Raman spectroscopy: Recent advancements, techniques and applications

Vibrational Spectroscopy 57 (2011) 163–176

Contents lists available at ScienceDirect

Vibrational Spectroscopy journal homepage: www.elsevier.com/locate/vibspec

Review

Raman spectroscopy: Recent advancements, techniques and applications Ruchita S. Das ∗ , Y.K. Agrawal Gujarat Forensic Sciences University, Institute of Research and Development, Sector 18 A, Gandhinagar 382007, Gujarat, India

a r t i c l e

i n f o

Article history: Received 14 March 2011 Received in revised form 12 July 2011 Accepted 9 August 2011 Available online 17 August 2011 Keywords: Vibrational spectroscopy Raman spectrum Samples Science Analysis Detection

a b s t r a c t Vibrational spectroscopy has proven itself to be a valuable contributor in the study of various fields of science, primarily due to the extraordinary versatility of sampling methods. Raman measurement gives the vibrational spectrum of the analyte, which can be treated as its “fingerprint,” allows easy interpretation and identification. Over the last years, there has been tremendous technical improvement in Raman spectroscopy, as overcome by the problems like fluorescence, poor sensitivity or reproducibility. This article reviews the recent advances in Raman spectroscopy and its new trend of applications ranging from ancient archaeology to advanced nanotechnology. It includes the aspects of Raman spectroscopic measurements to the analysis of various substances categorized into distinct application areas such as biotechnology, mineralogy, environmental monitoring, food and beverages, forensic science, medical and clinical chemistry, diagnostics, pharmaceutical, material science, surface analysis, etc. Advances in the instrumental design of Raman spectrometers coupled with newly developed sampling methodologies have also been described which enable trace level detection and satisfactory analysis. © 2011 Elsevier B.V. All rights reserved.

Contents 1. 2. 3.

4.

5.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Basic principles: Mechanism and Instrumentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Advanced Raman techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Surface-enhanced Raman spectroscopy (SERS) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Confocal Raman microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Coherent anti-Stokes Raman scattering (CARS) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. Resonance Raman spectroscopy (RRS) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5. Raman sensing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Forensic science . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Diagnostics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4. Materials science . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.1. Superconductors and Semiconductors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.2. Carbonaceous materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.3. Polymers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.4. Environmental materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.5. Crystalline study . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.6. Molecules and Molecular systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.7. Archaeological material . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5. Pharmaceuticals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6. Nanotechnology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

∗ Corresponding author. Tel.: +91 9328859853; fax: +91 7923256252. E-mail address: das [email protected] (R.S. Das). 0924-2031/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.vibspec.2011.08.003

164 164 164 164 165 166 167 167 167 168 169 169 171 171 171 171 171 172 172 172 172 174 174 174

164

R.S. Das, Y.K. Agrawal / Vibrational Spectroscopy 57 (2011) 163–176

1. Introduction In 1928, an Indian physicist Chandrashekhara Venkata Raman discovered the phenomena of inelastic scattering of light, known as the Raman effect. This explains the shift in wavelength of a small fraction of radiation scattered by molecules, having different frequency from that of the incident beam [1]. This shift in wavelength depends upon the chemical structure of the molecules responsible for scattering. Raman spectroscopy utilizes scattered light to gain knowledge about molecular vibrations which can provide information regarding the structure, symmetry, electronic environment and bonding of the molecule, thus permits the quantitative and qualitative analysis of the individual compounds [2]. This paper is dedicated to the techniques and various applications related to Raman spectroscopy in different fields of science, including a short introduction of its principle and instrumentation, which will help in better understanding of its analytical versatility. 2. Basic principles: Mechanism and Instrumentation Fig. 2. Instrumentation.

The irradiation of a molecule with a monochromatic light always results in two types of light scattering, elastic and inelastic. In elastic scattering, there occurs no change in photon frequency or without any change in its wavelength and energy. Conversely, the other is inelastic scattering which is accompanied by the shift in photon frequency due to excitation or deactivation of molecular vibrations in which either the photon may lose some amount of energy or gains energy [3]. Thus, three types of phenomena can occur [4] (Fig. 1). First, when light is incident on a molecule, it can interact with the molecule but the net exchange of energy (E) is zero, so the frequency of the scattered light is the same as that of the incident light (E = Eo ). This process is known as Rayleigh scattering. Second, the light can interact with the molecule and the net exchange of energy is the energy of one molecular vibration. If the interaction causes the light photon to gain vibrational energy from the molecule then the frequency of the scattered light will be higher than that of the incident light (E = Eo + Ev ), known as anti-Stokes Raman scattering. Third, if the interaction causes the molecule to gain energy from the photon then the frequency of the scattered light will be lower than that of the incident light (E = Eo − Ev ), this process is known as Stokes Raman scattering. A Raman spectrometer is composed of light source, monochromator, sample holder and detector. The factors which affect the analysis on Raman spectra may include high signal-to-noise ratio, instrument stability and sufficient resolution. The development of effective FT Raman spectrometers using NIR or red excitation lasers solved the problem of avoiding fluorescence that affects the Raman signals. On the other hand, the development of highly

sensitive detectors in conjunction with coupling of optical fibres and microscopes enhanced the capacity of analysis [5]. Two major technologies are used to collect the Raman spectra, Dispersive Raman spectroscopy and Fourier transform Raman spectroscopy, with difference in their laser sources and the way by which Raman scattering is detected and analysed (Fig. 2). Both these techniques have unique advantages and the method that best suit the sample should be preferred [6–8]. Several type of lasers can be used as the excitation source, like argon ion (488.0 and 514.5 nm), krypton ion (530.9 and 647.1 nm), He:Ne (632.8 nm), Nd:YAG (1064 nm and 532 nm) and diode laser (630 and 780 nm). Use of 1064 nm near-IR (NIR) excitation laser causes lower fluorescent effect than visible wavelength lasers [9]. In the most basic, a molecule is Raman active when there is a change in polarizability during the vibration. In addition, symmetry of a molecule is also one of the basic requirements for obtaining Raman spectra, as the symmetric stretches are more intense in Raman spectra. Functional groups such as -C–X (X = F, Cl, Br or I), –C–NO2 , –C–S–, –S–S–, –C C–, –C S–, –N N–, –S–H–, –CN, etc., exhibit more polarizability changes, give strong Raman signals [10]. Sometimes, Raman spectroscopy has been found coupled with many hyphenated analytical techniques like high-performance liquid chromatography [11], microchromatography [12], scanning tunneling microscopy [13] atomic force microscopy [14], etc.; give possibility for useful analysis at trace level studies. Thus, with the revolutionary developments in Raman instrumentation, it is now possible to acquire spectra more quickly on equipment which is affordable and easier to use than in the past. The following will summarise different techniques associated with the Raman spectroscopy and related applications.

3. Advanced Raman techniques 3.1. Surface-enhanced Raman spectroscopy (SERS)

Fig. 1. Mechanism of Raman scattering.

SERS is one of the most sensitive tools for the detection of adsorbate molecules on roughened metal surfaces which produces a large enhancement to the Raman scattering signal (Fig. 3). This enhancement effect was studied independently by Jeanmaire et al. [15] and Albrecht et al. [16] in 1977 and each proposed a different mechanism for the observed enhancement. Accordingly, two factors were assumed to be responsible for the enhancement of signals:

R.S. Das, Y.K. Agrawal / Vibrational Spectroscopy 57 (2011) 163–176

165

Fig. 3. Surface enhanced Raman scattering.

1. Electromagnetic enhancement (EM), associated with the roughened metal surface [15]. 2. Chemical enhancement (CHEM), due to the electronic coupling of molecules adsorbed on the roughened metal surfaces (or involves changes to the adsorbate electronic states due to chemisorption of the analyte or chemical bond formation between metal surface and molecules under observation) [16]. However, the electromagnetic effect is supposed to be more dominant and sometimes it is also called as “first-layer effect,” because it requires direct contact between analyte molecule and metal surface [10,17]. Both these factors can be well understood by the concept of surface plasmon which is located in the metals like Ag (silver) or Au (gold). When this plasmon oscillate perpendicular to the surface of the metal causes scattering, reflects the roughness of the surface which can either be the physical roughness or may be produced by some nano particles [18–20]. As the signals obtained by normal Raman scattering is usually very weak, therefore in order to get more detectable or increased signals, one prefers SERS technique. This technique can yield information on how molecules interact with surfaces, which allow detection of very low concentrations of analytes. This specially prepared metal surfaces like gold, silver and copper increase the intensity of the Raman signal up to 104 to 106 fold which enables faster and higher accuracy detection of biological and chemical samples [21]. The Raman signal enhancement is maximized when metal grains are smaller than the incident laser wavelength with optimized geometry [22]. Gold caps (nearly 50–400 nm in diameter) were supposed suitable for exploiting the tip or surface enhanced Raman scattering effects, since they assume the right size on nanometer scale with almost ideal hemispherical shape as the signal enhancement relies on the geometry of the metal particles [23]. Other concepts were also given for the enhancement of the signals, suggest that the enhancement effect is mainly due to the formation of nanojets, obtained when the laser focused on microsphere of appropriate diameter. This may lead to highly localized electromagnetic field causing enhancement [24]. SERS has been applied successfully for the study of Ni (nickel) and Pt (platinum) electrodes having different surface roughness, helps in obtaining good quality surface Raman signals from transition metals. Thus Raman spectroscopy will be further developed as a versatile mean for characterizing interfacial processes on rough surfaces in electrochemical and other surface environments for both fundamental and practical applications [25]. Quantitative analysis of a fungicide thiram has been done by using the substrate of silver (Ag+ ) nano-particles, prepared by radiolysis of Ag+ aqueous solution without addition of aggregating or stabilizing substances, as it might limit the spurious bands in Raman spectra. The detection thus obtained was at low concentration and without the interference of impurities of the medium [26]. Likewise, the Raman intensity of malachite green isothiocyanate (MGITC) adsorbed on gold (Au3+ ) surface was increased nearly up to 20 folds which explore the sensitivity of detection for adsorbed species on single crystal surface [24]. SERS sensor allows the feasibility of detection of volatile organic compounds (VOC), prepared by coating thiol with the substrate of SERS mounted on thermoelectric cooler. The purpose of this coating

Fig. 4. Confocal Raman microscope.

was to protect the substrate from degradation, to provide an internal calibration standard and to attract contaminants of interest. This can facilitate the detection of VOC such as chlorinated solvents, methyl t-butyl ether (MTBE) and aromatics with their characteristic SERS response [27]. Shell-isolated nanoparticle-enhanced Raman spectroscopy is described for the amplification of Raman signals by gold nanoparticles with an ultrathin silica or alumina shell. A monolayer of such nanoparticles is spread over the surface, later probed in order to obtain high quality Raman spectra. This can support the application of SERS in material and life sciences, as well as in the field of food safety, drugs, explosives and environment pollutants [28]. 3.2. Confocal Raman microscopy The first confocal scanning microscope was invented in 1955 by Marvin Minsky, in order to avoid thin-slicing of brain tissues [29]. In confocal Raman microscopy, laser light from the probe-head is made focused on the sample through microscope objective. The backscattered Raman signal is refocused onto a pinhole aperture that acts as a spatial filter. The filtered Raman signal then returns to the spectrometer where it is dispersed on a CCD (charge coupled device) camera to produce a spectrum [30] (Fig. 4). It must be remembered that the fluorescence background should be sufficiently low to affect weak Raman signals [31]. This technique possesses a number of applications by providing three-dimensional image of chemical composition with micrometer resolution and clear image quality. The three-dimensional imaging through turbid medium, high spatial resolution and rapid identification of micrometer sized biological specimens can be examined using a confocal Raman microscope [32–34]. It is a non-invasive method to obtain detailed information about the molecular composition of different tissues with high spatial resolution, provides optical section of the tissues without physical dissection. This technique can also combine with other Raman techniques to provide detailed information about molecular compositions of confocal images. This technique can also be combined with other with other Raman techniques to provide detailed information about the molecular composition in the confocal image.

166

R.S. Das, Y.K. Agrawal / Vibrational Spectroscopy 57 (2011) 163–176

The combination of confocal Raman microspectroscopy and confocal scanning laser microscopy (CSLM) has been presented as a novel non-invasive method to obtain information about molecular composition in relation to skin architecture. In addition to this, detail in vivo concentration profiles of water and natural moisturizing factors for stratum corneum has also been described which are directly related to the skin architecture. Thus, the arrangement of two different techniques can provide the basis for a wide range of applications in fundamental skin research, as well as in pharmacology, dermatology and cosmetics as well as for non-invasive analysis of blood analytes [35]. Confocal Raman microscopy has been found to be very effective in the analysis of the distribution of chemical moieties within polymeric coil coatings because it give strong Raman scattering which provides characteristic profiling of pigment. Such kind of analysis was performed with both dry and oil immersion objectives, and it was observed that the resolution gets affected by both the intrinsic as well as extrinsic factors of the objective lens. It has also been observed that the use of an oil immersion objective improves depth resolution and minimised the refractive effect which favours multiple analysis of samples. However, oil may contaminate the surface coating. On the contrary, the dry method yields the lowest depth resolution and allows non-destructive analysis [36]. Likewise, this technique can be used to measure the spatial and temporal evolution of a drying coating film of paint. Comparison between confocal Raman microscopy and nuclear magnetic resonance (NMR) technique has done for the examination of two alkyd paint coatings, one was organic solvent based and another was water based. The NMR and confocal Raman microscopy have shown good similarities in profiles of paint films which were obtained by comparing the profiles of disappearance of the double bonds of the unsaturated fatty acid side chains of alkyd molecules. Thus, the technique can be used to analyse spatial variations of paint properties within a thin film [37]. Moreover, this technique offers as an important alternative to conventional spectroscopic techniques that provide elemental/atomic composition of hazardous components in cosmetic products like eyeliner. Raman spectra of such cosmetic samples have been measured between 150 and 3000 cm−1 at room temperature, showed the presence of lead (II) sulphide (PbS) which is a weak Raman scatterer at room temperature and is therefore susceptible to laser-induced degradation when intensely irradiated. The use of confocal Raman microscopy exhibits superior rejection of fluorescence and thus the stray light is rejected, so only the desired Raman signal is passed to detector which helps in proper analysis of the sample [38]. 3.3. Coherent anti-Stokes Raman scattering (CARS) CARS was first reported by P.D. Maker and R.W. Terhune in 1965 and they called it ‘three wave mixing experiments’ [39]. But the name coherent anti-Stokes Raman Scattering was assigned by Begley et al. in 1974 [40]. This technique allows vibrational imaging with high sensitivity, high spectral resolution and threedimensional sectioning capabilities. It is a nonlinear diagnostic technique that relies on inducing Raman coherence in the target molecule using two lasers, probed by a third laser which generates a coherent signal in the phase-matching direction at a blue-shifted frequency. This phenomenon can be explained on the basis of some equations. Incident radiation consists of two overlapping coherent monochromatic beams of frequencies (ωp1 ) and (ωs ) with (ωp1 > ωs ). When (ωp1 − ωs = ωvib ) (molecular vibration frequency), observed normal Raman scattering, but when probed by a third laser (ωp2 ) generates a coherent laser-like signal in the phase-matching direction, produces strong anti-Stokes signal [ωas = (ωp1 + ωp2 ) − ωs ] with high spatial resolution [41] (Fig. 5).

Fig. 5. Coherent anti-Stokes Raman scattering.

This has been proven as a valuable tool for measuring temperature and major species concentration in reacting flows [42]. An overview about the advances made over the last few decades in the development and applications of nanosecond, picosecond, and femtosecond laser-based CARS spectroscopy in gas-phase reacting flows has been demonstrated well [43]. CARS permits probing of different molecular bonds, in various biological systems. Examples include, imaging of C–H stretching vibration present in the lipid bilayer of the cell membranes [44,45], P–O vibration (at 1090 cm−1 ) in chromosomes and more recently, imaging of live tissues [46]. Additionally, CARS microscopy has already been used for imaging a number of delicate biological samples and processes. It provides excellent sensitivity, high spatial resolution and inherent chemical specificity, gives vibrational and spectral information at low laser power also [47]. The detected signal in a CARS microscope is a result of coherent superposition of the signal waves generated in focus and can be used as an imaging tool for chemical mapping of biological cells and tissues with high sensitivity [48]. It is a key technique in non-invasive cellular imaging as it permits imaging of intrinsic biomolecules without chemical labelling or without the complications of photobleaching [44,49]. It also has significant potential for biomedical applications, since chemical imaging of live skin tissues has been achieved [50]. The nanosecond laser based CARS technique is used in the measurement of temperature and multiple species concentration detection. On other hand, dual-pump CARS, triple-pump CARS, dual-broadband CARS and dual-pump dual-broadband CARS were applied mainly to detect multiple species in one experimental setup. In picosecond CARS, the probe beam is temporally delayed with respect to the pump and Stokes beams to suppress the nonresonant background. Additionally, picosecond pulse laser works above 800 nm, which avoids multiphoton damage of specimen and allows deep penetration in thick samples. While temperature and concentration measurements of diatomic and triatomic gas phase molecules can be performed using a single femtosecond laser beam. Thus, by tuning into characteristic vibrational resonance in samples, higher scanning speed and fast image acquisition time makes CARS the technique of choice [51]. Furthermore, new developments in Electronic Resonance Enhanced CARS (ERE-CARS) spectroscopy have paved the way for detection of minor species such as NO [52,53], C2 H2 [54], etc. One more approach is multiplex CARS (M-CARS) microscopy that has been used to characterize neat liquids, polymer blends, domains in lipid droplets and bacterial spores. In this technique a spectrally broad femtosecond pulse is used for Stokes transition which can provide sensitivity over 300–1500 cm−1 of spectral bandwidth at each spatial pixel [55–57].

R.S. Das, Y.K. Agrawal / Vibrational Spectroscopy 57 (2011) 163–176

With the advent of time there has been remarkable development in the CARS technique. Recently with the use of pulse train, the time required for obtaining broadband CARS signals is reduced to one third as compared to previous studies without using pulse trains. The pulse train was created by shaping optical pulses with a pulse shaper and their waveforms were measured by a crosscorrelation frequency-resolved optical gating method. Pulse train was generated from a photonic crystal fibre (PCF) with different centre wavelengths and used as Stokes optical pulses in CARS which can be used to measure broadband CARS signals. Such kind of study has been done to create pulse train by a pulse shaper to generate multiple soliton pulses with different centre wavelengths and different delay times from a PCF. This helped in obtaining five soliton pulses necessary for broadband CARS spectroscopy; generated using only two phase patterns and the time required for the measurement was reduced [58]. These pulses can also be used as Stokes pulses in broadband CARS (B-CARS) measurements. The B-CARS imaging technique is used to measure spectral signatures of individual cells at least fivefold faster than spontaneous Raman microspectroscopy and can be used to generate maps of biochemical species in cell. It offers the same inherent chemical contrast as spontaneous Raman signal but with increased acquisition rate. B-CARS imaging technique has been studied to obtain complete resonant vibrational spectra of a single cell with 50-ms individual pixel dwell times. This acquisition time is at least fivefold faster than spontaneous confocal Raman microspectroscopy of cells at similar spatial resolution. The spectra thus obtained were in the range of 600–3200 cm−1 . This covers both fingerprint and CH-stretch regions that provide both morphological as well as rich biochemical information. This improved spectral range and signal intensity opens the door for more widespread use of vibrational spectroscopic imaging in biology and clinical diagnostics [59]. Furthermore, the non-resonant background can easily overwhelm less intense signals from other molecular vibrations which contain important information about the vibrational energy levels of molecule or sample specimen. This non-resonant scattered light can interfere coherently with the resonant signal which may distort the band shapes. Many approaches have been established for reducing the non-resonant background, like epi-detection, temporal delay of probe with respect to the pump and Stokes fields [60], and polarization control of the CARS signal [61]. One more technique has been given for interferometric suppression of the non-resonant background based on interferometric mixing of the CARS signal with a local oscillator, generated within the sample. In this technique liquid benzonitrile was selected as a model system for removing the non-resonant signals, which gives prominent peak throughout the vibrational spectrum, provides good sensitivity and chemical species identification [62].

3.4. Resonance Raman spectroscopy (RRS) Resonance Raman spectra obtained when the energy of photon of an exciting laser beam matches approximately with the energy require for electronic transition (Fig. 1). The vibrations which cause enhancement in Raman bands fall into two or three general classes. The most common class is Franck-Condon enhancement, in which a component of normal coordinate of the vibration is in the direction of molecule which expands during an electronic excitation. The more the molecule expands along the axis (when it absorbs light), the larger enhancement factor occurs. Vibrations which couple two electronic excited states produce enhanced resonance signal. This mechanism is called vibronic enhancement. In both cases enhancement factors roughly follow the intensities of the absorption spectrum [63].

167

Fig. 6. Fibre Raman sensor.

Resonance Raman scattering has been extensively exploited in the analysis of various chromophoric biological samples like enzymes, various parts of biomolecules and protective pigments of photosynthetic organisms. Similarly, it can also be used for obtaining high quality pre-resonance Raman spectra of bacteriochlorophyll chromophores in photosynthetic proteins from purple bacteria without sample degradation [64]. Discussion and comparison of RRS and normal Raman spectroscopy has been reviewed well. This has explored the utility of RRS as a reliable and versatile analytical technique, as in the analyses of carotenoids in biological matrices, analyses of pigments and dyes in forensic investigations, in bioanalytical and life sciences, study of metalloproteins, etc. Furthermore, this technique is made to couple with other separation (chromatographic and electrophoretic) and detection (UV) techniques [65]. Sometimes the extreme enhancement factors can be realized by combining RRS and SERS to SERRS as SE(R)RS (Surface Enhanced Resonance Raman Spectroscopy), where the laser excitation wavelength not only coincide with the plasmon band, but also with the absorption wavelength of the analyte [66]. 3.5. Raman sensing Advances in low-cost fibre optics and miniaturized detectors led to rapid increase in the use of Raman spectroscopy with greatest advantage of remote sensing. This technique is basically associated with optical fibres to transport Raman signals by collecting the scattered photons, simultaneously filtering out Rayleigh scattering (Fig. 6). Fibre system includes single-fibre and multiple fibres in which the laser excitation is transmitted along one fibre and the scattered radiations are transmitted to the detector along different fibres [4]. Thus, gives the feasibility of online monitoring of process streams and hazardous reactions [67]. Background discrimination, sample volume, and probe sensitivity has been investigated as a function of laser source and fibre design [68,69]. Other studies have focused on coupling efficiency, damage threshold, and sensitivity for UV Raman fibre probes in the presence of adsorbing materials [70]. Fibre-optic Raman can be used to determine the amount of organic vapour that partitions into solid-phase extraction medium [71]. Remote micro imaging has also been accomplished using a coherent optical microfiber array as a probe [72]. 4. Applications Raman spectroscopy is becoming increasingly important in a broad range of scientific disciplines. Brief discussion of these applications in various field of science have been discussed below.

168

R.S. Das, Y.K. Agrawal / Vibrational Spectroscopy 57 (2011) 163–176

4.1. Forensic science The recent technological advancements in Raman spectrometer have provided a reason for exploring its use in forensic science. Analysis of fibres, explosives [73], drugs [74], paints [75,76], inorganic fillers, lipsticks [77] and other materials have been done successively by using Raman techniques. Confocal Raman Microscopy is used for the examination of various kinds of biological fluids like dry traces of semen, vaginal fluid, sweat, saliva and blood. These fluids can be differentiated from one another by visual comparison of their Raman spectra, as each body fluid has its own composition that can give specific Raman signal. An even dry trace of human and canine semen exhibits different Raman signatures. This can help in better interpretation of crime scene exhibit especially body fluids, as these analyses are non-destructive in nature and offers the possibility of further testing. This study was done on only one sample of each body fluid and did not taken into account any variations that may occur between different donors of the same fluid [78]. However, in another study heterogeneity was obtained within a sample as well as among multiple donors for human semen samples. NIR is used to measure spectra of purely dried human semen samples from multiple donors. Statistical analysis of spectra obtained from samples showed that the dry semen is heterogeneous and its Raman spectra can be presented as a linear combination of a fluorescent background with three different spectral components varies with donor. Thus, no single spectrum could effectively represent an experimental Raman spectrum of dry semen in a quantitative way. However, combination of three spectral components can be considered to be a spectroscopic signature for semen during forensic analysis [79]. Examination of ink is very important aspect for the investigation of questioned documents. Sometimes it is necessary to apply chemical methods that normally cause partial destruction of the examined material, but Raman spectroscopy provides direct identification of inks on documents for determination of sequence of the handwritten lines. The investigations are mostly often aimed at authenticating the document or at determining its age or origin. This could be possible due to some characteristics of the ink such as luminescence in visible or in infrared light. A good discrimination between inks has been obtained using Raman spectroscopy, possibly used to differentiate between inks of the same colour [80]. Fibres are also one of the commonly encountered evidence during forensic investigation especially black/grey and blue cotton type fibres. The fibres are generally coloured by reactive dyes, as dyes can able to form non-polar bonds with fibres, which can produce difficulty in extraction during analysis. Raman spectroscopy measures the vibrational states of non-polar bonds through the use of high intensity lasers and can be used as analytical tool for fibre examination [81,82]. Different aspects of fibre analysis were studied via Raman Spectroscopy, observed variations due to the method of mounting the dye particles, spectral degradation, fluorescence, resonance enhancement and laser wavelengths. Results have shown good spectra of dyed cotton fibres without interference from cotton substrate, thereby provides molecular information about the dye used in the fibre [83]. Similarly, Raman measurements can be performed directly on mounted fibres without encountering much interference from the mounting medium. Forensic cases were described where the comparison of Raman spectra of reference fibres and suspect fibres has been discussed in detail. The results suggested Raman spectroscopy as a powerful tool for Forensic fibre examination (Fig. 7) [84]. Some studies have also explored the use of resonance, non-resonance, and surface-enhanced Raman spectroscopy for forensic fibre examination, including both dyed and

Fig. 7. Baseline corrected Raman spectra of (a) the brown cotton fibres found on the suspect, and (b) those found on the victim.

non-dyed textile fibres, as it can provide detailed vibrational profiles which gives characteristic “signature” to dyes [85]. Raman spectroscopy has been proved as a good complementary method for detection of drugs of abuse in fingerprints. This kind of study was done on five drugs of abuse that is codeine phosphate, cocaine hydrochloride, amphetamine sulphate, barbital and nitrazepam. Detection of these drugs was successfully achieved and clearly distinguished using their Raman spectra obtained from the substances in cyanoacrylate-fumed fingerprints (polymer is deposited on the fingerprint material for enhancing the visibility). Interestingly, it was observed that the spectra obtained from the fingerprints were of a similar quality to the spectra obtained from the substances under normal sampling conditions [86]. Likewise, quantitative determination of caffeine in different energy drinks has been achieved by FT-Raman spectroscopy, provides fast and alternative mean to chromatographic method with higher sampling frequency. In order to quantify caffeine, spectra were obtained directly between 3500 and 70 cm−1 with Raman bands between 573 and 542 cm−1 (corrected using a baseline between 580 and 540 cm−1 ), obtained limit of detection as 18 mg/l [87]. Raman spectroscopy can also offer a good flexibility in the analysis of hazardous environmental samples. Detection of cyclotrimethylenetrinitramine (RDX) is achieved by SERS, extending its broad application areas in the examination of other explosives also. Analysis of RDX was done with gold (Au) nanoparticles (90–100 nm in diameter) as SERS substrates, with the detection level of 0.15 mg/l from contaminated groundwater sample. Thus, it can be potentially used as a valuable tool for rapid screening and characterization of energetics in the environment, such as tri nitro toluene (TNT), perchlorate, pertechnetate, and uranium in groundwater at low concentrations [88]. SERS has also been used to detect and distinguish explosives in the solution using azo dyes. These dyes contain electron-donating moieties give efficient diazo coupling and have a strong silver complexing group to attach the product molecule to SERS substrate, allows detection up to nM (nano molar) concentration [89]. Thus, it can explore the studies related to environmental monitoring, security and biological materials like nuclear waste testing [90]. Raman has also been proved as a powerful tool in the analysis of adulterated fuel samples like diesel or biodiesel blends with vegetable oil, used in remote quality control. Calibration models based on multivariate analysis such as principal component regression and partial least square regression in combination to vibrational spectroscopy has been applied for examining the adulteration with complete accuracy; especially with the help of appropriate choice of spectral regions [91].

R.S. Das, Y.K. Agrawal / Vibrational Spectroscopy 57 (2011) 163–176

4.2. Biology The greatest advantage of using Raman spectroscopy in bioanalysis is the wealth of information contained in each spectrum. Even Raman micro-spectroscopy can provide useful biochemical information regarding live cells, without the need of fixatives, markers or stains [92]. This can be related to the interactions with toxic agents or drugs, disease, cell death and differentiation. Raman spectrum of a cell can produce “fingerprint” of its biochemical composition so, if any toxic agent causes biochemical changes it appears in the Raman spectra [93]. It has been proved that the Raman spectra of biological macromolecules arise from the molecular vibration of either the backbone chains or the side chains. The wavenumbers of the Raman bands lie in a region between 200 and 3000 cm−1 . Even conformation or secondary structure can also be determined by analysing Amide I and Amide III vibrations for polypeptides and protein backbone chain. Similarly, in polynueleotides and nucleic acids the wavenumbers of phosphate diester stretch of phosphate furanose chain varies between 814 cm−1 for A conformation and 790 cm−1 for B conformation [94]. Hemoglobin-oxygen saturation in living tissue is determined by measuring the ratio of intensities of resonance Raman bands aroused from oxygenated and de-oxygenated hemoglobin [95]. Similarly, CARS also provides a non-invasive determination of the blood oxygenation (oxygenation state of hemoglobin) in individual vessel inside bulk tissues [96]. It can also be used for detection of bacterial spores by obtaining the molecular specific signals, as a marker molecule (dipicolinic acid) for bacterial spores [97]. In addition, CARS has provided a non-destructive analysis of embryonic stem cells within a growing culture. It has also given specific backbone vibrations for DNA (788 cm−1 ), RNA (811 cm−1 ) and proteins [98]. Even, Raman spectra can provide information about the structure, function and kinetics of protein by identification of its vibrational bands. The high sensitivity of deep UV resonance Raman spectroscopy makes it a valuable tool for studying biological systems under different physiological conditions [99–102]. It is also possible to determine the metabolic activity of cell mitochondria. The Raman band at 1602 cm−1 reflects respiratory activity of mitochondria in living cells and it was observed that the intensity of the band decreases with time after addition of sodium azide, act as a respiratory inhibitor [103]. Study of these biomolecules with Raman spectroscopy helps in evaluation of the quality of natural food products, as it provides structural information about the change in proteins, water and lipids of muscle food which occurs during deterioration. As compared to the traditional techniques like protein solubility, viscosity, water holding capacity, instrumental texture methods and peroxide value calculation which were used for the determination of quality of muscle foods were destructive in nature. In contrast, Raman spectroscopy provides valuable and non-destructive analysis of samples [104]. Due to the extraordinary versatility of sampling methods vibrational spectroscopy has proven itself to be a valuable contributor in the study of biophysical interfaces, which can help in obtaining the spectra of oriented monolayer films. Advances in the Raman techniques coupled with newly developed sampling methodologies have enabled to probe ever-smaller and ever-thinner samples. This will allow the examination and identification of unenhanced bio-membrane monolayer spectra, by giving molecular structure of membrane [105]. Advances in instrumentation have increased the performance of Raman instruments in bioanalysis. The specificity of the technique makes it an attractive tool for biomedical analysis which can provides information applicable to almost any biomolecule, that too without labelling [106]. It can also become a useful tool in the early detection of cells exposed to human papillomavirus (HPV) [107]. Likewise, Raman spectra of other viruses have also been stud-

169

ied in an effort to elucidate the mechanism of replication in the host organism where the side chains and secondary structure of proteins were clearly observed [108]. Analysis of individual bacteria, complex mixture of spores and vegetative cells can be done by Raman chemical imaging (RCI). One of the major advantages of this technique is that it resides primarily during analysis of samples in complex backgrounds without the need of physical isolation or purification of the sample [109]. A study of interactions between lysozyme and whey protein indicates the presence of hydrophobic interactions. This can be evidenced by intensification of spectral bands assigned to CH and CH2 bending vibrations which arose with the changes in disulfide vibrational frequency and through lowering in R-helix and alpha-sheet contents [110]. The structure of lipid-cholesterol vesicles (as a function of pressure) have been investigated using Raman wavenumbers, band shapes and splittings in the lipid bilayer [111]. Likewise, Filipin (which binds specifically to cholesterol) is used as an extrinsic Raman probe molecule for determination of cholesterol distributions within rat eye lenses [112]. Intake of glucose into the cell could be determined by attaching quinoline red dye with cell membranes. Quinoline was used as labelling agent to bind with the analyte of interest. Thus, results in increasing Raman cross-section of weak scatterers because peak position of dye spectrum changes when it binds to the cell. Moreover, when the cell membrane gets energized and glucose is transported into cell there occurs with change in the intensity of the spectra. Hence, use of Raman labels can be applied for studies at molecular level [100]. It was believed that the physical properties of nails changes because of change in keratin structure. Thus, examination of change in molecular structure of intact moisten nails has been studied through NIR-FT-Raman spectroscopy. The results obtained revealed that nails have water holding capacity, which causes increase in intensity ratio of the X(OH)/X(CH2 ) bands. This imply about the water–protein interaction which leads to change in protein geometry [113]. The Interaction of calf-thymus DNA with aspirin was investigated by Fourier transform Infrared (FTIR) and laser Raman difference Spectroscopy, provided information about the drug binding sites, sequence preference and changes in secondary structure of DNA. In addition to this, structural variations of aspirin-DNA complexes in aqueous solution have also been determined. Spectroscopic evidence showed that low aspirin concentration was helpful for drug-DNA interaction which is mainly through the backbone PO2 groups and the AT base pairs, obtained phosphate vibration at 1227 cm−1 and A-T bands at 1663 and 1609 cm−1 , with no major helix destabilisation [114]. Macroscopic resolution allows examination of the chemistry of individual cells by mapping their images. These images can contain full spectral information at each pixel so that the distribution of components within the cell can be visualized based on their Raman signature. This is extremely valuable to researchers as biochemical changes can be observed during a cell’s life cycle or when a cell is damaged or cancerous. Using confocal Raman microscopy, the changes in a variety of cells, including bacteria and eukaryotes can be monitored over time and comparison between healthy and diseased tissue states can be done easily [115]. 4.3. Diagnostics Raman spectroscopy has widely showed its utility as a diagnostic technique, offering various advantages for analysis of variety of biomedical materials. NIR provides a rapid and non-destructive analysis to assess the thyroid stimulating Hormone (TSH) in blood, given good reliability to the experiment [116]. FT Raman spectroscopy is used for examining and characterising the outermost layer of the human skin, stratum corneum provided valuable data related to the nature of healthy and diseased skin, transdermal

170

R.S. Das, Y.K. Agrawal / Vibrational Spectroscopy 57 (2011) 163–176

Fig. 9. Raman spectra of stomach mucosal tissues: (a) mean spectra of (—) normal and (- - -) malignant tissues.

Fig. 8. Raman spectra of (a) non-pathologic, (b) atheromatous and (c) calcified artery tissues.

drugs, pollutant permeation, mechanism action of penetration enhancers, etc. [117]. Histochemical analysis of biological tissues have been reviewed, explores the advantages of Raman spectroscopy in endoscopic imaging and quantisation of biochemical constituents during clinical studies. This involves disease diagnosis which involves analysis of tissue proteins, lens, cornea, blood constituents, biological stones, hard tissues, arterial disease etc. [31]. Cancer studies in gynaecological tissues, soft tissues, breast, colon, bladder and brain has also been summarized [118]. A method had been reported on the use of near-infrared (NIR) Fourier transform (FT) Raman spectroscopy to analyse normal human epidermal keratinocytes before and after malignant transformation. Raman spectral differences between isolated DNA of immortalized and malignant transformed cell indicates some specific alteration during malignant transformation. Thus this make possible to observe progressive structural changes in either DNA or protein components of cancerous tissues. These changes in Raman spectrum can build area of interest in the evaluation of newer anticancer agents [119]. Raman spectra of various tissues were used to differentiate between normal and diseased ones, as well as different chemical states of organs and cells. Even white and grey matter of the brain can be easily differentiated with micrometer resolution in spite of being complex and heterogeneous. This can furnish an opening in the diagnosis of brain related disorders like Parkinson’s and Alzheimer’s disease by examining the differences between healthy and diseased tissues [120]. Studies have also revealed the facts, that the Raman band at 1628 cm−1 indicates the presence of b-amylose proteins obtained from post-mortem paraffinated brain tissues, thought to be responsible for the onset of Alzheimer’s disease [121]. In vivo diagnosis of atherosclerosis in human arteries has been done by using Near-infrared Raman spectroscopy (NIRS). It develops with the deposition of calcified mineral layers which decreases the elasticity of the artery. Each one of these biochemical depositions give a well distinct Raman spectrum which provides the information to differentiate between calcified artery tissue, atherosclerotic and non-pathological calcified mineral layer (Fig. 8). Thus, it is possible to obtain satisfactory tissue classification for in vivo clinical applications [122]. A similar kind of study has been

performed in vivo using optical fibre technology. The Raman spectra were collected from normal and atherosclerotic coronary artery samples in different stages of disease progression from explanted transplant recipient hearts. This provides information about the morphologic composition of intact human coronary artery without the need of excision and microscopic examination [123]. Moreover, it can also provide a new and important means for analyzing the chemical composition (lipid and calcium salt content) of the arterial wall, which may help in selecting appropriate treatment [124]. Similarly, Raman technique can be applied to quantitative analysis of cholesterol and cholesteryl esters in human atherosclerotic lesions [125]. Raman techniques can be used as an alternative tool to cancer detection, as it has been used for determination of the nature of tumour (as benign and malignant tumour has different spectra) and examination of paraffin-embedded skin biopsies. Raman spectra provide wealthy information about the chemical composition of biological samples by admitting Raman signature to every constituent. This can also help in building proper strategies during treatment of cancer by targeting the specific site. [126–131]. Moreover, the feasibility of discriminating normal and malignant stomach mucosal tissues has been studied. The mean Raman spectra of normal and malignant tissues exhibit significant differences in amide I, -CH2 , and amide III regions. The major spectral variation of normal tissue with respect to malignant tissue was observed due to weak amide I, slightly red shifted -CH2 , an intense band at 1303 cm−1 and hump at 1276 cm−1 . It has also been observed that the normal tissue has high lipid content, while malignant tissue is rich in protein, which can help in differentiation (Fig. 9) [132]. The Raman spectroscopy method was also validated for the discrimination of normal and malignant tissues in cervical cancers. It was observed that normal cervix tissues were characterized by strong, broad amide I, broader amide III and strong peaks at 853 and 938 cm−1 , which can be attributed to structural proteins such as collagen. While the malignant tissue spectra with respect to normal tissue was relatively weaker and sharper amide I, minor red shift in -CH2 and sharper amide III, indicated the presence of Deoxyribonucleic acid (DNA), lipids and non-collagenous proteins [133]. It has also shown good results during the evaluation of renal tumours, provided complete and accurate information of differentiation between normal and tumoural renal tissue, low-grade and high-grade renal tumours, and histologic subtype of renal cell carcinoma [134]. Even kidney stones which form due to the reduction of cystine to cysteine, can be well diagnosed with the help of Raman instruments [135]. Analysis of calcium oxalate-type kidney stones is also possible using Raman techniques [136]. Thus, Raman spectroscopy can provide important information about the molecular and supramolecular structure of living tissues including hard tis-

R.S. Das, Y.K. Agrawal / Vibrational Spectroscopy 57 (2011) 163–176

sues like bone [137–139]. SERS has been proved useful in detecting the specific Anthrax biomarker, protective antigen (PA), after using a short 16-amino acid peptide chain in place of an antibody as it has been observed that the peptides are more stable than antibodies under various biological conditions and are easily synthesized for a specific target. Thus, the peptides were made conjugated on gold nano-particles labelled with a Raman tag (dithiobis-Nsuccinimidyl propionate, DTSP) to specifically recognize target biomarkers against biological and environmental pathogens [141]. 4.4. Materials science Among different vibrational techniques, Raman spectroscopy appears as privileged method for the analysis in material science. This field provides a varied and challenging array of samples ranging from Super-conductors to Archaeological materials. Being as indispensable characterization procedure, Raman technique is readily applicable to any material system. Thus, this section explores the tremendous possibilities of Raman signature in the area of material science [140]. 4.4.1. Superconductors and Semiconductors Semiconductors are materials that have intermediate electrical conductivity between that of metals (conductors) and insulators (non-conductors). Raman measurements have been reported effective for structural characterization and investigation of properties of semiconductor [141,142]. The effect of reduced dimensionality on the shape, size and position of the first order phonon bands of semiconductors can also be described through their Raman spectrum. The same has been used for the study of non-destructive measurement stress, crystal lattice disorder, phase-separation of supersaturated solid solutions and homogeneity of materials [143]. Likewise, superconducting materials have also been widely studied through Raman spectroscopy [144,145]. Study of semiconductors such as nanowires and nanocones revealed that the Raman enhancements are diameter, wavelength and polarization dependent, which can play an important role for engineered photonic and sensing applications [146]. 4.4.2. Carbonaceous materials Natural carbon exhibits its distinct properties in two forms; the first is diamond which is an insulator having the greatest mechanical strength and the second is graphite, a brittle material that conducts electricity at room temperature. While recently synthesized carbon form that is carbon nanotube (CNT), contains both these properties (strength and conductivity). Raman scattering is one of the prime techniques for the characterization of carbon nanotube, which includes study related to their vibrational, electronic and optical properties. Even metallic and semiconducting tubes can be distinguished from their high-energy Raman spectra that can also help in their characterization [147,148]. A sharp Raman band for the cubical structure of diamond was also observed at 1332 cm−1 [149]. Thus, it is also possible to study the defective structures of diamonds and other similar materials. Likewise, spectra of carbon nanotubes and fullerene type carbonaceous materials can provide various information regarding their structure and properties [150]. Raman spectra of nine different carbonaceous materials of astronomical relevance have also been determined. In which eight out of nine samples showed two main bands falling around 1600 and 1350 cm−1 . This implies that all the materials are composed of randomly oriented structural units varying from 50 to 80 A˚ in size. Some bands were also observed between 800 and 1700 cm−1 which is of C C or CHn (n = 1, 2, 3) functional groups, provides information about astronomically relevant materials [151]. Laser-Raman

171

spectra of carbonized coals, carbons, and graphite have also been observed near 1590 and 1360 cm−1 [152]. 4.4.3. Polymers Characterization of the structure, environment and dynamics of polymeric materials has also been obtained through Raman spectroscopy [153]. Study of the chemical composition and structure of polymer materials such as Kevlar, involves illuminating the surface with an incident laser light. This helped in studying its characteristics scattering including the effect of incident wavelength, polarization and laser power. This allows the development of non-destructive evaluation technologies based on the interaction of laser light with Kevlar for detection of in situ deterioration [154]. Raman characterization of plastic explosive materials (trinitro toluene and di-nitro toluene) have been reported by SERS using 3D alumina membrane with cylindrical nanopores which allows molecular level detection of trace amount of explosives and other relevant chemical compounds [155]. Raman spectra and cross sections of spider silk fibres produced by two different species were measured as one of the characteristic feature to differentiate between species. Comparison of polymer fibres formed by electro-spinning (technique which produces continuous fibres from a polymer solution through the action of an electrical field) from mixture of polymer solutions produced homogeneous Raman spectra [156]. The spectra can be obtained even in cases of immiscible polymers through confocal Raman microscopy for analyzing the structure of electrospun fibres [31]. 4.4.4. Environmental materials Hazardous materials in trace amount can find their way in the water system causes very severe effects. Resonance Raman spectroscopy can be applied for on-line monitoring of NO2 − /NO3 − in wastewater (using such excitation radiation practically eliminates fluorescence from other species present in the wastewater) with the detection limits below 200 ppb for both the analytes [157]. SERS can be applied to detect contaminants at femtomolar concentrations over practical time scales in highly controlled environment. Qualitative identification of TNT (2,4,6-trinitrotoluene), fullerenes and numerous other compounds often rely on detection of a specific SERS spectrum to infer the presence of contaminants in environmental samples [158]. Even uranium can be detected in aqueous media by SERS technique. A method has been developed for the rapid screening of uranium in environmental samples, using a new SERS substrate, based on (aminomethyl)phosphonic acid (APA)modified gold nanoparticles. The intensity of uranyl band was observed at 830 cm−1 which is proportional to the concentration of uranium in solution, achieved detection limit close to 8 × 10−7 M with good reproducibility. This allows the detection of uranium in a highly contaminated groundwater with dissolved salts [159]. Similarly, the same technique has been reported for the detection and monitoring of perchlorate (ClO4 − ) in groundwater and surface water with detection limit of 10–100 ␮g/L [160]. Laser spectroscopy with tunable IR laser source is used as sensitive techniques for the detection of various molecules in atmosphere. This was made possible by photoacoustic Raman spectroscopy (PARS), where the thermal relaxation of molecules from the upper vibrational state was detected as an acoustic wave. Accompanied with two visible lasers with frequencies (ωp ) and (ωs ) were used instead of an IR laser, and the frequency difference (ωp − ωs ) was tuned equal to the Raman shift frequency (ωR ) [161]. Likewise, nonlinear Raman spectroscopy can be applied for monitoring various kinds of factories, chemical plants, gas pipelines, any leakages of inflammable gases such as H2 , CH4 , and H2 S. This gives the possibility to excite the vibrational level of gas molecules in the IR region without the use of tunable IR laser [162]. Optical detection methods in combination with Raman spec-

172

R.S. Das, Y.K. Agrawal / Vibrational Spectroscopy 57 (2011) 163–176

troscopy, fluorescence spectroscopy and digital imaging have been experimented to detect the trace levels of harmful biological agents like Bacillus anthracis, Yersinia pestis, and Francisella tularensis by their Raman signatures [163]. 4.4.5. Crystalline study Micro-Raman spectroscopy has been used to study the spatial variation of internal stress in synthetic diamond, including single crystal and polycrystalline specimens. It also provides direct information on the state of stress and was successfully employed to examine the stress distribution around cracks produced by indentation in the diamond [164]. Coherent Raman measurements of nanoshock in solids have been reported for the determination of shock-induced deformation in materials [165]. Specimens of sintered TiO2 ceramics were investigated on the basis of IR Raman signals for detecting the impure grains of Al2 O3 (including composition, size and shape variations). Micro-Raman configuration can offer the ability to monitor changes in chemical composition and morphology at microscale which helps in investigation and characterization of the materials. Moreover, this technique has the ability to recognise oxidizing and reducing production conditions, potentially save costs and optimise raw material consumption [166]. Temperature-dependent characteristic spectra of a BSO crystal were studied by Raman spectroscopy. It has been observed that the vibration mode of the longest bond Bi-O (1) in crystal shifts from 542 to 512 cm−1 when temperature was increased (from room temperature to 1123 K). In addition, the 58 cm−1 mode of Bi atoms (in crystal lattice) decreases rapidly when the temperature is higher than 873 K, indicates the breaking of crystal at high temperature [167]. Study of Raman spectra of crystalline domoic acid (DA) confirms its existence in zwitterionic form. DA is a natural occurring neurotoxin present in the marine ecosystem. The variations in the spectra were also observed attributed by hydration, the degree of protonation and crystallinity [168]. 4.4.6. Molecules and Molecular systems By plotting the excitation profile of Raman bands can help in the determination of coupling of electronic and vibrational motions of the molecule. In addition, structural changes results from a change in the electronic state of the scattering species and multiple metal–metal (MM) bonding can also be studied. Unique molecular information regarding oxygen–nitrogen, dinitrogen bridged complexes, and linear chain halogen-bridged complexes have also been examined successfully [169]. Apart from these applications it was also used to examine the solubility mechanism of fluorine in depolymerized silicate liquids, quenched glasses in CaO–CaFr–SiO system as an evidence for concomitant polymerization of the liquid [170]. Moreover, Raman scattering is a very sensitive technique to probe local atomic environments. Indeed, the properties of the vibrational modes are basically determined by the mass, bond type and symmetry of constituting atoms in the elemental unit [171]. Laser Raman spectroscopy holds great promise for trace level detection of surface planetary minerals especially oxy-anionic mineral such as silicates, carbonates, sulphates and phosphates [172]. A wide range of coloured main group metals, transition metal co-ordination and transition metal organometallic complexes has been studied by FT-Raman spectroscopy which gives good quality spectra in less time. This suggests its utility as a routine spectroscopic tool for inorganic as well as organic research and teaching laboratories [173]. The structure of supercritical H2 O has also been studied through Raman spectroscopy [174]. 4.4.7. Archaeological material Analysis of archaeological objects by Raman spectroscopy is a rapidly developing technique, provides non-destructive examination and investigation of such invaluable objects. Non-destructive

analysis of ancient skin tissue samples has been extensively studied by Raman spectroscopy. Partially desiccated or wet samples of skin were analysed when subjected to varying degrees of degradation (depending on the burial environment). Comparisons of Raman spectra between healthy and diseased contemporary skin specimens make possible to detect whether the skin preservation used were natural or assisted by chemicals (used in mummification processes). This can either represent a natural process of drying, which is often seen for mummies found in hot or cold deserts, or an artificial process where the body has been treated with different substances to promote preservation. In all the spectra of mummified skin observed changes due to degradation in protein structure. A progressive loss of protein amide I (1640–1680 cm−1 ) and amide III (1220–1290 cm−1 ) band intensities indicates loss of protein and changes in the secondary protein structure. This implies that most of the changes in molecular structure of the skin took place in a relatively short time interval during the natural mummification process. Such information is of great importance in archaeological conservation and can shed light on historical practices also [175]. Similar kind of study has been done for characterization of about 5200 year old skin sample. Contemporary skin was used for comparison in freeze-dried condition (for giving the similar environment) and its molecular structure was compared with that of Iceman skin. The results showed that the proteinaceous moiety of the ancient skin have degraded considerably, although olefinic bonds might oxidised with less or no alteration in the lipoidal component. It has been observed that there were significant alterations in the nature of lipid component of ancient skin. The spectral region 3200 cm−1 to 2700 cm−1 provides C–H stretching modes of the lipid alkyl chains, observed with reduced intensity and reduced width in case of old skin sample indicates that the tissue has lost some of its components. In addition to this the loss of olefinic C–H stretching (around 3060 cm−1 ) indicates that the unsaturated lipid component of older skin has degraded [176]. Pigment identification in artefacts has been successfully studied by obtaining their Raman spectra in order to identify and differentiate ochres (most important earth pigments) found extensively in Byzantine hagiography. Compared to other techniques such as X-ray fluorescence, particle-induced X-ray, gamma-ray emission and scanning electron microscopy, Raman spectroscopy has proved to be more significant particularly with respect to analysis time, sensitivity, specificity, amount of sample, spatial resolution, and immunity to interference [177]. In addition, pigment structure and degradation of manuscript has been identified using Raman techniques [178,179]. A significant effect was observed in the construction of novel micro-Raman-dedicated spectrometers united with holographic notch filters and charge coupled device detectors. This made it a powerful technique for studying variety of materials ranging from epitaxial semiconductor thin films to optoelectronics devices and also from biomaterials of interest to medical science [171]. 4.5. Pharmaceuticals Mid-IR and Raman spectroscopy are versatile tools in pharmaceutics and bio-pharmaceutics, with a wide field of applications ranging from characterization of drug formulations to elucidation of kinetic processes in drug delivery (Fig. 10). It could also provide fast detection and identification of counterfeit medicines. Since many blister package materials provide suitable spectral windows for both exciting and scattering radiations [180]. New developments in applications of Raman spectroscopy for studying drug delivery systems, in particular topical drug delivery have been reviewed [181]. Well-established standard methods coupled with Raman spectroscopy enables to study drug release in semisolid formulations, drug penetration, and influence of penetra-

R.S. Das, Y.K. Agrawal / Vibrational Spectroscopy 57 (2011) 163–176

173

Fig. 11. Example of API peak detection and tablet peak detection which has been correctly identified and symbolised, reflects the genuine tablet, not the counterfeited.

Fig. 10. An FT-Raman spectrum for characterization of antidepressant drugs belongs to the same class (selective serotonin reuptake inhibitors (SSRIs)): (A) Citalopram, (B) Sertraline.

tion modifiers. This approach is also applicable for in vivo studies and in characterizing the structure of colloidal drug carrier systems. The interaction of therapeutic drugs and their target biomolecules is of great interest in pharmaceutical research. Shift in Raman bands and changes in band intensity are used to study the kinetics and mechanism of these drug-target reactions [182–185]. Quality testing for tablet composition and uniformity are excessively critical issues at manufacturing stage in pharmaceutical industries because composition and uniformity are the main concerns. A non-destructive and cost-effective analysis by Raman technique played an important role for increasing the quality of products. Thus, Raman spectroscopy reduces time to analyse active ingredient composition of tablets while increasing the number of tablets tested with increased confidence level [186]. The crystal forms present in the drug products can be identified by NIR FT-Raman spectroscopy. Determination of polymorphic forms in a number of commercial drug products have been obtained containing polymorphic drug compounds like sorbitol, mannitol, famotidine, acemetacin, carbamazepine, meprobamate and phenylbutazone. The identification of crystal forms in drug products can be carried out on basis of comparison of peak positions, intensities and shapes of reference spectra with the spectra obtained from the drug product by their varying peak positions. Thus, supports the feasibility of analytical assays to identify crystal forms in drug product and can be easily exploited routinely for monitoring phase changes [187]. The quantitative measurements for solid state of pharmaceutical compounds have also been studied by Raman spectroscopy. Most of the organic molecules exhibits clear and well-resolved bands offer possibility of both qualitative as well as quantitative analysis. This feature is essential for pharmaceutical applications where final dosage forms are typically complex blends of numerous additives and excipients. Thus, Raman spectroscopy supports as a reliable tool in various industrial applications for monitoring and controlling solid elaboration processes [188]. Quantitative analysis of active pharmaceutical ingredient (API) in pharmaceutical products is the most essential part of pharmaceutical analysis. A simple linear regression method is developed and statistically validated using FT-Raman spectroscopy for the direct and non-destructive quantitative analysis that too without sample preparation of the active pharmaceutical ingredient (API),

medroxyprogesterone acetate (MPA) in an aqueous pharmaceutical suspension. The results thus obtained were compared with simple linear regression model of HPLC. The results suggested the reliability of the FT-Raman over the HPLC method [189]. Similarly quantitative analysis of amiodarone hydrochloride based on solutions with known concentrations has also been done, obtained limit of detection (LOD) = 2.11 mg/ml. However, this limit is much higher than the corresponding limit obtained from HPLC methodology, but the proposed method was also validated on Raman spectroscopy by evaluating the linearity of the calibration line as well as its accuracy and precision [190]. Raman spectroscopy has also shown its utility in Process analytical technologies (PAT) commonly used for manufacturing processes, for monitoring the synthesis of API, identification of raw materials and quantitative determination in finished dosage forms [191]. Likewise, the analysis of pharmaceutical solid dosage forms can also be carried out easily. The study has been made for identification of tablets with the applications in release of final products in quality control and detection of counterfeits. Extending, it also gives application in the development of calibration strategies for automatic identification of tablets from their API peak stored in database spectra collection. This can permit the identification of tablets and other pharmaceutical products in future to discriminate counterfeits from genuine stuff (Fig. 11) [192]. Determination of diphenhydramine hydrochloride (DPH), the active ingredient of Benadryl has been studied through FT-Raman spectroscopy. The reliability of this method was verified by testing it against the conventional HPLC, obtained with satisfactory results even in the presence of numerous excipients. The main difference between these two techniques was with their LOD = 0.81 ␮g/ml for HPLC, while LOD = 0.14 mg/ml for FT-Raman spectroscopy, but involves tedious sample preparation in former process [193]. Studies related to structure and interactions of the drug molecules can be well resolved with Raman spectroscopy by examining the shift in characteristic vibrational frequencies. This involves drug reactivity, such as hydrogen bonding, proton transfer, charge transfer and ion-molecule attraction, allows better understanding of the binding properties of drug in biological medium [194]. Other than drug analysis, there are also various substances which have profound use in pharmaceutical industries like surfactants, detergents, cosmetic products, etc. Their analysis is of particular interest with Raman spectroscopy as they contain water, which is a weak Raman scatterer and does not contribute significantly to solvent effects or band broadening phenomena. Thus, solutes can be measured in aqueous solutions facilitating more

174

R.S. Das, Y.K. Agrawal / Vibrational Spectroscopy 57 (2011) 163–176

Fig. 12. (A) Typical Raman spectrum of C60 molecule measured with 514.5 nm laser excitation; (B) typical Raman spectrum of single-walled carbon nanotubes.

effective spectral subtractions or manipulations and quantitative analysis without sample preparation [195]. 4.6. Nanotechnology Raman spectroscopy undoubtedly emerged as a relevant tool for probing and characterization of nanomaterials like nanosensor, nanotube and nanowire, as these are intensively studied in various areas of science due to their unique mechanical, electrical and chemical properties. For better understanding of nanomaterials like nanoceramic, nanocomposite, glassy material and semiconductor, techniques like micro Raman spectroscopy have been successfully applied for their better characterization [141,196,197]. Based on theoretical considerations and comparisons with absorption and luminescence data RRS cross-sections were derived and used to estimate the relative abundance of different species in the sample. It was observed that the total level of semiconductive single wall carbon nano tubes (SWCNTs) was about 11 times higher than that of the metallic species [147]. The same approaches was used to test the selectivity of DNA wrapping as a purification method which has given a mixture enriched with semiconductive SWCNTs [198]. Raman spectroscopy is one of the leading investigation techniques for the characterization of covalently functionalized metallic and semiconducting carbon nano tubes (CNTs). However, CNTs need to be functionalized for handling, sorting (according to electronic or structural characteristics) and property adjustments [199,200]. Semi-empirical quantum simulations of water or methanol absorption on CNTs and fullerenes have been studied profoundly with experimental Raman spectra, exploring the applications of CNTs and fullerenes in selectively absorbing nanofilters and substrates. The Raman spectrum of both the species can provide valuable information regarding the structure and properties of carbon species (Fig. 12) [201]. In addition, one of the latest applications of Nanotechnology is the development of improved chemical and biological sensors. Recent developments in the design and application of two types of optical nanosensors have been reviewed. It was based on localized surface plasmon resonance (LSPR) spectroscopy which measures change in local refractive index and surface-enhanced Raman scattering (SERS) by measuring the vibrational bands of the target molecule [202]. 5. Conclusion Vibrational spectroscopy is an excellent method for identifying substances by providing unique fingerprint spectra. Raman spectroscopy is one of the fastest and non-destructive analytical technique that give the vibrational spectrum and physical or chemical information of virtually any matrix in any state of matter. Thus, in this article we have discussed with the various possibilities and potentials of Raman measurements, with special emphasis on

recent technical developments. In recent years, this technique has been explored with much technical advancement like wide range of laser wavelengths, sampling conditions, instrumentation and data-processing methods, applicable in various fields of science. Additionally, Raman signals can be collected with a small probe head linked with the (portable) apparatus by optical fibre gives the flexibility for online process monitoring. The growing interest in Raman spectroscopy is probably due to its major advantages as it brings direct results of the analysis without sample preparation and easy interpretation, making it as a time saving and cost-effective technique.

References [1] C.V. Raman, R.S. Krishnan, Nature 121 (1928) 501. [2] H.H. Willard, L.L. Merritt, J.A. Dean, F.A. Settle, Instrumental Methods of Analysis, seventh ed., CBS Publishers, New Delhi. [3] H.S. Kaur, Instrumental Methods of Chemical Analysis, third ed., Pragati Prakashan, Meerut, 2006. [4] D.E. Bugay, P.A. Martogolio Smith, in: S. Jickells, A. Negrusz (Eds.), Clarke’s Analytical Forensic Toxicology, third ed., Pharmaceutical Press, London, 2008, pp. 455–468 (Chapter 17). [5] S.F. Parker, Spectrochim. Acta 50A (1994) 1841–1856. [6] Y. Wang, R.L. McCreery, Anal. Chem. 61 (1989) 2647–2651. [7] R.L. McCreery, Raman Spectroscopy for Chemical Analysis, third ed., John Wiley, New York, 2000. [8] P. Hendra, C. Jones, G. Warner, Fourier Transform Raman Spectroscopy, Instrumentation and Applications, Ellis Harwood, New York, 1991. [9] J.R. Ferraro, K. Nakamoto, Introductory Raman Spectroscopy, second ed., Academic Press, Boston, 1994. [10] R.J. Dijkstra, F. Ariese, C. Gooijer, U.A. Brinkman, Trends Anal. Chem. 24 (2005) 304–323. [11] T.D. Nguyen Hong, M. Jouan, N. Quy Dao, M. Bouraly, F. Mantisi, J. Chromatogr. A 743 (1996) 323–327. [12] S.D. Cooper, M.M. Robson, D.N. Batchelder, K.D. Bartle, Chromatography 44 (1997) 257–262. [13] Z.Q. Tian, W.H. Li, M.B. Ren, B.W. Mao, J.G.J.G. Chen, J.Q. Mu, X.D. Zhuo, D. Wang, J. Electroanal. Chem. 401 (1996) 247–251. [14] M. Lopes, A. Candini, M. Urdampilleta, A.R. Plantey, V. Bellini, S. Klyatskaya, L. Marty, M. Ruben, M. Affronte, W. Wernsdorfer, N. Bendiab, J. ACS Nano 4 (2010) 7531–7537. [15] D.L. Jeanmaire, R.P. van Duyne, J. Electroanal. Chem. 84 (1977) 1–20. [16] M.G. Albrecht, J.A. Creighton, J. Am. Chem. Soc. 99 (1977) 5215–5219. [17] P. Kambhampati, C.M. Child, M.C. Foster, A. Campion, J. Chem. Phys. 108 (1998) 5013–5026. [18] U. Kreibig, M. Vollmer, Optical Properties of Metal Clusters, Springer series in Materials Science, No. 25, Berlin, 1995, pp. 26–76. [19] M. Moskovits, J. Raman Spectrosc. 36 (2005) 485–496. [20] C.L. Haynes, A.D. McFarland, R.P. Van Duyne, Anal. Chem. 77 (2005) 338A–346A. [21] D. Zeisel, V. Deckert, R. Zenobi, T. Vo-Dinh, Chem. Phys. Lett. 283 (1998) 381. [22] M. Micic, N. Klymyshyn, Y.D. Suh, H.P. Lu, J. Phys. Chem. B 107 (2003) 1574. [23] S.H. Christiansen, M. Becker, S. Fahlbusch, J. Michler, V. Sivakov, G. Andra, R. Geiger, J. Nanotechnol. 18 (2007) 1–6. [24] L. Xiumei, W. Xiang, L. Zheng, R. Bin, Acta Phys. Chim. Sin. 11 (2008) 1941–1944. [25] Q.J. Huang, X.Q. Li, J.L. Yao, B. Ren, W.B. Cai, J.S. Gao, B.W. Mao, Z.Q. Tian, Surf. Sci. 427/428 (1999) 162–166. [26] A. Torreggiania, Z. Jurasekovab, M. D’Angelantonioa, M. Tambaa, J.V. GarciaRamosb, S. Sanchez-Cortes, Colloid. Surf. A: Physicochem. Eng. Aspects 339 (2009) 60–67. [27] P.A. Mosier-Boss, S.H. Lieberman, Anal. Chim. Acta 488 (2003) 15–23.

R.S. Das, Y.K. Agrawal / Vibrational Spectroscopy 57 (2011) 163–176 [28] J. Feng Li, Y. Fan Huang, Y. Ding, Z. Lin Yang, S. Bo Li, X.S. Zhou, F.R. Fan, W. Zhang, Z.Y. Zhou, D.Y. Wu, B. Ren, Z.L. Wang, Z.Q. Tian, Lett. Nat. 464 (2010) 392–396. [29] E.H.K. Stelzer, R.W. Wijnaendts-van-Resandt, in: T. Wilson (Ed.), Confocal Microscopy, Academic Press, London, 1990, pp. 79–87. [30] T. Vankeirsbilck, A. Vercauteren, W. Baeyens, G. Van der Weken, Trends Anal. Chem. 21 (2002) 869–877. [31] K.R. Allakhverdiev, D. Lovera, V. Altstadt, P. Schreier, L. Kador, Rev. Adv. Mater. Sci. 20 (2009) 77–84. [32] C.J.H. Brenan, I.W.J. Hunter, J. Raman Spectrosc. 27 (1996) 561–571. [33] A.P. Esposito, C.E. Talley, T. Huser, C.W. Hollars, C.M. Schaldach, S.M. Lane, Appl. Spectrosc. 57 (2003) 868–871. [34] J.W. Chan, A.P. Esposito, C.E. Talley, C.W. Hollars, S.M. Lane, T. Huser, J. Anal. Chem. 76 (2004) 599–603. [35] P.J. Caspers, G.W. Lucassen, G.J. Puppels, J. Biophys. 85 (2003) 572–580. [36] W.R. Zhang, C. Lowe, R. Smith, Prog. Org. Coat. 66 (2009) 141–148. [37] S.J.F. Erich, J. Laven, L. Pel, H.P. Huinink, K. Kopinga, Prog. Org. Coat. 52 (2005) 210–216. [38] K.N. Jallad, H.G. Hedderich, J. Hazard. Mater. 124 (2005) 236–240. [39] P.D. Maker, R.W. Terhune, Phys. Rev. 137 (1965) A801–A818. [40] R.F. Begley, A.B. Harvey, R.L. Byer, Appl. Phys. Lett. 25 (1974) 387. [41] S.A. Schaertel, A.C. Albrecht, A. Lau, A. Kummrow, Appl. Phys. B: Lasers Opt. 59 (1994) 377–387. [42] J.X. Cheng, X.S. Xie, J. Phys. Chem. B 108 (2004) 827–840. [43] S. Roy, J.R. Gord, A.K. Patnaik, Prog. Energy Combust. Sci. 36 (2010) 280–306. [44] J.X. Cheng, Y.K. Jia, G.F. Zheng, X.S. Xie, Biophys. J. 83 (2002) 502. [45] X.L. Nan, J.X. Cheng, X.S. Xie, J. Lipid Res. 44 (2003) 2202. [46] H.F. Wang, Y. Fu, P. Zickmund, R.Y. Shi, J.X. Cheng, Biophys. J. 89 (2005) 581. [47] J.X. Cheng, A. Volkmer, L.D. Book, X.S. Xie, J. Phys. Chem. B 106 (2002) 8493–8498. [48] V.V. Krishnamachari, E.O. Potma, J. Opt. Soc. Am. A 24 (2007) 1138–1147. [49] J.X. Cheng, A. Volkmer, L.D. Book, X.S. Xie, J. Phys. Chem. B 105 (2001) 1277–1280. [50] C.L. Evans, E.O. Potma, M. Puoris haag, D. Cote, C.P. Lin, X.S. Xie, Proc. Natl. Acad. Sci. U. S. A. 102 (2005) 16807. [51] Fouad El-Diasty, Vib. Spectrosc. 55 (2011) 1–37. [52] S.F. Hanna, W.D. Kulatilaka, Z. Arp, T. Opatrny, M.O. Scully, J.P. Kuehner, Appl. Phys. Lett. 83 (2003) 1887–1889. [53] W.D. Kulatilaka, N. Chai, S.V. Naik, N.M. Laurendeau, R.P. Lucht, J.P. Kuehner, Opt. Lett. 31 (2006) 3357–3359. [54] N. Chai, S.V. Naik, W.D. Kulatilaka, N.M. Laurendeau, R.P. Lucht, S. Roy, Appl. Phys. B: Lasers Opt. 87 (2007) 731–737. [55] H.A. Rinia, K.N.J. Burger, M. Muller, Biophys. J. 95 (2008) 4908–4914. [56] M. Bonn, M. Muller, K.N.J. Burger, J. Raman Spectrosc. 40 (2009) 763–769. [57] G.I. Petrov, R. Arora, M.O. Scully, Proc. Natl. Acad. Sci. U. S. A. 104 (2007) 7776–7779. [58] K. Tada, N. Karasawa, Opt. Commun. 282 (2009) 3948–3952. [59] S.H. Parekh, Y.J. Lee, K.A. Aamer, M.T. Cicerone, Biophys. J. 99 (2010) 2695–2704. [60] A. Volkmer, L.D. Book, X.S. Xie, Appl. Phys. Lett. 80 (2002) 1505–1507. [61] J.L. Oudar, R.W. Smith, Y.R. Shen, Appl. Phys. Lett. 34 (1979) 758–760. [62] T.W. Kee, H. Zhao, M.T. Cicerone, Opt. Express 14 (2006) 3631–3640. [63] P.R. Carey, Biochemical Applications of Raman and Resonance Raman Spectroscopies, Academic Press, New York, 1982, pp. 169–183. [64] T.A. Matitoli, A. Hoffmann, D.G. Sockalingum, Spectrochim. Acta A 49 (1993) 785–799. [65] E.V. Efremov, F. Ariese, C. Gooijer, J. Anal. Chim. Acta 606 (2008) 119–134. [66] W.E. Smith, P.C. White, C. Rodger, G. Dent, in: I.R. Lewis, H.G.M. Edwards (Eds.), Handbook of Raman Spectroscopy: From the Research Laboratory to the Process Line, Marcel Dekker, New York, USA, 2001, p. 733. [67] N.Q. Dao, M. Jouan, J. Sens. Actuators B 11 (1993) 147–160. [68] T.F. Cooney, H.T. Skinner, S.M. Angel, Appl. Spectrosc. 50 (1995) 836–996. [69] T.F. Cooney, H.T. Skinner, S.M. Angel, Appl. Spectrosc. 50 (1996) 849–860. [70] L.S. Greek, H.G. Schulze, C.A. Haynes, M.W. Blades, R.F.B. Turner, Appl. Opt. 35 (1996) 4086–4095. [71] K.J. Ewing, G. Nau, T. Bilodeau, D.M. Dagenais, F. Bucholtz, I.D. Aggarwal, Anal. Chim. Acta 340 (1997) 227–232. [72] H.T. Skinner, T.F. Cooney, S.K. Sharma, S.M. Angel, Appl. Spectrosc. 50 (1996) 1007–1014. [73] S.D. Harvey, M.E. Vucelick, R.N. Lee, B.W. Wright, Forensic Sci. Int. 125 (2002) 12–21. [74] C.M. Hodges, J. Akhavan, Spectrochim. Acta A: Mol. Spectrosc. 46 (1990) 303–307. [75] E.M. Suzuki, M. Carrabba, J. Forensic Sci. 1 (2001) 1053–1069. [76] G. Ellis, Spectrochim. Acta A 46 (1990) 221–241. [77] C. Rodger, D. Broughton, Analyst 123 (1998) 1823–1826. [78] K. Virkler, I.K. Lednev, Forensic Sci. Int. 181 (2008) e1–e5. [79] K. Virkler, I.K. Lednev, Forensic Sci. Int. 193 (2009) 56–62. [80] J.Z. Palus, M. Kunicki, Forensic Sci. Int. 158 (2006) 164–172. [81] P.L. Lang, J.E. Katon, J.F. O’Keefe, D.W. Schiering, J. Microchem. 34 (1986) 319–331. [82] I.P. Keen, G.W. White, P.M. Fredericks, J. Forensic Sci. 43 (1) (1998) 82–89. [83] J. Thomas, P. Buzzini, G. Massonnet, B. Reedy, C. Roux, Forensic Sci. Int. 152 (2005) 189–197. [84] L. Lepot, K. De Wael, F. Gason, B. Gilbert, Sci. Justice 48 (2008) 109–117.

175

[85] L.C. Abbott, S.N. Batchelor, J.R. Lindsay Smith, J.N. Moore, Forensic Sci. Int. 202 (2010) 54–63. [86] J.S. Daya, H.G.M. Edwards, S.A. Dobrowski, A.M. Voice, Spectrochim. Acta A 60 (2004) 1725–7130. [87] S. Armenta, S. Garrigues, M. de la Guardia, Anal. Chim. Acta 547 (2005) 197–203. [88] N.A. Hatab, G. Eres, B. Paul, Hatzinger, Gu Baohua, J. Raman Spectrosc. 41 (2010) 1131–1136. [89] F.T. Docherty, P.B. Monaghan, C.J. McHugh, D. Graham, W.E. Smith, J.M. Cooper, IEEE Sens. 5 (2005) 632–638. [90] E. Etz, S. Roberson, G. Gillen, Microsc. Microanal. 8 (2002) 1520–1521. [91] F.C.C. Oliveira, C.R.R. Brandao, H.F. Ramalho, L.A.F. da Costa, P.A.Z. Suarez, J.C. Rubim, Anal. Chim. Acta 587 (2007) 194–199. [92] C. Kraft, T. Knetschke, A. Siegner, R.H.W. Funk, R. Salzer, Vib. Spectrosc. 32 (2003) 75–83. [93] I. Notingher, J. Sens. 7 (2007) 1343–1358. [94] W.L. Peticolas, J. Biochimie 57 (1975) 417–428. [95] K.R. Ward, R.W. Barbee, P.S. Reynolds, I.P. Torres-Filho, M.H. Tiba, L. Torres, R.N. Pittman, J. Terner, Anal. Chem. 79 (2007) 1514. [96] H.A. Rinia, M. Bonn, E.M. Vartiainen, C.B. Schaffer, M. Muller, J. Biomed. Opt. 11 (2006), 050502-1–050502-3. [97] D. Pestov, M. Zhi, Z.E. Sariyanni, N.G. Kalugin, A.A. Kolomenskii, R. Murawski, G.G. Paulus, V.A. Sautenkov, H. Schuessler, A.V. Sokolov, G.R. Welch, Y.V. Rostovtsev, T. Siebert, D.A. Akimov, S. Graefe, W. Kiefer, M.O. Scully, Proc. Natl. Acad. Sci. U. S. A. 102 (2005) 14937–14938. [98] S.O. Konorov, C.H. Glover, J.M. Piret, J. Bryan, H.G. Schulze, M.W. Blades, R.F.B. Turner, Anal. Chem. 79 (2007) 7221. [99] S.A. Overman, G.J. Thomas, J. Raman Spectrosc. 29 (1998) 23–29. [100] P.R. Carey, J. Raman Spectrosc. 29 (1998) 7–14. [101] R. Callender, H. Deng, R. Gilmanshin, J. Raman Spectrosc. 29 (1998) 15–21. [102] X. Zhao, T.G. Spiro, J. Raman Spectrosc. 29 (1) (1998) 49–55. [103] L. da Chiu, M. Ando, H. Hamaguchi, J. Raman Spectrosc. 4 (2010) 2–3. [104] A.M. Herrero, Food Chem. 107 (2008) 1642–1651. [105] R.A. Dluhy, S.M. Stephens, S. Widayati, A.D. Williams, Spectrochim. Acta A 51 (1995) 1413–1447. [106] D. Pappas, B.W. Smith, J.D. Winefordner, Talanta 51 (2000) 131–144. [107] P.R.T. Jess, D.D.W. Smith, M. Mazilu, K. Dholakia, A.C. Riches, C.S. Herrington, Int. J. Cancer 121 (2007) 2723. [108] R. Tuma, G.J. Thomas, Biophys. Chem. 68 (1997) 17–31. [109] J. Guicheteau, S. Christesen, D. Emgea, A. Tripathi, J. Raman Spectrosc. 41 (2010) 1632–1637. [110] N. Howell, E. Li-Chan, Int. J. Food Sci. Technol. 31 (1996) 439–451. [111] J.D. Guo, T.W. Zerda, J. Phys. Chem. B 101 (1997) 5490–5496. [112] N.M. Sijtsema, J.J. Duindam, G.J. Puppels, C. Otto, J. Greve, Appl. Spectrosc. 50 (1996) 545–551. [113] S. Wessel, M. Gniadecka, G.B.E. Jemec, H.C. Wulf, J. Biochim. Biophys. Acta 1433 (1999) 210–216. [114] J.F. Neault, M. NaouP, M. Manfait, H.A. Tajmir-Riah, FEBS Lett. 382 (1996) 26–30. [115] C. Xie, D. Chen, Y.Q. Li, Opt. Lett. 30 (2005) 1800–1802. [116] C.M. Gutierrez, J.L. Quintanar, C. Frausto-Reyes, R.S. Berru, Spectrochim. Acta A 61 (2005) 87–91. [117] A.C. Williams, H.G.M. Edwards, B.W. Barry, Int. J. Pharma 81 (1992) 11–14. [118] R. Manoharan, Y. Wang, M.S. Feld, Spectrochim. Acta A 52 (1996) 215–249. [119] X. Gaoa, I.S. Butler, R. Kremer, Spectrochim. Acta A 61 (2005) 27–35. [120] C.W. Ong, Z.X. Shen, Y. He, T. Lee, S.H. Tang, J. Raman Spectrosc. 30 (1999) 91–96. [121] J. Sajid, A. Elhaddoui, S. Turrel, J. Raman Spectrosc. 28 (1997) 165–169. [122] A.R. de Paula Jr., S. Sathaiah, Med. Eng. Phys. 27 (2005) 237–244. [123] H.P. Buschman, J.T. Matz, G. Deinum, T.J. Romer, M. Fitzmaurice, J.R. Kramer, A. van deer Laarse, A.V. Bruschke, M.S. Feld, J. Cardiovas. Pathol. 10 (2001) 59–68. [124] J.P. Salenius, J.F. Brennan, A. Miller, Y. Wang, T. Aretz, B. Sacks, R.R. Dasari, M.S. Feld, J. Vasc. Surg. 27 (1998) 710–719. [125] P. Weinmann, M. Jouan, N.Q. Dao, B. Lacroix, C. Groiselle, J.P. Bonte, G. Luc, Atherosclerosis 140 (1998) 81–88. [126] C.J. Frank, D.C.B. Redd, T.S. Gansler, R.L. McCreery, Anal. Chem. 66 (1994) 319–326. [127] N.J. Kline, P.J. Treado, J. Raman Spectrosc. 28 (1997) 119–124. [128] D.C.B. Redd, Z.C. Feng, K.T. Yue, T.S. Gansler, Appl. Spectrosc. 47 (1993) 787–791. [129] C.J. Frank, R.L. McCreery, D.C.B. Redd, T.S. Gansler, Appl. Spectrosc. 47 (1993) 387–390. [130] M.D. Schaeberle, V.F. Kalasinsky, J.L. Luke, E.N. Lewis, J.W. Levin, P.J. Treado, Anal. Chem. 68 (1996) 1829–1833. [131] V. Vrabie, C. Gobinet, O. Piot, A. Tfayli, P. Bernard, R. Huez, M. Manfait, Biomed. Signal Process. Control 2 (2007) 40–50. [132] K.K. Kumar, A. Anand, M.V.P. Chowdary, Keerthi, J. Kurien, C.M. Krishna, S. Mathew, Vib. Spectrosc. 44 (2007) 382–387. [133] C.M. Krishna, N.B. Prathima, R. Malini, B.M. Vadhiraja, R.A. Bhatt, D.J. Fernandes, P. Kushtagi, M.S. Vidyasagar, V.B. Kartha, Vib. Spectrosc. 41 (2006) 136–141. [134] K. Bensalah, J. Fleureau, D. Rolland, O. Lavastre, N.R. Leclercq, F. Guille, J.J. Patard, L. Senhadji, R. de Crevoisier, J. Eur. Urol. 58 (2010) 602–608. [135] V.R. Kodati, A.T. Yu, Appl. Spectrosc. 44 (1990) 837–839.

176

R.S. Das, Y.K. Agrawal / Vibrational Spectroscopy 57 (2011) 163–176

[136] V.R. Kodati, G.E. Tomasi, J.L. Turumin, A.T. Yu, Appl. Spectrosc. 44 (1990) 1405–1411. [137] B. Schrader, B. Dippel, S. Fendel, S. Keller, T. Liichte, M. Riedl, R. Schulte, E. Tatsch, J. Mol. Struct. 408/409 (1997) 23–31. [138] M.H. Schweitzer, M. Marshall, K. Carron, D.S. Bohle, S.C. Busse, E.V. Arnold, D. Barnard, J.R. Horner, J.R. Starkey, Proc. Natl. Acad. Sci. U. S. A. 94 (1997) 6291–6296. [139] A. Bertoluzza, P. Brasili, L. Castri, F. Facchini, C. Fagnano, A. Tinti, J. Raman Spectrosc. 28 (1997) 185–188. [140] K. Ryu, A.J. Haes, H.Y. Park, S. Nah, J. Kim, H. Chung, M.Y. Yoona, S.H. Han, J. Raman Spectrosc. 41 (2010) 121–124. [141] G. Gouadec, P. Colomban, Prog. Cryst. Growth Charact. Mater. 53 (2007) 1–56. [142] G. Abstreiter, Appl. Surf. Sci. 50 (1991) 73–78. [143] D. Nesheva, J. Optoelectro. Adv. Mater. 7 (2005) 185–192. [144] J. Zhu, F. Xu, S.J. Schofer, C.A. Mirkin, J. Am. Chem. Soc. 119 (1997) 235–236. [145] Y. Dai, J.S. Swinnea, H. Steinfink, J.B. Goodenough, A. Campion, J. Am. Chem. Soc. 109 (1987) 5291–5292. [146] L. Cao, L. Laim, P.D. Valenzuela, R.R. Martin, S.S. Nonmann, E.M. Gallo, B. Nabet, J.E. Spanier, J. Raman Spectrosc. 38 (2007) 697. [147] A. Jorio, A.P. Santos, H.B. Ribeiro, C. Fantini, M. Souza, J.P.M. Vieira, C.A. Furtado, J. Jiang, R. Saito, L. Balzano, D.E. Resasco, M.A. Pimenta, Phys. Rev. B 72 (2005) 075207/1–075207/5. [148] C. Thomsen, S. Reich, J. Topics Appl. Phys. 108 (2007) 115–234. [149] M.C. Polo, J. Cifre, G. Sanchez, R. Aguiar, M. Varela, J. Polo, Appl. Phys. Lett. 67 (1995) 485–487. [150] A. Kasuya, Y. Saito, Y. Sasaki, M. Fukushima, T. Maeda, C. Horie, Y. Nishina, Mater. Sci. Eng. A 218 (1996) 46–47. [151] S. Fonti, A. Blanco, E. Bussoletti, L. Colangeli, M. Lugara, V. Mennella, V. Orofino, G. Scamarcio, Infrared Phys. 30 (1990) 19–25. [152] R.A. Friedel, G.L. Carlson, Fuel 51 (1972) 194–198. [153] J.K. Agbenyega, G. Ellis, P.J. Hendra, W.F. Maddams, C. Passingham, H.A. Willis, Spectrochim. Acta A 46 (1990) 197–216. [154] K. Prasad, D.T. Grubb, J. Appl. Polym. Sci. 41 (1990) 2189–2198. [155] H. Ko, S. Chang, V.V. Tsukruk, ACS Nano 3 (2009) 181–188. [156] J. Doshi, D.H. Reneker, J. Electros 35 (1995) 151–160. [157] A. Ianoul, T. Coleman, S.A. Asher, Anal. Chem. 74 (2002) 1458. [158] R.A. Halvorson, P.J. Vivesland, J. Environ. Sci. Technol. 44 (2010) 7749–7755. [159] C. Ruan, W. Luo, W. Wang, B. Gu, Anal. Chim. Acta 605 (2007) 80–86. [160] B. Gu, J. Tio, W. Wang, Y.K. Ku, S. Dai, Appl. Spectrosc. 58 (2004) 741–744. [161] J.J. Barrett, M.J. Berry, Appl. Phys. Lett. 34 (1979) 144–146. [162] Y. Oki, S. Nakazono, Y. Nonaka, M. Maeda, Opt. Lett. 25 (2000) 1040–1042. [163] K.S. Kalasinsky, T. Hadfield, A.A. Shea, V.F. Kalasinsky, M.P. Nelson, J. Neiss, A.J. Drauch, G.S. Vanni, P.J. Treado, Anal. Chem. 79 (2007) 2658–2673. [164] M.L. Fish, O. Massler, J.A. Reid, R. MacGregor, J.D. Comins, Diamond Relat. Mater. 8 (1999) 1511–1514. [165] G. Tas, S.A. Hambir, J. Franken, D.E. Har, D.D. Dlott, J. Appl. Phys. 82 (1997) 1080–1087. [166] S.S. Potgieter-Vermaak, J.H. Potgieter, R. Van Grieken, Cem. Concr. Res. 36 (2006) 656–662. [167] H.L. Qiu, A.H. Wang, J.L. You, X.J. Liu, H. Chen, S.T. Yin, Spectrosc. Spectral Anal. 25 (2005) 222–225.

[168] Y. Djaoued, M. Thibodeau, J. Robichaud, S. Balaji, S. Priya, N. Tchoukanova, S.S. Bates, Spectrochim. Acta A Mol. Biomol. Spectrosc. 67 (2007) 1362–1369. [169] R.J.H. Clark, J. Mol. Struct. 59 (1980) 137–145. [170] R.W. Lurn, Am. Miner. 73 (1988) 297–305. [171] S. Jimenez-Sandoval, J. Microelectron. 31 (2000) 419–427. [172] A. Wang, B.L. Jolliff, L.A. Haskin, J. Geophys. Res. 100 (1995) 21189–21199. [173] T.N. Day, P.J. Hendra, A.J. Rest, A.J. Rowlands, Spectrochim. Acta A 47 (9/10) (1991) 1251–1262. [174] Y. Ikushima, J. Corros. Eng. 49 (2000) 117–121. [175] H.G.M. Edwards, M. Gniadecka, S. Petersen, J.P.H. Hansen, O.F. Nielsen, D.H. Christensen, H.C. Wulf, Vib. Spectrosc. 28 (2002) 3–15. [176] A.C. Williams, H.G.M. Edwards, B.W. Barry, Biochim. Biophys. Acta 1246 (1995) 98–105. [177] D. Bikiaris, S. Sister Daniilia, O. Sotiropoulou, E. Katsimbiri, A.P. Pavlidou, Y. Moutsatsou, Chryssoulakis, Spectrochim. Acta A 56 (1999) 3–18. [178] R.J.H. Clark, P.J. Gibbs, K.R. Seddon, N.M. Brovenko, Y.A. Petrosyan, J. Raman Spectrosc. 28 (1997) 91–94. [179] L. Bugio, R.J.H. Clark, Spectroc. Acta Part. A Mol. Biomol. Spectrosc. 57 (2001) 1491–1521. [180] C. Eliasson, P. Matousek, Anal. Chem. 79 (2007) 1696. [181] S. Wartewiga, R.H.H. Neubert, Adv. Drug Deliv. Rev. 57 (2005) 1144–1170. [182] A. Torreggiani, G. Fini, J. Raman Spectrosc. 30 (1999) 295–300. [183] M.M. Glice, A. Les, K. Bajdor, J. Mol. Struct. 450 (1998) 141–143. [184] C.A. Butler, R.P. Cooney, W.A. Denny, Appl. Spectrosc. 48 (1994) 822–826. [185] A. Weselucha-Birczynska, K. Nakamoto, J. Raman Spectrosc. 27 (1996) 915–919. [186] W. Bonawi-Tan, D.A. Stuart Williams, J. Manuf. Syst. 23 (2004) 299–308. [187] M.E. Auera, U.J. Griessera, J. Sawatzki, J. Mol. Struct. 661/662 (2003) 307–317. [188] G. Fevotte, Trans. IChemE A Chem. Eng. Res. Des. 85 (2007) 906–920. [189] T.R.M. De Beer, G.J. Vergote, W.R.G. Baeyens, J.P. Remon, C. Vervaet, F. Verpoort, Eur. J. Pharm. Sci. 23 (2004) 355–362. [190] M.G. Orkoula, C.G. Kontoyannis, C.K. Markopoulou, J.E. Koundourellis, Talanta 73 (2007) 258–261. [191] J. Muller, K. Knop, M. Wirges, P. Kleinebudde, J. Pharma. Biomed. Anal. 53 (2010) 884–894. [192] Y. Roggo, K. Degardin, P. Margot, Talanta 81 (2010) 988–995. [193] M.G. Orkoula, C.G. Kontoyannis, C.K. Markopoulou, J.E. Koundourellis, J. Pharma. Biomed. Anal. 41 (2006) 1406–1411. [194] P.V. Huong, J. Pharma. Biomed. Anal. 4 (1986) 811–823. [195] J.G. Grasselli, F. Walder, C. Petty, G. Kemeny, J. Mol. Struct. 294 (1993) 207–210. [196] D. Roy, M. Chhowalla, H. Wang, N. Sano, I. Alexandrou, T.W. Clyne, G.A.J. Amaratunga, Chem. Phys. Lett. 373 (2003) 52. [197] P. Puech, A. Bassil, J. Gonzalez, Ch. Power, E. Flahaut, S. Barrau, Ph. Dernont, C. Lacabanne, E. Perez, W.S. Basca, Phys. Rev. B 72 (2005) 155436–155442. [198] C. Fantini, A. Jorio, A.P. Santos, V.S.T. Peressinotto, M.A. Pimenta, Chem. Phys. Lett. 439 (2007) 138. [199] M.S. Dresselhaus, G. Dresselhaus, R. Saito, A. Jorio, Phys. Rep. 409 (2005) 47. [200] R. Graupner, J. Raman Spectrosc. 38 (2007) 673–683. [201] M.S. Amer, J. Raman Spectrosc. 38 (2007) 721–727. [202] C.R. Yonzon, D.A. Stuart, X. Zhang, A.D. McFarland, C.L. Haynes, R.P. Van Duyne, Talanta 67 (2005) 438–448.