing cells, like tumor cells and tumor endothelial cells) in many types of transformed cells when compared with nontransformed cells [71, 75, 76]. The application of both the SERS and SEF techniques was aimed to obtain comprehensive information about vibrational and fluorescent properties of hypericin molecules incorporated into LDL and contribute to the more detailed characterization of the localization of hypericin inside LDL at different hypericin/LDL ratios.

Figure 19. Difference emission spectra obtained by subtracting the spectrum of hypericin in DMSO from the fluorescence spectrum at 50:1 a) and 300:1 b) hypericin/LDL ratios. c) Plot of the excimer emission in the absence (o) and in presence (■) of Ag NPs. d) Variation of SERS intensity with the hypericin/LDL ratio measured for the band at 1372cm-1. AgHx colloid, prepared using hydroxylamine hydrochloride as reducing agent, has been used in all SERS experiments.

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The fluorescence and Raman spectra of hypericin/LDL complexes were studied in water and on Ag NPs (Figure 18 (left) a and b, respectively). The experiments were realized at different hypericin/LDL ratios maintaining the constant concentration of LDL at 10-8 M. As can be seen, an increase of the intensity of the main fluorescence peak of hypericin (maximum at 599 nm) with the hypericin concentration is observed in water in the range of the concentration ratios between 1:1 and 50:1 (Figure 17, right). The fluorescence spectra observed in this interval correspond to the hypericin monomer, as corroborates the similarity with the spectrum of hypericin in DMSO [77]. For higher hypericin/LDL ratios (>50:1), a significant decrease of this fluorescence is detected. This result indicates that hypericin interacts with LDL under the monomeric form up to a 50:1 ratio, and above this value, hypericin molecules begin to aggregate upon accumulation inside the LDL structure (Figure 17 (right), inset scheme). The fluorescence quenching is attributed to the drug aggregation which is responsible for the significant decrease of hypericin fluorescence observed at these high concentration ratios, but partially also to the dynamic self-quenching of the singlet excited state of hypericin [77, 78]. No Raman bands of hypericin are observed in water due to the poor Raman emission in the absence of Ag NPs.

Figure 20. Fluorescence of the hypericin monomers (at 599 nm), the excimer emission of hypericin aggregates (at 709 nm), and the SERS intensity of 1372 cm -1 band as function of hypericin/LDL ratio. AgHx colloid, prepared using hydroxylamine hydrochloride as reducing agenthas been used for SERS experiments. These variations were correlated to the possible approach of the drug to the metal surface of Ag NPs. Spectrum were recorded on a Confocal Raman Microscope Renishaw RM2000, with macro objectives (f : 30mm) and using an excitation line provided by a Spectra Physics Model 165 argon ion laser with λexc=514,5 nm.

A broad emission with an apparent center at 709 nm is observed when hypericin/LDL ratio increases (Figure 18, left). This emission is better seen in the difference spectra (Figure 19 a and b) and can be assigned to the excimer emission, which is provoked by the formation of H-aggregates of hypericin, as previously demonstrated [42]. The fluorescence emission of hypericin/LDL complex undergoes a significant intensity decrease in the presence of Ag NPs (Figure 17, right b). This is a consequence of fluorescence quenching caused by the energy transfer between hypericin molecules localized inside LDL and metal colloid. The quenching was not compensated by an intensification of the fluorescence by SEF effect due to the long distance between

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hypericin molecules and the metal surface. This result points out that at low hypericin concentrations the drug is rather placed in the interior part of LDL, where the distance to the metal surface is too far as to undergo a SEF enhancement. On increasing the hypericin/LDL ratio it is observed an enhancement of the excimer emission (Figure 19 c). This enhancement is attributed to two different reasons: the increase of hypericin aggregates in the LDL particle, and the closer position of these aggregates in relation to the metal surface, since they tend to be localized in the external part of the LDL particles as the concentration of hypericin is increased. Both effects are further amplified as the hypericin/LDL ratio is increased. More detailed information about the host and the ligand structures can be obtained from the SERS spectra (Figure 18, right). The amplified SERS spectrum of free LDL is shown in Figure 18 (right) a. This spectrum displays strong features at 1006, 1154 and 1516 cm-1, which correspond to carotenoid compounds normally existing in LDL structure, in particular all-trans-β-carotene and lycopene [79, 80]. These bands are attributed to the in-phase C=C stretching vibrations in the central part of the chain, the C-C stretching also from bonds in the central part, and in-phase combinations of inplane rocking motions contributed by the CH3 groups [81, 82]. In addition, the bands at 1270, 1347 and 1430-60 cm-1 could be attributed to the aliphatic chains of lipidic species existing in LDL. The SERS spectrum of hypericin/LDL complex at the 10:1 ratio exhibited a mixture of bands due to both LDL and hypericin (Figure 18, right b). However, the SERS intensity of hypericin is weaker than the main bands of LDL due to the long distance of hypericin molecules from the NPs, suggesting again that the drug is located in the inner part of the LDL particle al this low ratio. The SERS intensity of hypericin, analyzed for the most intense band at 1372 cm -1, increases linearly with the increase of hypericin/LDL ratio (Figure 19 d). Two different effects are contributing to this: a) the increase of hypericin concentration, and b) the progressive localization of hypericin molecules in the outer shell of LDL, which are indeed closer to the metal surface than the hydrophobic core of LDL. The analysis of the SERS spectra affords detailed structural information since the Raman spectra can be considered as molecular fingerprints showing a different structure of the molecules. At relatively low hypericin/LDL ratios (< 50:1) the SERS spectrum of hypericin corresponds to a mixture of neutral and monoanionic forms, which is deduced on the basis of the results of our previous work, where pH dependence of the SERS spectra of hypericin was studied [42]. At high hypericin/LDL ratios (~200:1) the contribution of the monoanionic form, characterized by the SERS bands at 1249, 1293 and 1372 cm-1, is dramatically increased. This indicates that at very high concentration hypericin molecules are placed in a more polar environment, i.e. in the amphipathic outer part of the LDL particle as already proposed in previous work [42]. Since in this region phospholipids and apoB-100 proteins are localized, an interaction of the hypericinate with the polar groups of these biomolecules is likely occurring. In Figure 20 a summary picture of the variation of the fluorescence of monomer hypericin, excimer emission and SERS intensity of hypericin/LDL complex at different concentration ratios is shown. This scheme is based on the combination of the data coming out from the SERS band at 1372 cm-1, the fluorescence spectra of monomers (emission at 599 nm), and the excimer emission (apparent maximum at 709 nm). It is proposed that at low hypericin/LDL ratios (