Send Orders of Reprints at bspsaif@emirates.net.ae

108 Current Analytical Chemistry, 2013, 9, 108-127

1573-4110/13 $58.00+.00 © 2013 Bentham Science Publishers

Recent Advances in Raman Analysis of Plants: Alkaloids, Carotenoids, and Polyacetylenes

Malgorzata Baranskaa,*, Maciej Romana, Jan Cz. Dobrowolskib,c, Hartwig Schulz d and Rafal Baranskie,*

aFaculty of Chemistry, Jagiellonian University, Ingardena 3, 30-060 Cracow, Poland

bNational Medicines Institute, 30/34 Che mska Street, 00-725 Warsaw, Poland

cIndustrial Chemistry Research Institute, 8 Rydygiera Street, 01-793 Warsaw, Poland

dJulius Kühn-Institut, Institute for Ecological Chemistry, Plant Analysis and Stored Product Protection, Erwin-Baur-

Strasse 27, 06484 Quedlinburg, Germany

eDepartment of Genetics, Plant Breeding and Seed Science, University of Agriculture in Krakow, Al. 29 Listopada 54,

31-425 Krakow, Poland

Abstract: This paper demonstrates the special potential of Raman spectroscopy for the study of selected plant metabolites. Carotenoids, which are beneficial components in fruits and vegetables, have been shown to be a significant factor in lowering the risk of various types of cancer and ischemic heart diseases. On the other hand, alkaloids may have various effects on human health, e.g. caffeine is a mild stimulant of the central nervous system and as a result it can influence human behaviour. Polyacetylenes are highly cytotoxic against numerous cancer cell lines and demonstrate antifungal, anti-inflammatory and anti-platelet-aggregatory properties.

In most cases, vibrational measurements can be performed directly on plant tissues as well as on fractions isolated from the plant material by hydro-distillation or solvent extraction. Raman spectroscopy techniques allow obtaining spectra which present some characteristic key bands of individual components. Based on such markers related to individual plant substances, spectroscopic analyses in principle allow the discrimination of different species, and even chemotypes among the same species. Moreover, Raman microspectroscopy provides 2- and 3-dimensional images of the investigated plant samples. These maps can be directly compared to the corresponding visual images obtained from a light microscope and offer additional detailed information regarding the local distribution of specific compounds in the surface layers of the analyzed plant tissue.

Keywords: FT-Raman, Raman mapping, Imaging, In situ, Nondestructive analysis, Secondary Metabolites, Algae, Lichens, Fungi.

1. INTRODUCTION

Raman spectroscopy is a method where a sample is radi-ated with monochromatic visible, ultra-visible or near-infrared light emitted from a laser, which probes molecular vibrations (stretching, bending, deformation) and conse-quently information about the structure of chemical compo-nents present in the investigated sample is gathered. Func-tional groups of the molecules can be identified by their unique pattern of light scattering, called Raman spectrum, with the intensity of the signals proportional to their relative concentration in the sample. Plant tissues contain enzymes and coenzymes that may absorb in the visible spectral range however, when the excitation is applied in the near-infrared range (NIR), by using a Nd:YAG (neodymium doped *Address correspondence to these authors at the Dept. of Genetics, Plant Breeding and Seed Sci., University of Agriculture in Krakow, Al. 29 Listo-pada 54, 31-425 Krakow, Poland; Tel: +48 12-662-51-91; Fax: +48 12-662-52-66; E-mail: baranski@ogr.ur.krakow.pl

Faculty of Chemistry, Jagiellonian University, Ingardena 3, 30-060 Cracow, Poland; Tel: +48 12-663-22-53; Fax: +48 12-634-05-15; E-mail: baranska@chemia.uj.edu.pl

yttrium aluminium garnet) laser at 1064 nm, this effect can be avoided. Thus, in most cases the plant tissue does not excite fluorescence and its thermal breakdown can be re-duced to a minimum (Scheme 1). Additionally, using a Michelson interferometer and Fourier transform (FT) proc-essors for analysis of scattered light an enhancement of the recorded intensity by about two orders of magnitude com-pared to ‘classical’ dispersive spectrometers is observed [1]. The resolution and sensitivity of FT-Raman measurements is usually not as good as those obtained by dispersive instru-ments with excitation in the UV–Vis range. Resonance Ra-man effect in which the exciting laser wavelength is adjusted to the absorption range of particular chromophores within the sample results in a significant sensitivity enhancement. To avoid strong fluorescence and to enhance the sensitivity and selectivity, the Raman measurement can be combined with the absorption of the sample on nanometal particles. This technique is called SERS (surface-enhanced Raman scattering), and allows enhancement of the Raman signals from molecules absorbed on the metal surface by six orders of magnitude or more due to electromagnetic and chemical factors [2]. Another useful technique applied for in situ plant

Recent Advances in Raman Analysis of Plants Current Analytical Chemistry, 2013, Vol. 9, No. 1 109

analysis is Raman mapping or imaging that allows chemical maps of the investigated samples to be obtained, and these can be directly compared with sample visual images (Scheme 2). The derived two-dimensional maps provide detailed information regarding the distribution of specific compounds occurring in the surface layer of the sample [3].

Scheme 1. Raman spectra of thyme essential oil obtained from measurements using diode laser emitting at 830 nm that shows high sample fluorescence and low signal to background ratio, and using Nd:YAG laser emitting at 1064 nm.

The main advantage of Raman spectroscopy applied to fresh plant material is that water, due to its low polarizabil-ity, has low response in contrast to complementary method of infrared (IR) spectroscopy, where intense water bands dominate in the spectra and may overlay the signals from an analyte. Therefore, both dried and fresh plant samples can be used successfully for Raman measurements (Scheme 3) [4].

Combination of vibrational spectroscopy and hierarchical cluster analysis provides a fast, easy and reliable method for chemotaxonomy characterization. The PLS algorithm en-

ables for determination of main plant components with rela-tively high R2 and low SECV (Standard Error of Cross-Validation) values. The ability to rapidly monitor various plant components makes it possible to efficiently select high-quality single plants from wild populations as well as proge-nies of crossing experiments. Furthermore, the vibrational spectroscopy methods can also be used by the processing industry in order to perform fast quality checks of incoming raw materials as well as continuous controlling of the pro-duction [5].

Here, the recent progress in Raman spectroscopy analysis of alkaloids, carotenoids, and polyacetylenes in plants is reviewed. Alkaloids stand for a large and structurally diverse group of nitrogen-containing metabolites, and several of them are used for medicinal purposes. Only a few are de-rived from purines (e.g., caffeine), pyrimidines, or terpenes (e.g., aconitine), while the large majority of alkaloids are produced from amino acids. Carotenoids are represented by 750 naturally occurring compounds, which since the late 1920s (when constitution of first carotenoids were estab-

Scheme 2. Images of biological samples and their Raman maps colored according to the intensity of signals in the region of 1510-1530 cm-1 characteristic for carotenoids. Carrot root slice with the highest carotenoid content in phloem tissue (a, b) and chemomile inflorescence with carotenoids detectable in anthers only (c, d).

Scheme 3. Raman spectrum obtained from the measurement of an oil cell in Eucalyptus leaf showing, among many signals coming from plant matrix, the characteristic band at 1159 cm-1 assigned to 1,8-cineole based on the measurement of the pure standard and the spectrum of an orange spot on a Viola petal showing the shift of zeaxanthin band at 1156 cm-1 in comparison to the position at 1159 cm-1 characteristic for pure standard.

110 Current Analytical Chemistry, 2013, Vol. 9, No. 1 Baranska et al.

lished by Kuhn and co-workers and Karrer and co-workers [6]) had been elucidated. The number of minor natural caro-tenoids has been recently increased by more than 20 struc-tures per year [7]. The health promoting properties of fruits and vegetables can be explained by the presence of other bioactive compounds such as polyacetylenes that occur in plants in minor concentrations.

Analysis of many new compounds has been possible due to the spectacular advances in the analytical methods, their instrumentation, and the development of chemometric tech-niques. Especially chromatography isolation and research including with structure elucidation spectroscopy techniques have impacted the recent success in natural product analyses. Yet, non-invasive and non-destructive spectroscopy methods have innovated quite exceptional new field of analytical chemistry and Raman imaging spectroscopy is currently becoming one of the most powerful method especially for in-situ detection of natural product distribution in plant tis-sues.

2. ALKALOIDS

2.1. Classification of Alkaloids

In 2009 a consecutive book from Elsevier entitled The Alkaloids: Chemistry and Biology (Geoffrey A. Cordell, editor) has been published, where one chapter Determination of Alkaloids through Infrared and Raman Spectroscopy is dedicated to the determination of these compounds by Ra-man and IR spectroscopy [8]. That work can be regarded as a broad and complex review showing the contemporary state of knowledge, so here only the newest data on Raman analy-sis of alkaloids will be presented as well as some of the most significant results received from that previous period. Moreover, in this paper the same classification of alkaloids will be used as in the work cited above since it reflects the variability of this chemical group, however only selected representative alkaloids will be presented and discussed in each class.

2.2. Function in Plants

The alkaloids are low molecular weight nitrogen-containing compounds found mainly in plants, but also to a lesser extent in microorganisms and animals. Ecological role of alkaloids as very prominent class of defense compounds is well documented [9]. These secondary metabolites are usually multifunctional, in many cases even a single alkaloid can exhibit more than one biological function. During evolu-tion, the constitution of alkaloids has been modulated so that they usually contain more than one active functional group, allowing them to interact with several molecular targets and usually more than one group of enemies [9].

2.3. Selected Data on Raman Spectroscopy of Alkaloids

Purine Alkaloids: Caffeine

Caffeine occurs naturally and commonly in coffee beans, but also guarana seeds contain a relatively high concentra-tion of this alkaloid. As minor alkaloids, also theophylline and theobromine can be found in these plants. With Raman spectroscopy theobromine can be distinguished from caf-feine and theophylline by the presence of a band at 620 cm-1,

whereas the other two alkaloids show a strong feature at 556 cm-1. Additionally, a medium doublet for caffeine can be seen at 643 and 741 cm-1 whereas for theophylline a charac-teristic single Raman band occurs at 668 cm-1. Furthermore, the discrimination of anhydrous caffeine and its monohy-drate form can be done by comparing the intensities of the carbonyl bands occurring at 1654 and 1698 cm-1 (see Table 1 in ref. [10]).

Raman spectra of cacao seeds and their extracts also showed the presence of theobromine [11]. Marker bands of theobromine were detected at 1682 (C=N stretching), 1594 (C=C stretching), 1334 (C–N stretching), 1296 (C–N stretch-ing), 1225 (C–N stretching), 776 (O=C–C deformation), and 733 cm-1 (O=C–N deformation). Because the Raman spectra of powdered seed kernels do not show any spectral shift of theobromine, in comparison to the spectrum of the pure standard, it is assumed that this alkaloid occurs naturally in its free form (not coordinated with phenolic substances such as tannins). Beside theobromine, a small amount of caffeine was also found, but no theophylline was detected in the dried cacao powder.

It is worth mentioning that Raman spectroscopy has been successfully applied to detect small amounts of caffeine e.g. in commercial energy drinks [12]; furthermore in situ studies have been performed to analyse its distribution in pharma-ceutical products [13]. Reactivity and active sites of caffeine were studied based on the vibrational behavior of caffeine on a silver colloidal surface at different pH values [14,15].

Opioid Isoquinoline Alkaloids: Morphine

The main alkaloids present in poppy plant material and in related pharmaceutical products were successfully analyzed by Raman spectroscopy [16]. Raman spectra in the finger-print range between 700 and 1500 cm-1 show numerous sharp bands, which are mainly assigned to deformation and stretching vibrations of the alkaloid ring system. Because of their similar molecular structures, morphine, codeine, and thebaine demonstrate no significant differences in their Ra-man spectral profile, however, some characteristic bands can be found in the range of 1600–1650 cm-1 and 630–650 cm-1.

The Raman spectrum obtained from the poppy milk pre-sents very clearly characteristic morphine bands (peaks at 631, 1620 and 1642 cm-1). It has been found that for alka-loids occurring in higher amounts in poppy capsules (mor-phine, codeine, papaverine, and noscapine) the correlation coefficients between the spectral data and reference HPLC values demonstrate an acceptable predictive quality. Also the total alkaloid content of poppy samples could be determined with comparatively high accuracy (R2=0.90) [16]. SERS and conventional Raman spectroscopy methods were employed to investigate the absorption and orientation of morphine on a silver surface. The results suggested that the molecule had a charge transfer adsorption on silver island film, and both planes of its ‘T’ type structure had a rather perpendicular orientation to the substrate, mainly via the lone-pair elec-trons on the oxygen atoms [17].

Morphine is known as a highly potent opiate analgesic drug, which metabolizes in the body to morphine-3-O-glucuronide (antagonist to the analgesic effects of mor-phine). In order to investigate morphine, its metabolite and

Recent Advances in Raman Analysis of Plants Current Analytical Chemistry, 2013, Vol. 9, No. 1 111

salts as well as pharmaceutical product, Fourier transform Raman spectroscopy was used. Experimental spectra of morphine were interpreted with the help of quantum-chemical calculations. The results provided clear evidence of the benefits of Raman spectroscopy in the analysis of mor-phine occurring in various environments [18].

Natural Isoquinoline Alkaloid Dyes: Barberine

The main representative of isoquinoline alkaloids is ber-berine, a yellow coloring substance present in the roots of various Berberis species (Berberidaceae) shrubs [19]. Raman spectra from a pure berberine standard can be recorded with-out any problems, and show well-resolved signals, whereas reliable Raman data from samples colored with berberine are difficult to obtain due to high fluorescence and the low con-centration of the analyte [20]. This is the reason why the SERS technique was introduced to enhance the sensitivity of this alkaloid signals [21]. It was demonstrated that silver nanoparticles added as silver nitrate colloid to the sample, possess great affinity to the positively charged berberine molecule. The most intense signals of the ordinary Raman spectrum occur at 1397 and 1520 cm-1, and have been assigned as the in-plane and ring deformation vibrations, respectively. These lines are missing in the SERS spectrum, which is dominated by three bands at 729, 753, and 770 cm-1, assigned as out-of plane vibrations. Similar results for the SERS characterization of berberine were described by Wang and Yu [22].

Antimalarial Isoquinoline and Naphthyisoquinoline Alka-loids: Quinine

Quinine is a main alkaloid present in the bark of different Cinchona species (Rubiaceae), especially Cinchona pubes-

cens Vahl. [23]. The bark of trees of the Cinchona genus is the source of a variety of alkaloids, the most familiar of which are the following pairs of stereoisomers: cinchonine and cinchonidine, quinine and quinidine, as well as dihydro-quinidine and dihydroquinine. Quinine and quinidine have a bitter taste and due to this property they are used as food additives. The high potential of resonance Raman spectros-copy was demonstrated by the detection of small amounts of these alkaloids in plant materials, without any further sample preparation. The non-resonant Raman microscopy with exci-tation wavelengths in the visible (532 and 633 nm) and NIR (1064 nm) regions was unsuccessful to obtain adequate Ra-man features for identification purposes, due to a strong fluorescence background and due to noisy signals resulting from low laser power, respectively. The use of higher laser power resulted in destruction of the plant material. Non-resonant Raman spectroscopy suffers from low scattering cross section, but sometimes for the active agents, which are present at low concentration in the biological environment, the signals can be strongly enhanced if the excitation laser lies within an electronic absorption of the sample. So when UV resonance Raman microspectroscopy ( exc=244 nm) was applied for analysis of quinine in Cinchona bark the meth-odology allowed Raman spectra of the active agents to be obtained selectively, with enhanced signals by a factor of up to 106 compared to the nonresonant Raman signal of that plant. It was found that the UV resonance Raman spectrum obtained from Cinchona bark corresponds well with that of

pure isolated quinine. Moreover, in situ UV-RR microspec-troscopy is capable of differentiating between quinine and its diastereomer quinidine, structurally very similar active agents. This discrimination is possible based on a marker band at 831cm-1 in the Raman spectrum of quinine, which is shifted to 843 cm-1 in the case of quinidine. This vibration involves a banding motion within the side chain around the chiral centre of quinine. Vibrations belonging to the quino-line ring, important for its antimalarial activity in forming p–p interactions to hemozoin, and the vinyl group are reso-nantly enhanced in the UV-Raman spectra. The interpreta-tion of the experimental spectra was based on DFT calcula-tions combined with FT-Raman spectroscopy of the pure isolated standards. The solvent effect was correlated with the shift of the band at 1362 cm-1 to 1371 cm-1 for anhydrous quinine. This vibration is sensitive to the presence of an aqueous environment and is assigned to a C=C stretching mode [24].

Some other antimalarial alkaloids (dioncophylline A, di-oncophylline C, and dioncopeltine A) belonging to the group of naphthylisoquinoline alkaloids were isolated from the tropical liana Triphyophyllum pelatatum [25]. In order to locate those parts of the plant containing the highest concen-tration of these bioactive substances, Raman microspectro-scopy was found to be a very efficient tool capable of differ-entiating between various structurally similar naphthyliso-quinoline alkaloids, like it can be seen in Fig. (2) of refer-ence [26]. In this context, the signals registered at 1356 and 1613 cm-1 assigned to C=C stretching and C–H bending vi-brations were especially useful for the reliable distinction of the different alkaloid structures [27]. Raman spectra ob-tained from the individual pure alkaloid standards, as well as in vivo measurements on the plant tissue, were presented and discussed in detail. Most of the identified signals could be successfully assigned to various vibrational modes of the alkaloid molecular structures.

Tropane Alkaloids: Cocaine

Cocaine can be obtained from leaves of the coca plant (Erythroxylum coca Lam., Erythroxylaceae). FT-Raman spectra were recorded from a series of 33 solid mixtures con-taining cocaine, caffeine, and glucose (containing cocaine in amounts varying between 9.8 and 80.6 wt%) [28]. It was found that 98 % of the analyzed samples could be defini-tively classified according to their individual cocaine con-centration. Quantitative calibration models have been devel-oped using PLS (Partial Least Squares) algorithms, which allowed very precise predictions (RMSEP (Root Mean Squared Error in Probability)=4.1 %) of the cocaine content in the solid mixture. Cocaine was also investigated by SERS on silver tetrahydroborate and citrate colloids. A high-quality SERS signal was obtained at the concentration below 1 mg/mL [29].

Pyrrolidine Alkaloids: Nicotine

Tobacco alkaloids belong to the class of "true alkaloids" and are named after nicotine which has been detected in Nicotiana tabacum L. (Solanaceae) for the first time. Nico-tine is chiral and occurs naturally as S enantiomer, still often named (-)-nicotine. In the alkaloid fraction isolated from fresh leaves of Nicotiana tabacum, (S)-nicotine is present in

112 Current Analytical Chemistry, 2013, Vol. 9, No. 1 Baranska et al.

ca. 90 %, anatabine in ca. 4 %, (S)-nornicotine in ca. 2.5 % and anabasine in ca. 0.5 %. The other detected alkaloids (S)-N-methylanatabine, (S)-N-methylanabasine, myosmine, and anabaseine occur only in smaller amounts.

Early Raman measurements, started probably as far back as 1937 [30,31], were focused on establishing the spectra-structure relationships and registration of good quality spec-tra in a large spectral region [32,33]. A reference Raman spectrum of nicotine was published in mid 1970s [34]. In early 1980s, Raman spectrum was used to study the confor-mation of nicotine in solid state, aqueous solution and in interaction with a surface [35]. In mid 1990s, the nicotine SERS spectra on silver doped cellulose were used to detect its concentrations in water from 10-5 to 10-7 M [36], whereas on silver electrode the nicotine detection limit was estimated to be 7 ppb [37]. Now, the quantitative SERS analysis of nicotine at ppm/ppb levels can be done rapidly using the 1030 cm-1 band and stable and inexpensive polymer-encapsulated silver nanoparticles [37,38].

Molecular structures of nicotine, nornicotine, cotinine, anabasine were investigated using the Raman spectra and DFT calculations [39]. For all tobacco alkaloids the marker bands were selected and assigned. For the first time, the Ra-man Optical Activity (ROA) spectrum of (-)-nicotine in wa-ter solution was measured and interpreted based on DFT calculations. A good agreement between the experimental and simulated (B3LYP/aug-cc-pVDZ) ROA spectra were shown, for example, to identify two trans nicotine conform-ers to be present in the solution. This demonstrates a high efficiency of the ROA technique in detailed analyses of the chiral tobacco alkaloids. Additionally, an analysis of se-lected pharmaceutical products performed by Raman map-ping confirmed that the active substance in the pharmaceuti-cal tablet is a complex of nicotine with methacrylic acid whereas that in the plaster is the neutral nicotine in the trans conformation. The plant analysis showed an elevated distri-bution of nicotine near the edge of tobacco leaves [39].

Stimulated Raman scattering from liquid nicotine, to-gether with spontaneous Raman spectra, has been recorded [40]. The low frequency region was of special interest in that study. Raman intensity maxima occurred near 303, 260, and 115 cm-1, with the half-widths ranging from 15 to 30 cm-1. Such relatively sharp Raman bands almost certainly arise from intramolecular deformation or torsional oscillations of nicotine. Close inspection of the spectrum below 100 cm-1 revealed an inflection near 65–75 cm-1 and an accompanying wide region of downward concavity. These features may result from intermolecular effects. From the intensity, posi-tion, and half-width of the low-frequency Raman band it was possible to conclude that the band arises from intermolecular effects, a restricted rotational or vibrational motion of the nicotine molecule, or the closely related collisional mecha-nism. Of the two intermolecular mechanisms, the first has been strengthened by the observed good correlation between the integrated Raman intensity and the quantity. The spectral similarities between nicotine and pyridine below 100 cm-1 indicate that a restricted rotational motion is also reasonable as an explanation for the low-frequency nicotine band. The nicotine SERS spectrum on a silver–alumina substrate re-veals a sharp band at 1032 cm-1 (trigonal ring breathing), and

several broad bands similar to those for nicotinamide [41]. This feature is supported by the interaction of nicotine with the surface through the pyridine ring, while the characteristic five-membered pyrrolidine ring of the alkaloid stays far from the surface.

Piperidine Alkaloids: Piperine

Piperine is the main pungent in the green berries of pep-per (Piper nigrum L., Piperaceae). FT-Raman spectra ob-tained from green pepper berries, ground black pepper, and black pepper oleoresin predominantly show significant key signals of piperine [42]. Apart from the intense –C–H stretching vibrations between 2800 and 3100 cm-1, the main Raman signals occur in the fingerprint range between 1100 and 1630 cm-1. The aromatic and aliphatic –C=C– as well as –N–C=O stretching vibrations can be detected between 1580 and 1635 cm-1. The signals observed at 1448 cm-1 can be assigned to –CH2 bending vibrations, whereas the other bands in the range between 1100 and 1400 cm-1 are mainly due to –C–C– stretching (1153 cm-1), as well as –CH2 twist-ing and rocking vibrations (1295 and 1256 cm-1) of piperine molecules. Based on the individual key bands detected in the range between 1580 and 1635 cm-1, the distribution of piper-ine in a peppercorn can be analyzed in situ applying FT-Raman microscopic mapping. According to these measure-ments the pungent principle is distributed more or less in the whole perisperm of the green fruit; only the endosperm, lo-cated in the center of the berry, contains lower amounts of piperine. Applying ATR-IR and Raman measurements, chemometric equations have been developed for the calibra-tion of piperine content in pepper samples, presenting a comparatively high prediction quality (R2=0.86 and 0.84, respectively) [42].

3. CAROTENOIDS

3.1. Definition of Carotenoids

Carotenoids are a class of hydrocarbons (carotenes) and their oxygenated derivatives (xanthophylls). They may be formally derived from the acyclic lycopene C40H56 structure ( , -carotene, Scheme 4) modified by (i) hydrogenation, (ii) dehydrogenation, (iii) cyclization, (iv) oxidation, or any combination of these processes [43].

3.2. Biosynthesis and Function of Carotenoids in Plants

3.2.1. Carotenoid Biosynthesis

All known natural carotenoids are synthesized in 11 en-zyme-catalyzed carotenoid biosynthesis pathway [44-46] with some minor variations among species [47,48]. The first two steps are condensations of C40 carotenoid, 15-cis-phytoene, from two C20 compounds each formed from two C10 ones. Next, phytoene, infrequently accumulated in the plant tissues, is symmetrically desaturated to form di-cis- -carotene. Then, -carotene is once again desaturated to tetra-cis-lycopene and next isomerized to all-trans-lycopene [49].

From now on, the carotenoid biosynthesis pathway is split into the symmetric , - and asymmetric , -branches leading to - and -carotene, respectively. The rings are formed due to action of lycopene- -cyclase ( LCY) which

Recent Advances in Raman Analysis of Plants Current Analytical Chemistry, 2013, Vol. 9, No. 1 113

also co-operates with lycopene- -cyclase ( LCY) in forma-tion of -carotene.

Xanthophylls are formed from - and -carotene by hy-droxylation. Single hydroxylation of - and -carotene at the C3 atom leads to zeinoxanthin and -cryptoxanthin, respec-tively. Zeaxanthin is formed by the subsequent hydroxyla-tion of the other -ring of -cryptoxanthin, while lutein is produced by the hydroxylation of the ring of zeinoxanthin.

In the , branch, zeaxanthin is epoxidased to antherax-anthin and next to violaxanthin, yet, deepoxidase reverses this reaction in intense light. The last carotenoid of the , branch, 9-cis-neoxanthin, is converted from 9-cis-violaxanthin by a synthase. Both violaxanthin and neoxan-thin can be cleaved by a dioxygenase to ultimately produce the abscisic acid.

3.2.2. Light Harvesting, Photooxidative Protection, and Color Determination Functions

Light Harvesting

The major functions of carotenoids in plants, algae and purple bacteria are connected with photosynthesis which is a delicate interplay between light absorption, excitation energy transfer, dissipation of excess energy and electron transfer processes. Carotenoids in photosynthesis primarily play a: 1) light-harvesting role in the antenna complexes of the chloro-plasts and 2) protecting agent role against the harmful pho-tooxidative effects caused by bright light.

Plants, algae, and purple bacteria absorb light predomi-nantly using chlorophyll or bacteriochlorophyll, but the other pigments, such as carotenes and xanthophylls, are also in-volved in the phothosyntesis. Carotenoids are embedded in antenna-proteins forming light-harvesting complexes [50], which surround a photosynthetic center to focus photon en-ergies absorbed by the pigment, and to transport the excita-tion energy to the reaction center where it is converted into chemical energy driving the photosynthetic reaction [51].

Photooxidative Protection

When the absorbed high-light exceeds the capacity of the reaction center, photosynthetic organisms must dissipate the

excess energy to reduce dangerous byproducts. The dissipa-tion proceeds in non-photochemical quenching by carotenoid radical cations formed in the light harvesting complex. The cation radicals are scavenging harmful singlet oxygen and are quenching triplet states of intermediates always accom-panying to photosynthesis [52-54].

Plant Color Determinants

Carotenoids present in chromoplasts, give plants, algae, fungi and bacteria a characteristic yellow-red-orange color. Plant colors are addressed to insects, birds, and bats, which bring pollen grains to effect the pollination and thus the con-tinuation of the species [55]. So, plants must be visible from a distance for airborne pollinators, and must stand out from their surroundings. Carotenoids absorb at 400–500 nm de-termining color of some flowers (roses and marigolds), fruits (tomatoes), roots (carrot) and seeds (red peppers). Possibly, the autumn tree colors show insects strength and readiness of a plant to fight and that, in case of parasite attack, it may respond with a harmful substance [56,57].

3.3. Vibrations of Carotenoids

Carotenes are built from a poliene chain and cyclic end groups. Among linear polyenes, the most stable isomer of 1,3-butadiene and 1,3,5-hexatriene have the planar s-trans structure belonging to C2h symmetry [58-62]. Changes of the all-trans- , -dibutylpolyene Raman spectra are related to the number of the conjugated double bonds (NC=C) increasing from 3 to 12 and are reflected in four bands called 1, 2, 3 and 4 [63]. The 1 band is assigned to the in-phase C=C stretch and is 190 cm-1 downshifted as NC=C increases. The

2 band is assigned to a mixture of the C-C and C=C stretches of frequency insensitive to NC=C. The 3 and 4 bands are assigned to mixtures of CH in-plane bend as well as C=C and C-C stretches. In a series of unsubstituted polyenes, the intensity of the 3 band is much larger than that of the 4 band and oppositely in , -dibutyl substituted polyenes. This indicates that the 3 and 4 mode mixing is substitution dependent. The polyenes formed by polymeriza-tion of acetylene (trans-polyacetylenes), and consisting of all-trans NC=C conjugated segments with various conjugation

Scheme 4. A simplified carotenoid pathway restricted to the most abundant pigments in the plant kingdom.

114 Current Analytical Chemistry, 2013, Vol. 9, No. 1 Baranska et al.

lengths, give rise to the Raman bands similar to those of polyenes [60].

The resonance Raman spectra of -carotene and its syn-thetic homologs C30H44, C35H50, C50H68, and C60H80 of bixin, crocetin, capsanthin, the polyene antibiotic amphotericin B (fungizone), and of deuterio- -carotene showed that the fre-quency of the intense ethylenic mode at -1500 cm-1 is a genuine measure of the stiffness of the C=C bonds hence the frequency of the 1 mode is an indicator of bond alternation [61]. Changes of the observed Raman frequencies with in-creasing NC=C in unsubstituted polyenes [62], , -dibutylpolyenes [63], carotenoids and -carotene [63], rhodovibrin [64], spirilloxanthin [64], decapreno- -carotene [61] and dodecapreno- -carotene [65] have already been presented. The maximum of 11Bu 11Ag electronic absorp-tion in polyenes shifts to longer wavelengths as NC=C in-creases. Thus, the Raman spectrum taken with a red laser line provides Raman bands arising from a long segment, whose conjugation length is not accurately determined. In the Raman spectrum taken with the 632.8 nm excitation laser line, the bands observed at 1457, 1294, 1174 and 1066 cm-1 are clearly related to the infinite all-trans structure.

It has been also reported that infrared frequencies of the CH out-of-plane bending vibrations of olefins are sensitive to the number and position of hydrogen atoms attached to the C=C bond. The in-phase CH out-of-plane bending vibra-tions are observed in the range between 1009 and 1015 cm-1: 1009, 1011, 1010 and 1015 cm-1, for trans-hexatriene, trans, trans-1,3,5,7-octatetraene, all-trans-1,3,5,7,9-decapentaene and trans-polyene, respectively [60].

For the infinite all-trans structure, the 1174 cm-1 band is assigned to an optically inactive mode whereas the other four bands correspond to optically active ag ones. The un-usually low frequencies of the 1 bands are observed for the ground state (1'Ag), whereas for 2'Ag electronically excited state, unusually high frequencies of the corresponding modes are observed. This is due to the vibronic coupling between the 11

Ag ground and 21Ag excited state. The observed 1 fre-

quencies have been fitted by equation of the following gen-eral form:

AN

Kcm

CC+

+=

=

)( 1

1 (1)

where for trans-polyene A=1, = 1454 cm-1 and K = 727 cm-1, and for , -dibutylpolyenes (NC=C=7-12) A=0, = 1438 cm-1 and K = 830 cm-1 [66]). Thus, having the observed

1 frequency it is possible to make an estimation of the num-ber of the conjugated double bonds NC=C of the all-trans structures.

It has been recently shown by the B3LYP/6-311++Gcalculation that the position of the 1 band in C1–C15 polyenes as a function of the chain length (Cn), converges in the infinity to ca. 1495 cm 1 and the Raman activity of this band has been shown to increase rapidly if the number of double bonds exceeds nine [67].

The vibrational spectra of -carotene were registered and utilized in a very large amount of studies, however, system-atic theoretical interpretation of the vibrational spectra of -

carotene has been the subject of only a few papers [58-61]. In the first detailed theoretical IR and Raman study the nor-mal-coordinate calculations were performed by Saito and Tasumi for the all-trans and 15-cis isomers of -carotene [68]. Also, the bands of 7-cis, 9-cis, 13-cis, 9,13-di-cis, 9,13-di-cis, 9,15-di-cis and 13,15-di-cis -carotene isomers were assigned [68]. The -carotene, zeaxanthin, canthaxanthin, and astaxanthin spectra were then analyzed at the semiem-pirical AM1 level by Weesie et al. [69]. Based on the AM1 calculated excitation energies resonance and non-resonance Raman spectra of astaxanthin and 13C labeled astaxanthins as well as asymmetric 20-norastaxanthin were recorded and interpreted [69].

In 2003, the IR and Raman spectra of -carotene of Ci symmetry were interpreted by means of DFT/BPW91/6–31G* calculations [70]. It was found that, at the calculation level applied, the s-cis- -carotene (where the -ionene rings are twisted and the ring double bonds are in sterically fa-vored s-cis arrangement with respect to the polyene chain) is 8.8 kJ mol 1 more stable than the s-trans isomer (where the ring double bonds are in s-trans positions to the polyene chain) [70]. In 2005 the IR spectra calculations for all-trans-

-carotene (s-trans conformer) and its 15,15 -cis-isomer were performed at the DFT (B3LYP/6-31G*) level and in-terpreted by means of PED (Potential Energy Distribution) analysis [71]. The necessity for calculations of the whole -carotene molecule with -ionone rings for correct interpreta-tion of the vibrational IR spectra was stressed [71].

The Raman spectra of -carotene shows three prominent vibrational bands named 1, 2 and 3 (at 1524, 1157 and 1005 cm 1, respectively), which are attributed to the double bond C=C stretching mode (1524 cm 1), the C-C in-plane single bond stretching mode (1157 cm 1), and the C-H bend-ing mode (1008 cm 1), respectively. Recently, the B3LYP/6-31G* calculations have predicted two closely spaced 1 modes of -carotene [72]. The 1a mode is predicted at 1535 cm 1 and 1b mode at 1528 cm 1, whereas they are present in the experimental spectrum at 1525 cm 1 and 1519 cm 1, re-spectively. The former exhibits twice of the intensity of lat-ter. According to the DFT calculations both the 1a- and the

1b-mode originate from double bond C=C stretching of the polyene chain, although different C=C stretching coordinates are involved. The two bands have been shown to exhibit different excitation profiles: the shift to lower wavenumbers upon excitations from 2.3 to 2.5 eV is supposed to be due to the higher contribution of the 1b band, whereas the apparent upshift in the region around 488 nm can be attributed to the higher contribution of the 1a band [72].

The excitation profiles of three lycopene fundamentals 1, 2, and 3 in resonance Raman spectrum were examined

in various solvents in terms of two- and three-mode vibra-tional models [73-75]. Quite recently, the Raman spectra of all-trans and 13-cis lycopene encapsulated in cholesteric polyesters have been measured and interpreted by using the DFT/B3LYP/6-31G* method [76]. It has been shown that the polyesters induced conversion of the trans isomer into a cis one with the cis C=C bond in central positions of the isomer chain. It has been suggested that the conversion mainly leads to the 13-cis lycopene and that the polyesters revealed changes due to the ester groups showing a recon-

Recent Advances in Raman Analysis of Plants Current Analytical Chemistry, 2013, Vol. 9, No. 1 115

struction of the polyester chain accommodating lycopene [76].

The resonance Raman excitation profiles for astaxanthin were studied experimentally in early 1970s and it was dem- onstrated that there is a correlation between the development of vibrational structure in the absorption spectrum and in the excitation profile [77]. Raman spectra of two crystal forms of astaxanthin can be found in Refs. [78 and 79].

As already pointed out in 1985 [80] the resonance Raman spectra of carotenoids vary appreciably with cis-trans iso- merisation. When compared to the all-trans isomer, the order of increasing frequency of the 1 band correlates well with the blue shift of the absorption band caused by the chain perturbations. Generally, the centrally bent-isomers show larger Raman shifts than peripherally-bent isomers. The 2 region is very sensitive to the configuration of the chain. The spectral pattern is unique for each of the isomers studied and can be ascribed to changes in the mixing of the C-C stretch-ing vibrations and no 1 vs. 2 correlation is obtained.

3.4. Raman Spectroscopy of Carotenoids in Plants and Fungi

The reliability of the carotenoid analyses was compre-hensively reviewed in Current Analytical Chemistry in 2005 by Feltl et al. [81]. In that review, the most important prop-erties of modern techniques applied to analyses for caro-tenoids and their impact on the reliability of the analytical results were critically discussed. Several examples of ana-lytical procedures illustrating the characteristics of modern experimental approaches to carotenoid analyses as well as to various sample pretreatment and preconcentration methods, such as microwave-assisted extraction or supercritical fluid extraction were presented. The problems related to the chromatographic purification of carotenoids and getting reli-able standard materials were also dealt with. The focus was put on HPLC, capillary electrochromatography, detection techniques, UV/VIS spectrophotometry, electrochemistry, thermal lens spectrometry, mass spectrometry, NMR, and some aspects of vibrational spectrometry. Although in the Feltl et al. review little was reported on Raman spectro-scopic analyses of carotenoids in plants, in the presented here review the papers published before 2005 are discussed only rarely. Hereafter, the subchapter is dedicated to plant materials investigated by Raman spectroscopy such as vari-ous trees, flowers, herbs, bushes, grasses, vines and ferns, but also green, and red algae, fungi, and lichens. Addition-ally, also some aspects of Raman spectroscopy applied to bacteria identification is discussed.

3.4.1. Higher Plants

In 2007, a thorough review of the potential of Raman mi-croscopy and Raman imaging in plant research was pub-lished by Gierlinger and Schwanninger [82]. They stressed the role of preresonance intensity enhancement effect ac-companying the FT-Raman measurements of carotenoids enabling for detection of minute amounts of analyte with simultaneous elimination of the backgound increase pro-duced by fluorescence of the sample. They also emphasized that the position of C=C stretching vibrations is mainly in-fluenced both by the carotenoids polyene chain length and

terminal substituents, which however, are somehow convo-luted with their interactions with other plant constituents. Indeed, these advantages of FT-Raman spectroscopy applied to 2D Raman mappings of carotenoids in plants enabled to show in 2005 that the distribution of individual carotenoids (7-, 8-, and 9-double bond conjugated carotenoids) can be analyzed independently in the same sample [83]. Moreover, a high potency of Raman spectroscopy for in situ detection of unstable epoxycarotenoids was indicated. FT-Raman mi-crospectroscopy was applied to obtain detailed information on microstructure and chemical composition of fennel fruits, chamomile inflorescence and curcuma roots [84]. The caro-tenoids, monitored by observation of the 1525, 1156 and 1004 cm-1 bands, were shown to be located at the top of the flower heads. Interestingly, the light microscope image of this very region showed anthers full of pollen, whereas empty anthers did not contribute to the carotenoid signals (see Fig. 3 in ref. [84]).

For various living plant tissues of the size ranging from about 0.01 mm2 to 35 cm2 a heterogeneous distribution of carotenoids and changes induced by abiotic and biotic stresses were also monitored in situ by the NIR-FT-Raman imaging [85]. For example, in a pelargonium leaf the distri-bution of total carotenoids by using the 1525 cm-1 band and separately lycopene and -carotene by using the bands at 1510 cm-1 and 1524 cm-1, respectively, was demonstrated.

In a tomato (Lycopersicon esculentum Mill.) fruit, Ra-man mapping visualized the inhibition of lycopene biosyn-thesis and accumulation of -carotene in the tissue affected by sunscald physiological disorder [85]. A local carotenoid decline at infection site was displayed by Raman map of diseased sugarbeet (Beta vulgaris L.) leaf whereas the caro-tenoid accumulation was shown to be an evident response to Septoria petroselini infestation in parsley (Petroselinum crispum Mill. Nym.). Variation of lutein, -carotene and capsanthin content during bell pepper (Capsicum annum L.) fruit ripening were checked by single Raman spectra [85].

Only lately, lycopene, -carotene, and lutein in tomato have been analyzed and distinguished from each other by confocal Raman microspectroscopy [86]. Both, the physical state of carotenoids (crystalline or solvated) and the effect of processing on carotenoids in tomatoes were under study. It is emphesized that this type of information can be of great benefit to rationalize food processing and to define the state in which foods should be presented to the body to maximize uptake [86].

It was demonstrated that, without any preliminary sample preparation, carotenoids can be determined in cultivated and wild carrots (Daucus carota L.) simultaneously with polya-cetylenes (falcarinol and falcarindiol), pectin (polysaccha-ride with the characteristic -C-O-C- skeletal mode of alpha-anomer carbohydrates band at 854 cm 1) and starch (detected at ca. 480 cm 1) [87]. Moreover, the simultaneous in situ measurements of relative content and distribution of -, -carotene, lutein, and lycopene were performed in carrot root sections [88]. Such measurements were possible due to the ability to separate different Raman bands to -carotene (1520 cm 1), lycopene (1510 cm 1) and -carotene/lutein (1527 cm 1). Clearly, the level of -carotene was heteroge-neous across root sections of orange, yellow, red and purple

116 Current Analytical Chemistry, 2013, Vol. 9, No. 1 Baranska et al.

carrot roots. In the secondary phloem -carotene increased gradually from periderm towards the core, but declined fast in cells close to the vascular cambium. On the other hand, -carotene/lutein were deposited in younger cells with a higher rate than -carotene while lycopene in red carrots accumu-lated throughout the whole secondary phloem at the same level. The results support the concept of developmental regulation of carotenoid genes in carrot root and showed the potency of the Raman spectroscopy to supply essential in-formation on carotenogenesis useful for molecular investiga-tions on gene expression and regulation.

The potency of the Raman mapping combined with clus-ter analysis have recently been shown by non-destructive FT-Raman direct in-situ discrimination of flavonoids, antho-cyanins, and carotenoids occurring in differently colored flower petals of pansy cultivars (Viola x wittrockiana) [89]. For the sake of comparison, the pigment extracts of the pet-als were separated by thin-layer chromatography (TLC) and investigated by classical Raman measurements. Hierarchical cluster analysis of the reference pigments spectra taken from the flower petals and from the TLC extracts allowed dis-crimination of flavonols from anthocyanins, and the two of them from carotenoids. Relative concentration of these com-pounds in petals was successfully determined by the Raman mapping supported by cluster analysis. The correlation be-tween the distribution patterns observed in visible and Ra-man mapping images was very satisfactory [89].

Quite recently, the first report on direct in situ Raman microspectroscopic detection of carotenoids at the subcellu-lar level has been published [90]. Single carotenoid crystals sequestered in a carrot cell were measured by using an FT-Raman spectrometer equipped with a microscope and 40 objective. The observed characteristic bands centered at 1518 cm 1 and 1156 cm 1 proved the crystals to be predomi-nantly formed from -carotene. Thus, the current Raman microspectroscopy techniques allow for identification and analysis of compounds localized in cytoplasm by taking measurements directly from a single plant cell [90].

The nonlinear coherent anti-Stokes Raman scattering (CARS) microscopy have recently been introduced by Brackmann et al. for 3D imaging of carotenoids in fresh and thermally treated plant tissues [91]. The CARS microscopy measurements of carotenoids were done for orange-fleshed sweet potato, carrot, and mango, and in the case of potato they were related to the plant-matrix morphology by simul-taneous second-harmonic generation microscopy of starch granules. This has given the opportunity for quantification of sizes, shapes, densities, and location of different types of carotenoid bodies. Heterogeneous rod-shaped bodies with high carotenoid densities were indicated for potato and car-rot by the CARS signals, whereas the carotenoid-filled lipid droplets in mango appeared as homogeneous low-density aggregates of rounded shape. Interestingly, it was found that after thermal processing of the potato, -carotene density and morphology remain intact despite of significant changes of the surrounding starch granules [91].

The non-destructivity of the classical FT-Raman meas-urements are still being explored by many laboratories. For example, the presence of FT-Raman bands at 1520, 1157, and 1007 cm 1 in the spectra of rose hips were assigned to

C=C, C–C, and C–CH3 stretching vibrations bands of -carotene as the main constituent, however, the presence of a C9 carotene was also suggested [92]. On the other hand, in-side the seed the Raman spectra showed the presence of un-saturated fatty acids and fatty products comprising cis iso-mers. The classical FT-Raman measurements of a range of naturally occurring carotenoids in over 50 specimens of plant tissue were performed in the same laboratory [93]. Based on a range of standard extracts, they showed that there may be a serious problem in the interpretation of the spectro-scopic data which can be attributed to significant wavenum-ber shifts caused by carotenoid interactions with organic tissues. On the other hand, the potential of ATR-FTIR (At-tenuated Total Reflectance FTIR) and Raman spectroscopies was applied to evaluate changes in olive skin, flesh and stone during different stages of fruit development and matu-ration [94]. Besides carotenoids, triglycerides, water, and phenolic compounds were monitored by evolution of charac-teristic bands. Although the authors showed that the oil ac-cumulation can be followed using both FTIR and Raman spectroscopy, the increase of the content in carotenoids and phenolic compounds with olive growing and their decrease during fruit ripening can be successfully monitored by means of the Raman bands at 1525 and 1605 cm-1, respec-tively [94].

Raman spectroscopy has been also used for chemical characterization and discrimination of pollen grains of dif-ferent trees species [95-97]. Out of different excitation laser wavelengths, the red He-Ne laser yielded high-quality single pollen Raman spectra with several bands assignable to such pollen components as carotenoids, proteins, nucleic acids, carbohydrates, and lipids. The multivariate classification based on principal component and hierarchical cluster analy-ses demonstrated the validity of the approach for discrimina-tion between different pollen species [95]. It was also shown, that in contrast to purification-based analyses, the nondestructive Raman spectroscopy approach allows (i) to analyze various classes of molecules simultaneously at mi-croscopic resolution and (ii) to acquire fingerprint-like chemical information that was used for the classification of pollen from different species. In this way, it was possible to obtain a Raman signature supporting classification in accord with biological systematics and introduce chemistry-based online pollen identification methods [96]. An interesting study was devoted to Raman signatures of the carotenoid component of individual pollen grains from different species of trees obtained as differences in the strong pre-resonant Raman spectra measured before and after photodepletion of the carotenoid molecules [97]. The in situ evidence of inter-species differences in content of pollen carotenoids, pro-vided without prior purification, was confirmed by reso-nance Raman spectra taken from the HPTLC plates used for separation of carotenoids in pollen extracts. Thus, the in situ, extraction-free routine of carotenoid analysis enhance sensi-tivity and structural selectivity of single pollen grains ex-amination. Quite recently, the silver nanoparticles have been generated in citrate reduction of the sporopollenin bio-polymer of ragweed (Ambrosia artemisiifolia) and rye (Se-

cale cereale). Based on these very nanoparticles, the acquisi-tion of the SERS spectra and vibrational characterization of

Recent Advances in Raman Analysis of Plants Current Analytical Chemistry, 2013, Vol. 9, No. 1 117

the sporopollen in local molecular structure has become pos-sible [98].

The Raman spectroscopy, faster and cheaper than classic chemical analysis, has been also used for determination of carotenoids in plants for industrial purposes [99]. The caro-tenoid Raman 1524 cm 1 band was observed for fresh ligules and for ligule powder of four Tagetes erecta L. varieties: white, cream-colored, yellow and orange, whereas the total xantophyll concentration was determined by chemical analy-sis on dry ligules. By relating the two measurements by ex-ponential models, a rapid and inexpensive quantitative Ra-man analysis was prepared. Last but not least, the Raman spectroscopy is continuosly being applied for characterisa-tion of new carotenoids isolated from plants [100].

3.4.2. Algae

Algae is a diverse group of organisms in which tissues are not organized into distinct organs as found in higher plants. Usually, they are photosynthetic, autotrophic, unicel-lular or multicellular organisms. The green algae (prokary-otic cyanobacteria) are nowadays classified as bacteria, and thus the term algae is now restricted to Eukaryota, however in this review, both eucariotic and prokaryotic algae are dis-cussed together.

Raman spectroscopy, microscopy, and imaging have been sucessfully applied to monitor the carotenoid distribu-tion in algae and physico-chemical changes in the carotenoid state [101-105], photosynthetic activity of algae [106-108], and to observe accumulation of different compounds in algal cells [109, 110].

Astaxanthin, a non-provitamin A carotenoid, is the major carotenoid in marine animals and it is produced by a few organisms, like unicellular green algae (Haemaotococcus pluvialis), bacterium (Paracoccus) and yeast (Xanthophyl-lomyces dendrorhous-asexual stage of Phaffia rhodozyma), in a -carotene ketolase promoted reaction. Raman imaging has lately been shown to be a highly selective and sensitive method of in situ and in vivo monitoring of astaxanthin dis-tribution in the cyst form of H. pluvialis [101]. Moreover, the changes of astaxanthin structure upon thermal stress of H. pluvialis can be in situ analyzed for both, a single algal cell and a multicellular sample by Raman spectroscopy tech-niques [102]. Indeed, from -100ºC to 150ºC, a significant shift of the Raman 1 C=C band at ca. 1520 cm-1 is accom-panied by rapid increase of the 1190 and 1160 cm-1 band intensity ratio. These changes are in line with the DFT pre-dicted conformer population changes which have lead to the conclusion that astaxanthin is initially bound in the H-form aggregates (with the trans conformations of the end-rings), and, upon thermal stress of the algae, it interconverts into the more stable gauche forms [102].

A Raman spectroscopy characterization of carotenoid pigments in algal species was recently supported by multi-variate hyperspectral analysis. Although, the carotenoids mixture present in the studied algae did not fully match the reference spectrum, Raman measurements combined with chemometric techniques provided enough information to successfully perform the analysis [103]. The confocal Ra-man microscopy combined with multivariate analysis was used for monitoring of astaxanthin and -carotene in H. plu-

vialis. Use of chemometric techniques enabled to resolve the resonance–enhanced Raman signatures of astaxanthin and

-carotene from chlorophyll fluorescence. In turn, it was possible to locate the two carotenoids independently of each other in various life cycle stages of the living cells. Interest-ingly, chlorophyll was found only in the chloroplast, whereas astaxanthin was identified within globular and punctuate regions of the cytoplasmic space, and -carotene was co-located with both the chloroplast and astaxanthin in the cytosol [104].

The photosynthetic activity of the algae was studied by optical and electronic microscopy applied to Klebsormidium flaccidum algal cells encapsulated with gold intra-cellular nanoparticles within silica gels [105]. The microscopy tech-niques used indicated the entrapped cells to maintain their ability to reduce gold salts. The encapsulation process pro-tects the cells from lethal effects arising from gold toxicity. Moreover, the first in situ imaging of entrapped cells using Raman spectroscopy allowed the investigation of the influ-ence of the gold colloids on the photosynthetic system of the algae. It is suggested, that a coupling of sol–gel encapsula-tion and Raman imaging may account for development of novel photosynthesis-based cellular biosensors [105].

Different fucoxanthin chlorophyll complexes were de-tected in diatom Cyclotella meneghiniana alga using reso-nance Raman spectroscopy [106], yielding the strongest ever spectroscopic evidence that the number of pigments previ-ously estimated may even be doubled. The relative en-hancement of the resonance Raman bands for 406.7 versus 413.1 nm further allows some distinction of blue- versus red-absorbing chlorophylls, respectively [106].

The Dunaliella tertiolecta eucariotic algae, grown either under nutrient-replete conditions or starved of nitrogen for four days, were next analyzed by Raman spectroscopy methods to monitor changes in chlorophyll a and -carotene concentrations [107]. The starved cells indicated a decline in chlorophyll a and an increase of -carotene in chlorophytes. This findings were supported by various chemometric meth-ods [107]. Different nutrition conditions were also applied for study on Chlorella sorokiniana and Neochloris oleo-abundans algae considered as candidates for biofuel produc-tion [108]. Raman signals due to lipids (triglycerides) were identified in nitrogen-starved algae and they were absent in healthy organisms, whereas the Raman signals of caro-tenoids were found to be present in all of the analyzed sam-ples [108]. Raman microspectroscopy was also used for in vivo predicting of the nutrient status of individual algal cells [109]. The use of 780 nm laser excitation line revealed en-hanced bands attributable to chlorophyll a- and -carotene which, under nitrogen limitation, became less intense and more prominent, respectively. Although spectra from N-replete and N-starved cell populations varied, each distribu-tion was distinct enough to be accurately predicted by Ra-man spectroscopy combined with chemometric techniques [109].

3.4.3. Lichens and Fungi

Lichens are symbiotic organisms composed of a fungus with a photosynthetic partner usually either green alga or cyanobacteria. Sixteen lichen specimens growing on new

118 Current Analytical Chemistry, 2013, Vol. 9, No. 1 Baranska et al.

basaltic lava fields and wood substrates have been recently analyzed by Raman spectroscopy [110]. Among the pig-ments and biomolecules produced in extreme environment, the Raman spectroscopy analysis indicated presence of chlo-rophyll and a carotenoid (lutein or astaxanthin), but a range of protective pigments, such as atranorin, usnic acid, gyro-phoric acid, parietin and pulvinic acid dilactone, have been identified too. These findings enabled authors to conclude that several survival strategies were adopted by lichens to combat radiation insolation, desiccation and high tempera-tures [110].

The Raman spectroscopy was also used to detect caro-tenoids in fungi themselves. In 2009, the intracellular moni-toring of carotenoids in filamentous fungi Blakeslea trispora was performed by using the strongest Raman 1 carotenoid band at 1519 cm 1 [111]. A good linear correlation between the carotenoid content determined by Raman, UV-Vis, and HPLC techniques was shown but Raman analysis was not as accurate as HPLC. However, the Raman method allowed a rapid sampling and analysis of carotenoids thus avoided the prolonged and tedious classical isolation procedures required for carotenoid determination by HPLC and UV-Vis spec-troscopy [111].

Rhodotorula glutinis is a pigmented yeast which is known to accumulate large amounts of carotenoids under certain culture conditions [112]. The molecular mechanism of regulating carotenogenesis in yeast is still not well under-stood. To directly detect carotenoids in single live R. glutinis cells, the authors developed the laser tweezers Raman spec-troscopy. It was found that the carotenoid and lipid accumu-lation occurred mainly when the cell growth was inhibited by nutrient limitation. Moreover, the carotenoid concentra-tion changed together with the concentration of nucleic ac-ids, which increased in the first phase and decreased in the last phase of the culture [112].

4. POLYACETYLENES

4.1. Polyacetylene Definition

Polyacetylenes are a class of bioactive secondary plant metabolites that contain at least two, usually conjugated, triple carbon-carbon bonds in their structures. However, the most common are diacetylenes with two conjugated triple bonds such as falcarinol (panaxynol), falcarindiol and panaxydol (Scheme 5). Nowadays, the number of known natural polyacetylenes reaches thousands [113-115].

4.2. Synthesis, Biosynthesis, and Function in Plants

The isotopic studies revealed that fatty acids and polyketides (secondary metabolites of bacteria, fungi, plants, and animals) are precursors of most of the natural polyacety-lenes [116]. The first isolated natural alkyne was tariric acid CH3(CH2)10C C(CH2)10COOH. It was obtained from seeds of Brazilian Picramnia tariri DC plant (Simaroubaceae) in 1892 [117, 118].

The first synthesis of diacetylene compounds was described by Carl Glaser and was called Glaser coupling [119]. Later, diiododiacetylene and some other diacetylenes were synthesized by Bayer [120]. Nowadays, diacetylene

syntheses are performed using the Cadiot-Chodkiewicz cou-pling, in which the terminal acetylene group is coupled with a terminal halogenoacetylene group [121, 121]. The Cadiot-Chodkiewicz coupling has been also used to synthesize di-acetylenes with complex organosilocon substituents [122, 124]. Presently, the modern diacetylene syntheses allow to substitute this compound by a large number of groups in-cluding metals [125-131]. For instance, dimetal diacetylenes such as dilithium derivative have been shown to be of use in obtaining polymers [131-133].

Naturally occurring polyacetylenes are synthesized by plants as important protective compounds against insects, fungi and diseases. They are formed from unsaturated fatty acids and are built up from acetate and malonate units. Fal-carinol-type polyacetylenes are synthesized from oleic acid by dehydrogenation leading to the C18-acetylenes crepenynic acid and dehydrocrepenynic acid that is then transformed to C17-acetylenes by -oxidation. Further oxidation and dehy-drogenation leads to obtaining falcarinol and related C17-acetylenes of the falcarinol-type [134].

The localization of polyacetylenes in exterior tissue lay-ers is consistent with their role in providing an antifungal shield for young roots. These compounds play an important role in plant defense against, e.g. predators, fungi and other plant pathogens. Polyacetylenes are also recognized to be phytoalexins, low molecular weight compounds produced by plants to respond to microbial attack, disease state, or abiotic stress (e.g., UV irradiation, metal salts, detergents) [116]. For instance, accumulation of suspected polyacetylenic phy-toalexins, including falcarinol, was observed in tomato fruits and leaves induced with Cladosporium fulvum, Verticillium albo-atrum, and F. oxysporum [116]. Fungal stimulation of polyacetylene accumulation has been seen in some plant families such as Asteraceae, Fabaceae, and Apiaceae. Polya-cetylenes have been also shown in a number of cases to be allelochemicals, compounds that affect the metabolism and growth of plants subacutely, e.g. agrocybin strongly inhibits the growth of wheat, soybean, while dehydrofalcarinol was identified as one of the seed germination inhibitory compo-nents from Artemisia capillaries. Furthermore, several ex-amples of plant-derived polyacetylenes involved in antiher-bivory and insecticidal activities depending on the light con-ditions have been reported, e.g. thiarubrine was toxic to mosquito larvae and tobacco hornworm under dark condi-tions, whereas the toxicity of a thiophenic diacetylene to the A. atropalpus larvae became apparent under near-UV irra-diation. In a limited number of cases, polyacetylenic natural products from plants have been also shown to act as allo-mones, i.e. transspecific chemical signals that induce a be-havioral change in another species beneficial to the producer [116].

4.3. Occurrence, Physical and Chemical Properties, and

Application

Natural polyacetylenes are widely spread in plants from Apiaceae family [135], i.e. in carrot roots [136-138], parsley (Petroselinum crispum L.) [138], celery (Apium graveolens L.) [137], from Asteraceae family such as Chrysoma pauci-flosculosa [139], Echinacea purpurea L. Moench. and Echi-nacea pallida L. Moench [140,141], Acroptilon repens L.

Recent Advances in Raman Analysis of Plants Current Analytical Chemistry, 2013, Vol. 9, No. 1 119

DC [142], as well as in plants from East Asia like ginseng (Panax ginseng C.A. Mey) [143-147] Angelica dahurica Fisch. ex Hoffm., Araliaceae Juss. [148-149], and Centella asiatica L. Urban [150]. Furthermore, aliphatic C17-polyacetylenes of the falcarinol-type have been also detected in plants from Solanaceae family such as tomatoes (Solanum lycopersicum) and aubergines (Solanum melongena) [151-153].

Polyacetylenes are not basic components in plants. They occur in low concentrations, e.g. the concentration of polya-cetylenes in carrots ranges from 20 to 100 mg kg-1 fresh weight (FW) [154]. According to Czepa et al. [155], the most abundant polyacetylene in carrot roots is falcarindiol with a concentration range from 16 to 84 mg kg-1 FW. Fal-carinol and falcarindiol 3-acetate (Scheme 5) are present in somewhat lower concentrations, 5-31 and 7-41 mg kg-1 FW, respectively. Moreover, the polyactylene content depends on cultivar and, for instance, yellow carrot roots contain less polyacetylenes than orange ones. Natural polyacetylenes were identified in ginseng roots that have been used in tradi-tional Asian medicine for the past 2000 years [156]. In gen-eral, the concentration of falcarinol in this plant has been determined to be between 200 and 560 mg kg-1 dry weight (DW), while panaxydol occurs at concentrations of 180-950 mg kg-1 DW [157,158]. However, Kitagawa et al. [159] re-

ported the concentration of falcarinol in fresh ginseng even in amounts of 700 mg kg-1 DW that is approximately 5 times higher than in carrots. In parsley, apart from falcarindiol (2 mg g-1 freeze dried tissue; FD) two other polyacetylenes were found: 8-O-methylfalcarindiol and panaxydiol (Scheme 5) in amounts of 0.4 and 0.1 mg g-1 FD, respectively, while falcarinol was not detectable. However, four polyacetylenes were detected in celery, i.e. falcarinol, falcarindiol, 8-O-methylfalcarindiol, and panaxydiol, with concentrations of 0.2-1.6, 2.1-4.6, 0.04-0.17, and 0.02-0.06 mg g-1 FD, respec-tively. Finally, fennel and parsnip contain two polyacety-lenes, i.e. falcarinol and falcarindiol, in amounts of 0.04 and 0.24 mg g-1 FD in fennel, respectively, and 1.6 and 5.8 mg g-1 FD in parsnip.

Simple diacetylenes, such as diacetylene [160,161], methyldiacetylene [162], cyanicdiacetylene [163-165], and methylcyanicdiacetylene, have been discovered in interstel-lar space. The other diacetylenes potentially observable in interstellar space (not discovered yet because of very small or zero dipole moment) such as isocyanodiacetylene [166], dicyanodiacetylene [167,168], isocyanocyanicdiacetylene [169], diisocyano diacetylene [170], as well as dicarbonyl diacetylene [171], have been synthesized in laboratory con-ditions.

Scheme 5. Structures of falcarinol-type diacetylenes and other exemplary mono- and polyacetylenes.

120 Current Analytical Chemistry, 2013, Vol. 9, No. 1 Baranska et al.

Diacetylenes must be handled with special care when performing experimental studies. Some of them explode or decompose after heating over their melting points or under light. Isolation of natural diacetylenes by preparative chro-matography is also difficult because, extracted from tissues, they either oxidize or decompose under light or pH change. Hence, only a few physicochemical properties of these com-pounds are established.

Diacetylenes are used for polymerization in solid state leading to polydiacetylenes constituted by alternating triple and double bonds in trans configuration [172,173]. The po-lymerization proceeds through 1,4-addition and can be in-duced by heat [174, 175], pressure, UV radiation [176, 177], or high-power radiation as well as interaction with some gases [177]. Under special conditions, perfectly conjugated macroscopic polymer monocrystals can be obtained by po-lymerization of diacetylenes [177]. The electric conductivity of polyacetylenes substituted by halogens has been already confirmed [178]. The substituted polydiacetylenes have still been tested for their potential use as conductive materials [179].

4.4. Vibrations of Polyacetylenes

Although the concentration of polyacetylenes in plants is in the order of 0.01-1.00 % fresh weight, depending on the individual species, they can be successfully detected by means of FT-Raman spectroscopy. Diacetylenes have two characteristic stretching vibrations of the –C C–C C– sys-tem, i.e. symmetric and antisymmetric. If the considered molecule has a symmetry centre, the symmetric mode will be active in Raman, while the antisymmetric one will be seen in IR. A very strong and polarized band due to –C C–C C– symmetric stretching vibration can be observed in a Raman spectrum in the region of 2200-2300 cm-1 [180-183]. Because only rare functional groups scatter in this very re-gion the presence of polyacetylenes in plant tissues can be clearly illustrated by in situ Raman mapping. Both, the num-ber of conjugated –C C– bonds in polyacetylenes and sub-stituents influence the number of Raman bands, their fre-quencies and intensities [182]. Thus, the shape of Raman spectrum in the region of ca. 2200 cm-1 allows recognition of the type of substituent as well as identification of the polya-cetylene type. For instance, in the Raman spectrum the most abundant polyacetylenes, i.e. falcarinol, falcarindiol, and panaxydol, have very similar characteristic bands, but shifted by a few cm-1. The band maximum for falcarinol ap-pears at 2258 cm-1 and for falcarindiol at 2252 cm-1 (addi-tional –OH group in the -position of the diacetylenic sys-tem) [182], whereas the Raman spectrum of panaxydol shows a band at 2260 cm-1 [182]. It has been noticed that some mono- and diacetylenes with aliphatic substituents have a very strong band at 2230 cm-1, but some of them show also a weak satellite band at about 2190 cm-1, which may be due to the 13C isotopic substitution of the triple bonds. Although the Raman spectrum of the unsubstituted triacetylene showed two very strong bands, at 2212 and 2019 cm-1, due to the in-phase stretching vibration of all three tri-ple bonds and the out-of-phase vibration of the same bonds, the strongest bands in the spectra of some triacetylenes have been shown in the region of 2190–2118 cm-1

[182]. In the case of tetraacetylenes, two symmetric and antisymmetric –

C C– modes occur, while for diacetylenes substituted by a substituent of mass similar to that of the –C C– moiety, several additional couplings occur and the spectral picture in the (C C) region is more complicated [184].

4.5. Raman Spectroscopy of Polyacetylenes in Plants

It has been shown that apart from easy detection of poly-acetylenic compounds, it is also possible to distinguish the main polyacetylenes occurring in the same plant material using FT-Raman spectroscopy [182]. Two similar polyacety-lenes, i.e. falcarinol and falcarindiol, usually co-exist in plants so that it could be difficult to detect them simultane-ously in situ. However, the shift of the characteristic band positions in the registered Raman spectra for both polyacety-lenes is 6 cm-1 and it is sufficient to differentiate between both compounds. Similarly to that, other polyacetylenes can be distinguished on the basis of the position of characteristic bands. Furthermore, the Raman mapping technique gives a good opportunity to study the distribution of polyacetylenic compounds through different tissues within the plant.

Falcarinol and falcarindiol have been experimentally de-tected, for instance, in cultivated carrot roots (Daucus carota

L. ssp. sativus). The spectrum taken from the roots of the high-carotene orange roots showed a few overlapping bands in the region of 2260 and 2250 cm-1. However, the maxi-mum of the Raman signal registered in phloem tissue close to the vascular cambium was observed at 2257 cm-1. It seems that falcarinol was the main polyacetylene present in that area [137]. Other studies have shown the dependence of the location of carrot root measurement on the spectral shape. Thus, a higher number of polyacetylenic compounds has been expected in the studied plant materials.

The distribution of polyacetylenes has been investigated using the Raman mapping technique. It has been reported that the accumulation of polyacetylenes was located in the outer section of the root, i.e. in secondary phloem tissue close to the vascular cambium as well as in pericyclic paren-chyma tissue close to the periderm. On the other side, xylem parenchyma contained only small amounts of polyacetylenes [137]. Heterogeneous distribution of polyacetylenes over a root cross-section is in good agreement with reports based on chromatographic measurements [155]. Thus, heterogene-ous and tissue-specific occurrence of these compounds in live material has been visualized and proved by Raman mapping. The localization of the polyacetylenes in carrot roots can be explained by the presence of vascular bundles in a young secondary phloem and pericycle oil channels in the vicinity of the periderm. These channels could be responsi-ble for the transport and accumulation of polyacetylenes [185]. It should be mentioned that the accumulation of poly-acetylenes relates to the high concentration of carotenes, but the maximal concentrations of them were identified not at the same areas. Moreover, carrots containing high amounts of carotenoids possess also high levels of polyacetylenes. Measurements performed along the diameter of a trans-versely cut carrot root showed the distribution of both polya-cetylenes and carotenoids as well as their relative concentra-tions in the investigated plant material. The presented plot for the polyacetylene distribution was not symmetrical. It seems that measurements performed at the vascular bundles,

Recent Advances in Raman Analysis of Plants Current Analytical Chemistry, 2013, Vol. 9, No. 1 121

occasionally distributed in the phloem tissue, show higher concentration of polyacetylenes (see Fig. 4 in ref. [137]).

Wild carrot species (D. carota ssp. maritimus, D. carota

ssp. gummifer, D. carota ssp. commutatus) have been also investigated with regard to the accumulation of polyacety-lenes. Raman spectra of the wild carrot species roots have shown a symmetric band at 2253 cm-1 indicating that fal-carindiol is the main polyacetylene in that carrot. Further-more, the position and the structure of the band have not been dependent on the location of the measurement. Recent work has shown differences in the Raman spectra of two wild carrot species (D. carota ssp. gummifer, D. carota ssp. maximus) [186]. The spectra exhibited characteristic band usually around 2252 cm-1, however, the positions of band maxima obtained for individual carrots differed slightly. For one species the observed band maximum was shifted to 2248 cm-1, but registered spectra often showed slight asymmetry and sometimes even distinctive broadening. It may suggest the presence of other falcarindiol conformers or a specific interaction with the plant matrix [186]. The lack of Raman band at 2258 cm-1 in the spectra of wild carrot roots, which is characteristic for falcarinol, implies that falcarinol was either absent in roots or present at too low concentration so that its band could overlapp with the more intense band of falcarindiol. Indeed, lower concentration of falcarinol in comparison with falcarindiol has been already confirmed by chromatographic measurements for wild carrot species [187]. Raman mapping has shown the difference between the distribution of polyacetylenes in wild and cultivated or-ange carrots. It has been reported that the whole phloem tis-sue was rich in polyacetylenes, but the maxima could have been also observed near the pericyclic parenchyma. Analo-gous distribution of polyacetylenes has been shown in roots of other wild carrot species. However, in recent studies, the distribution of polyacetylenes has not been shown to be uni-form within the phloem tissue and areas with high concen-tration of these compounds have been presented to occur occasionally [186]. Additionally, that study did not pointed out higher concentration of polyacetylenes close to the periderm. The significant amounts of falcarindiol found in wild carrot species can be related to their resistance to some diseases, for instance, root knot nematodes [188].

Main polyacetylenes have been also found in other plants from the Apiaceae family. Three widespread vegetables, i.e. parsnip (Pastinaca sativa L.), celeriac (Apium graveolens var. rapaceum L.), and parsley (Petroselinum crispum Mill. Nyman ex A.W. Hill), were investigated to identify and show the distribution of polyacetylenic compounds [189]. Several polyacetylenes were identified in parsnip using chromatographic methods such as falcarinol and falcarindiol (occurring in the highest concentrations) as well as falcari-none and falcarinolone, the ketone derivatives of falcarinol and falcarindiol, and polyacetylenic C18 ketoaldehyde (Scheme 5) [190]. The Raman spectra obtained from various points of the parsnip roots showed distinct signals in the re-gion of 2180 – 2270 cm-1 [189]. The spectral pattern and positions of bands maxima depended on the localization within the root. Since only experimental spectra of falcarinol and falcarindiol can be found in literature, the unequivocal assignment of other detected bands to the individual com-

pounds are not possible. The distribution of polyacetylenic compounds in parsnip roots was studied by Raman mapping technique. It was found that polyacetylenes were not uni-formly distributed over the roots and they accumulated mainly in the phloem tissue [189]. Furthermore, the accumu-lation of these compounds was not uniform even within the same tissue.

Falcarinol and falcarindiol were also identified in cele-riac roots in relatively high concentrations applying HPLC methods. They were accompanied by 8-O-methylfalcarindiol and panaxydiol. Additionally, two other polyacetylenes, fal-carinone and falcarinolone, have been proven to occur in celeriac roots, but there is no information on their concentra-tion in this plant. Raman spectra from various points of the root showed the same pattern and a band maximum at about 2252 cm-1 [189]. Thus, the results indicated that falcarindiol dominates in celeriac root and other polyacetylenes were not detectable in the investigated material. The Raman mapping technique was also applied to study the polyacetylene distri-bution within the whole transversely cut root. It was shown that polyacetylenes were accumulated only in a few small areas occasionally localized close to the peel [189].

Raman spectroscopy has been successfully applied for investigations of polyacetylenic compounds in parsley roots. Four polyacetylenes have been already found in this plant, namely, falcarindiol (present at the highest concentration), falcarinol, 8-O-methylfalcarindiol, and panaxydiol. Similarly to parsnip, Raman spectra taken from parsley roots showed the diversity of patterns and positions of the maxima. Apart from characteristic bands for falcarinol and falcarindiol, other bands that can be assigned to other polyacetylenes or to some effects connected with plant matrix was detected [189]. A Raman map and the related image of a parsley root showed a quite uniform distribution of polyacetylenes in the phloem tissue close to the secondary cambium. However, areas with higher concentration of these compounds were present occasionally.

Raman spectroscopy can be successfully applied for studies of other plants as it was shown for ginseng roots, where two polyacetylenes, i.e. falcarinol and panaxydol, were detected [143]. Although Raman spectra of the isolated compounds exhibited characteristic bands at about 2260 cm-1, a band at 2237 cm-1 was observed in the spectrum taken from fresh ginseng root that is about 20 cm-1 lower than for the isolated polyacetylenes. Furthermore, the Raman spec-trum of freeze-dried roots exhibited a band with a maximum at 2258 cm-1, while the Raman spectrum obtained after hy-dration of the freeze-dried roots showed again the band at 2237 cm-1. The most possible explanation of this phenome-non is an interaction between polyacetylene molecules and plant components (e.g. metal ions) in presence of water re-sulting in the formation of a stable metal complex. Observed phenomenon can be easily followed using Raman mapping technique. Raman maps, colored according to the individual band intensities, have reflected the changes after drying and then after hydration of the freeze-dried roots. Thus, Raman spectroscopy seems to be a powerful tool to investigate structural changes in situ without any preliminary prepara-tion of a sample. The distribution of polyacetylenes in gin-seng roots is similar to that for carrot roots [143].

122 Current Analytical Chemistry, 2013, Vol. 9, No. 1 Baranska et al.

Falcarindiol was also identified in the Raman spectrum of Angelica dahurica root, where this polyacetylene is re-sponsible for the antibiotic activity of this Chinese drug, on the basis of the position of the characteristic band at 2252 cm-1 [149].

However, other polyacetylenic compounds can be also found in plants. For instance, in the Raman spectrum of the flower of Erigeron neglectus a strong band at 2198 cm-1 was observed. It has been attributed to the lachnophyllum lactone that is a monoacetylenic derivative of 2-furanone. The Ra-man spectrum of lachnophyllum lactone derivative with a double bond in the side chain (Scheme 5) showed a very strong band about 20 cm-1 lower compared to the real com-pound at 2178 cm-1 which may be caused by the difference in the molecular structure [180].

In the roots of another plant, Coreopsis grandiflora, a band at 2194 cm-1 that comes from a monoacetylene with a substitution by a thiophene ring was observed. It was found that a compound with one thiophene ring substituted by two monoacetylenes (Scheme 5, thiophenic diacetylene) has a similar spectrum with a band maximum at 2192 cm-1 [180].

Measurements of the flowers of the ox-eye daisy (Chry-santhemum leucanthemum) produced Raman spectra includ-ing a doublet with maxima at 2172 and 2221 cm-1, while the spectrum of the isolated trans-dehydromatricariaester (Scheme 5) had the same pattern with band maxima at 2173 and 2222 cm-1 [180].

The blossoms of other plant species from the Asteraceae family, i.e. cornflower (Centaurea cyanus), exhibited a dou-blet with maxima at 2172 and 2199 cm-1 in their Raman spectra that is in good accordance with the spectrum of (6E)-tetradec-6-ene-8,10,12-triyn-3-one (Scheme 5). In the Ra-man spectrum of this compound the maxima of the doublet were present at 2179 and 2220 cm-1, however, the ratio of the band intensities was reverse suggesting that another polyacetylene or another isomer was also present in the blos-soms [180].

Other examples of polyacetylenes in plants are the flower heads of Centaurea ruthenica and the leaves of Carthamus lanatus. They showed polyacetylene bands of which the po-sition (2167 cm-1) was nearly identical with that of triacety-lenic diacetate presented in Scheme 5 (2166 cm-1) [180].

Characteristic polyacetylenic bands were also observed at 2196 cm-1 in the spectrum of the flower of apache beggar-ticks (Bidens ferulifolia) that is exclusively used as horticul-tural plant mainly in Guatemala and Mexico. Other species of the genus Bidens are widely used in Chinese medicine, such as Bidens pilosa and Bidens campylotheca. Several lipophilic polyacetylenes and polyacetylene glucosides were identified in these species that are likely to be responsible for the described medicinal properties [191,192]. A phenyl sub-stituted triine (hepta-1,3,5-triyn-1-ylbenzene, Scheme 5) was found in Bidens pilosa species [193] and its Raman spectrum was similar to that obtained from the flower of apache beg-garticks (2191 cm-1 comparing to 2196 cm-1 for the plant) [180]. The Raman mapping technique confirmed also the distribution of polyacetylenes reported earlier for chamomile (Matricaria chamomilla L.), where cis and trans-spiroethers were found in the flower. It has been discovered that polya-

cetylenes occur mainly in the stamen, whereas carotenes occur obviously in the petals.

CONCLUSION

Raman spectroscopy has several advantages of many other analytical methods. As shown in this review, Raman measurements provide detailed and comprehensive informa-tion on plant chemical components that can be successfully assessed in both fresh and processed material. Raman map-ping enables visualization of the distribution of individual compounds even in live and intact plant tissues. Moreover, determination of metabolites at cellular level is feasible due to recent progress in the instrumental development. These unique features make Raman technique attractive also for quality control of processed plant products as well as raw material used in pharmaceutical or food industry [194,195]. Raman spectroscopy is considered as a non-destructive ap-proach in contrast to chromatographic methods that are de-structive to biological material and require sample prepara-tion techniques such as extraction or distillation of the inves-tigated compound. This advantage can be of particular sig-nificance as live or at least intact biological material can be analyzed using Raman spectroscopy before it is subjected to other destructive techniques.

CONFLICT OF INTEREST

The author(s) confirm that this article has no conflicts of interest.

ACKNOWLEDGEMENTS

This research was supported by the Ministry of Science and Higher Education (grant no. N204013635, 2008-2011). JCzD is grateful for partial support by Grant of Ministry of Higher Education for statutory activity of National Medi-cines Institute. The research was supported by the Interna-tional PhD-studies program at the Faculty of Chemistry Jagiellonian University within the Foundation for Polish Science MPD Program co-financed by the EU European Regional Development Fund.

REFERENCES

[1] Schrader, B. In Infrared and Raman Spectroscopy. Methods and Applications, Schrader, B., Ed.; VCH: Weinheim, 1995.

[2] Creighton, J.A. In Spectroscopy of Surface, Clark, R.J.; Hester, R.E., Eds.; Wiley: New York, 1988; Vol. 21.

[3] Baranska, M.; Schulz, H.; Rösch, P.; Strehle, M.A.; Popp, J. Identi-fication of secondary metabolites in medicinal and spice plants by NIR-FT-Raman microspectroscopic mapping. Analyst, 2004, 129, 926-930.

[4] Laserna, J.J. In Modern techniques in Raman spectroscopy, Laserna, J.J., Ed.; John Wiley & Sons: New York, 1996.

[5] Schulz, H.; Baranska, M. Identification and quantification of valu-able plant substances by IR and Raman spectroscopy. Vib. Spect., 2007, 43, 13-25.

[6] Karrer, P.; Jucker, E. In Carotenoids, Elsevier: New York, 1950. [7] Maoka, T. Recent progress in structural studies of carotenoids in

animals and plants. Arch. Biochem. Biophys., 2009, 483, 191-195. [8] Baranska, M.; Schulz, H. In The Alkaloids Chemistry and Biology,

Cordell, G.A., Ed.; Elsevier: The Netherlands, 2009; Vol. 67; pp 217-255.

[9] E. Fattorusso, O. Taglialatela-Scafati (Eds.) Modern Alkaloids, Wiley: Weinheim 2008.

Recent Advances in Raman Analysis of Plants Current Analytical Chemistry, 2013, Vol. 9, No. 1 123

[10] Edwards, H.G.M.; Farwell, D.W.; de Oliveira, L.F.C.; Alia, J.M.; Le Hyaric, M.; de Ameida, M.V. FT-Raman spectroscopic studies of guarana and some extracts. Anal. Chim. Acta, 2005, 532, 177-186.

[11] Edwards, H.G.M.; Villar, S.E.J.; de Oliveira, L.F.C.; Le Hyaric, M. Analytical Raman spectroscopic study of cacao seeds and their chemical extracts. Anal. Chim. Acta., 2005, 538, 175-180.

[12] Armenta, S.; Garrigues, S.; Guardia, M. Solid-phase FT-Raman determination of caffeine in energy drinks. Anal. Chim. Acta, 2005, 547, 197-203.

[13] Baranska, M.; Proniewicz, L.M.; Raman analysis of caffeine alka-loid. Vib. Spectrosc., 2008, 48, 153-157.

[14] Pavel, I.; Szeghalmi, A.; Moigno, D.; Cinta, S.; Kiefer, W. Theo-retical and pH Dependent Surface Enhanced Raman Spectroscopy Study on Caffeine. Biopolymers, 2003, 72, 25-37.

[15] Cinta Pinzaru, S.; Pavel, I.; Leopold, N.; Kiefer, W. Identification and characterization of pharmaceuticals using Raman and surface-enhanced Raman scattering. J. Raman Spectrosc., 2004, 35, 338-346.

[16] Schulz, H.; Baranska, M.; Quilitzsch, R.; Schütze, W. Determina-tion of alkaloids in capsules, milk and ethanolic extracts of poppy (Papaver somniferum L.) by ATR-FT-IR and FT-Raman spectros-copy. Analyst, 2004, 129, 917-920.

[17] Li, P.-W.; Zhang, J.; Zhang, L.; Mo, Y.-J. Surface-enhanced Ra-man scattering and adsorption studies of morphine on silver island film. Vib. Spectrosc., 2009, 49, 2-6.

[18] Baranska, M.; Kaczor, A. Morphine studied by vibrational spec-troscopy and DFT calculations. J. Raman Spectrosc., 2012, 43, 102-107.

[19] Hofenk de Graaf, J. H. In The Colourful Past, Archetype Publica-tions Ltd.: London, 2004; p 171-213.

[20] Burgo, L.; Clark, R.J.H. Library of FT-Raman spectra of pigments, minerals, pigment media and varnishes, and supplement to existing library of Raman spectra of pigments with visible excitation. Spec-

trochim. Acta A, 2001, 57, 1491-1521. [21] Leona, M.; Lombardi, J.R. Identification of berberine in ancient

and historical textiles by surface-enhanced Raman scattering. J. Raman Spectrosc., 2007, 38, 853-858.

[22] Wang, S.L.; Yu, J.S. Chem. Res. Chin. Univ., 2002, 23, 1676-. (Wang, S.L.; Yu, J.S. SERS Studies on the Interaction of Berberine with ct DNA. Chem. J. Chin. Univ. -Chin., 2002, 23, 1676.)

[23] Raynes, K. Bisquinoline antimalarials: Their role in malaria che-motherapy. Int. J. Parasitol., 1999, 29, 367-379.

[24] Frosch, T.; Schmitt, M.; Popp, J. In situ UV resonance Raman micro-spectroscopic localization of the antimalarial quinine in cin-chona bark. J. Phys. Chem. B, 2007, 111, 4171-4177.

[25] Bringmann, G.; Rübenacker, M.; Vogt, P.; Busse, H.; Assi, L.A. Dioncophylline C from the roots of Triphyophyllum peltatum, the first 5,1 -coupled dioncophyllaceae alkaloid. Phytochemistry, 1992, 31, 4019-4024.

[26] Frosch, T.; Schmitt, M.; Schenzel, K.; Faber, J.H.; Bringmann, G.; Kiefer, W.; Popp, J. In vivo localization and identification of the antiplasmodial alkaloid dioncophylline a in the tropical liana Triphyophyllum peltatum by a combination of fluorescence, near infrared fourier transform Raman microscopy, and density func-tional theory calculations. Biopolymers, 2006, 82, 295-300.

[27] Frosch, T.; Schmitt, M.; Noll, T.; Bringmann, G.; Schenzel, K.; Popp, J. Ultrasensitive in situ tracing of the alkaloid dioncophylline A in the tropical liana Triphyophyllum peltatum by applying deep-UV resonance raman microscopy. Anal. Chem., 2007, 79, 986-993.

[28] Ryder, A.G.; O’Connor, G.M.; Glynn, T.J. Quantitative analysis of cocaine in solid mixtures using Raman spectroscopy and chemometric methods. J. Raman Spectrosc., 2000, 31, 221-227.

[29] Perez, R.; Ruperez, A.; Laserna, J.J. Evaluation of silver substrates for surface-enhanced Raman detection of drugs banned in sport practices. Anal. Chim. Acta, 1998, 376, 225-263.

[30] Tan, C.; Wang, J.; Wu, T.; Qin, X.; Li, M. Determination of nico-tine in tobacco samples by near-infrared spectroscopy and boosting partial least squares. Vib. Spectrosc., 2010, 54, 35-41.

[31] Dezelic, M. The viscosities of liquid mixtures with pyrrole as a component. Trans. Faraday Soc., 1937, 33, 713-719.

[32] Walrafen, G.E. Low-Frequency Raman Spectra from Nicotine. J. Chem. Phys., 1971, 55, 4677-4678.

[33] McDevitt, N.T.; Fateley, W.G. The character of low frequency (250-40 cm-1) raman bands from organic liquids and their compari-son to far infrared absorption. J. Mol. Struct., 1970, 5, 477-490.

[34] Nicholas, M.L.; Powell, D.L.; Williams, T.R.; Thompson, R.Q.; Oliver, N.H. Reference raman spectra of eleven miscellaneous pes-ticides. J. Assoc. Off. Anal. Chem., 1976, 59, 1266-1275.

[35] Aslanian, D. Vibrational spectroscopic approach to the study of acetylcholine and related compounds. Life Sci., 1983, 32, 2809-2825.

[36] Ishikawa, H.; Imai, Y.; Kurokawa, Y. Preparation of Ag particle-doped cellulose acetate gel membrane as a surface-enhanced Ra-man scattering active substrate. Vib. Spectrosc., 1995, 8, 445-449.

[37] Barber, T.E.; List, M.S.; Haas, J.W.; Wachter, E.A. Determination of nicotine by surface-enhanced Raman scattering (SERS). Appl. Spectrosc., 1994, 48, 1423-1427.

[38] Bell, S.E.J.; Sirimuthu, N.M.S. Rapid, quantitative analysis of ppm/ppb nicotine using surface-enhanced Raman scattering from polymer-encapsulated Ag nanoparticles (gel-colls). Analyst, 2004, 129, 1032-1036.

[39] Baranska, M.; Dobrowolski, J.Cz.; Kaczor, A.; Chruszcz-Lipska, K.; Gorz, K.; Rygula, A. Tobacco alkaloids analyzed by Raman spectroscopy and DFT calculations. J. Raman Spectr., 2012, 43, 1065-1073.

[40] Crabb, T.A.; Newton, R.F.; Jackson, D. Stereochemical studies of nitrogen bridgehead compounds by spectral means. Chem. Rev., 1971, 71, 109-126.

[41] Pal, T.; Narayanan, V.A.; Stokes, D.L.; Vo-Dinh, T. Surface-enhanced Raman detection of nicotinamide in vitamin tablets. Anal. Chim. Acta, 1998, 368, 21-28.

[42] Schulz, H.; Baranska, M.; Quilitzsch, R.; Schütze, W.; Lösing, G. Characterization of peppercorn, pepper oil, and pepper oleoresin by vibrational spectroscopy methods. J. Agric. Food Chem., 2005, 53, 3358-3363.

[43] Nomenclature of Carotenoids (Rules Approved 1974) International Union of Pure and Applied Chemistry and International Union of Biochemistry, Issued by the IUPAC Commission on the Nomen-clature of Organic Chemistry and the IUPAC—IUB Commission on Biochemical Nomenclature, Butter Worths; London, 1974.

[44] Hirschberg, J.; Cohen, M.; Harker, M.; Lotan, T.; Mann, V.; Pecker, I. Molecular genetics of the carotenoid biosynthesis path-way in plants and algae. Pure Appl. Chem., 1997, 69, 2151-2158.

[45] Hirschberg, J. Carotenoid biosynthesis in flowering plants. Curr. Opin. Plant Biol., 2001, 4, 210-218.

[46] Sandmann, G. Carotenoid Biosynthesis and Biotechnological Ap-plication. Arch. Biochem. Biophys., 2001, 385, 4-12.

[47] Ronen, G.; Carmel-Goren, L.; Zamir, D.; Hirschberg, J. An alter-native pathway to -carotene formation in plant chromoplasts dis-covered by map-based cloning of Beta and oldgold color muta-tions in tomato. Proc. Natl. Acad. Sci., 2000, 97, 11102-11107.

[48] Gallagher, C.E.; Matthews, P.D.; Li, F.; Wurtzel, E.T. Gene Du-plication in the Carotenoid Biosynthetic Pathway Preceded Evolu-tion of the Grasses. Plant Physiology., 2004, 135, 1776-1783.

[49] Park, H.; Kreunen, S.S.; Cuttriss, A.J.; DellaPenna, D.; Pogson, B.J. Identification of the carotenoid isomerase provides insight into carotenoid biosynthesis, prolamellar body formation, and photo-morphogenesis. Plant Cell., 2002, 14, 321-332.

[50] Green, B.R.; Parson, W.W. In Light-harvesting Antennas in Pho-tosynthesis, Green, B.R.; Parson, W.W., Eds.; Kluwer Academic Publisher: Dordrecht, 2003.

[51] Bode, S.; Quentmeier, C.C.; Liao, P.-N.; Hafi, N. ; Barros, T.; Wilk, L.; Bittner, F.; Walla, P.J. On the regulation of photosynthe-sis by excitonic interactions between carotenoids and chloro-phylls. PNAS, 2009, 106, 12311-12316.

[52] Polívka, T.; Sundström, V. Ultrafast dynamics of carotenoid ex-cited states - from solution to natural and artificial systems. Chem Rev., 2004, 104, 2021-2071.

[53] Wormit, M.; Dreuw, A. Quantum chemical insights in energy dissipation and carotenoid radical cation formation in light harvest-ing complexes. Phys. Chem. Chem. Phys., 2007, 9, 2917-2931.

[54] Wormit, M.; Harbach, P.H.P.; Mewes, J.M.; Amarie, S.; Wachtveitl, J.; Dreuw, A. Excitation energy transfer and carotenoid radical cation formation in light harvesting complexes — A theo-retical perspective. Bioch. Biophys. Acta, 2009, 1787, 738-746.

124 Current Analytical Chemistry, 2013, Vol. 9, No. 1 Baranska et al.

[55] Mlodzinska, E. Survey of Plant Pigments: Molecular and Envi-ronmental Determinants of Plant Colors. Acta Biol. Cracoviensia Ser. Bot., 2009, 51, 7-16.

[56] Hamilton, W.D.; Brown, S.P. Autumn tree colours as a handicap signal. Proc. Royal Soc. London. Biol. Lett., 2001, 268, 1489-1493.

[57] Lee, D.W.; Gould, K.S. Why leaves turn red. Amer. Sci., 2002, 90, 524-531.

[58] Furukawa, Y.; Takeuchi, H.; Harada, I.; Tasumi, M. Molecular Force Fields of s-trans-1,3-Butadiene and the Second Stable Con-former. Bull. Chem. Soc. Jpn., 1983, 56, 392-399.

[59] Panchenko, Yu.N.; De Maré, G.R. Vibrational Analysis of Buta-1,3-Diene and its Deutero and 13C Derivatives and some of Their Rotational Isomers. J. Struct. Chem., 2008, 49, 235-244.

[60] Furukawa, Y. In The Chemistry of Dienes and Polyenes, Rap-poport, Z., Ed.; Wiley & Sons: New York, 1997; Vol. 1.

[61] Rimai, L.; Heyde, M.E.; Gill, D. Vibrational spectra of some caro-tenoids and related linear polyenes. Raman spectroscopic study. J. Am. Chem. Soc., 1973, 95, 4493-4501.

[62] Ci, X.; Pereira, M.A.; Myers, A.B. Resonance Raman spectra of trans 1,3,5 hexatriene in solution: Evidence for solvent effects on excited state torsional motion. J. Chem. Phys., 1990, 92, 4708-4717.

[63] Schaffer, H.E.; Chance, R.R.; Silbey, R.J.; Knoll, K.; Schrock, R.R. Conjugation Length Dependence of Raman Scattering in a Se-ries of Linear Polyenes: Implications for Polyacetylene. J. Chem. Phys., 1991, 94, 4161-4170.

[64] Hashimoto, H.; Koyama, Y.; Hirata, Y.; Mataga, N. S1 and T1 species of -carotene generated by direct photoexcitation from the all-trans, 9-cis, 13-cis, and 15-cis isomers as revealed by picosec-ond transient absorption and transient Raman spectroscopies. J. Phys. Chem., 1991, 95, 3072-3076.

[65] Naruse, M.; Hashimoto, H.; Kuki, M.; Koyama, Y. Triplet Excita-tion of Precursors of Spirilloxanthin Bound to the Chromatophores of Rhodospirillum Rubrum as Detected by Transient Raman Spec-troscopy. J. Mol. Struct., 1991, 242, 15-26.

[66] Merlin, J.C. Resonance Raman spectroscopy of carotenoids and carotenoid containing systems. Pure Appl. Chem., 1985, 57, 785-792.

[67] Kupka, T.; Ming Lin, H.; Stobinski, L.; Chen, C.-H.; Liou, W.-J.; Wrzalike, R.; Flisaka, Z. Experimental and theoretical studies on corals. I. Toward understanding the origin of color in precious red corals from Raman and IR spectroscopies and DFT calculations. J. Raman Spectrosc., 2010, 41, 651-658.

[68] Saito, S.; Tasumi, M. Normal-coordinate analysis of -carotene isomers and assignments of the Raman and infrared bands. J. Ra-

man Spectrosc., 1983, 14, 310-321. [69] Weesie, R.J.; Merlin, J.C.; Lugtenburg, J.; Britton, G.; Jansen, F.J.;

Cornard, J.P. Semiempirical and Raman spectroscopic studies of carotenoids. Biospectroscopy., 1999, 5, 19-33.

[70] Schlücker, S.; Szeghalmi, A.; Schmitt, M.; Popp, J.; Kiefer, W. Density functional and vibrational spectroscopic analysis of -carotene. J. Raman Spectrosc., 2003, 34, 413-419.

[71] Berezin, K.V. Nechaev, V.V. Calculation of the IR spectrum and the molecular structure of -carotene. J. Appl. Spectr., 2005, 72, 164-171 (Zh. Prikl. Spektrosk., 2005, 72, 157-164.).

[72] Tschirner, N.; Schenderlein, M.; Brose, K.; Schlodder, E.; Mrogin-ski, M.A.; Hildebrandt, P.; Thomsen, C. Raman excitation profiles of -carotene - novel insights into the nature of the 1-band. Phys. Stat. Sol. B, 2008, 245, 2225-2228.

[73] Hoskins, L.C. Resonance Raman excitation profiles of lycopene. J. Chem. Phys., 1981, 74, 882-885.

[74] Cotting, J.E.; Hoskins, L.C.; Levana, M.E. Effects of inhomogene-ous broadening on the resonance Raman excitation profile of lyco-pene. J. Chem. Phys., 1982, 77, 1081-1086.

[75] Hoskins, L.C. Resonance Raman Spectroscopy of Beta-Carotene and Lycopene: A Physical Chemistry Experiment. J. Chem. Educ., 1984, 61, 460-462.

[76] López-Ramírez, M.R.; Sanchez-Cortes, S.; Pérez-Méndezc, M.; Blanch, G. Trans-cis isomerisation of the carotenoid lycopene upon complexation with cholesteric polyester carriers investigated by Raman spectroscopy and density functional theory. J. Raman Spec-

trosc., 2010, 41, 1170-1177. [77] Salares, V.R.; Mendelsohn, R.; Carey, P.R.; Bernstein, H.J. Corre-

lation between the absorption spectra and resonance Raman excita-tion profiles of astaxanthin. J. Phys. Chem., 1976, 80, 1137-1141.

[78] Leigh, S.; Leigh, M.L.S.; Van Hoogevest, P. Crystal forms of astaxanthin. US Patent, 2009, US7563935 B2.

[79] Guo, J.; Jones, M.J.; Ulrich, J. Polymorphism of 3,3'-dihydroxy-

, -carotene-4,4'-dione (Astaxanthin). Chem. Eng. Res. Design, 2010, 88, 1648-1652.

[80] Merlin, J.C. Resonance Raman spectroscopy of carotenoids and carotenoid containing systems. Pure Appl. Chem., 1985, 57, 785-792.

[81] Feltl, L.; Pacáková, V.; tulík, K.; Volka, K. Reliability of Caro-tenoid Analyses: A Review. Curr. Anal. Chem., 2005, 1, 93-102.

[82] Gierlinger, N.; Schwanninger, M. The potential of Raman micros-copy and Raman imaging in plant research. Spectroscopy, 2007, 21, 69-89.

[83] Schulz, H.; Baranska, M.; Baranski, R. Potential of NIR-FT-Raman Spectroscopy in Natural Carotenoid Analysis. Biopolymers, 2005, 77, 212-221.

[84] Baranska, M.; Schulz, H.; Rösch, P.; Strehle, M.A.; Popp, J. Identi-fication of secondary metabolites in medicinal and spice plants by NIR-FT-Raman microspectroscopic mapping. Analyst, 2004, 129, 926-930.

[85] Baranski, R.; Baranska, M.; Schulz, H. Changes in carotenoid content and distribution in living plant tissue can be observed and mapped in situ using NIR-FT-Raman spectroscopy. Planta, 2005, 222, 448-457.

[86] Pudney, P.D.A.; Gambelli, L.; Gidley, M. Confocal Raman mi-crospectroscopic study of the molecular status of carotenoids in tomato fruits and foods. Appl. Spectr., 2011, 65, 127-134.

[87] Baranska, M.; Schulz, H.; Baranski, R.; Nothnagel, T.; Christen-sen, L.P. In situ simultaneous analysis of polyacetylenes, caro-tenoids and polysaccharides in carrot roots. J. Agric. Food. Chem., 2005, 53, 6565-6571.

[88] Baranska, M.; Baranski, R.; Schulz, H.; Nothnagel, T. Tissue-specific accumulation of carotenoids in carrot roots. Planta, 2006, 224, 1028-1037.

[89] Gamsjaeger, S.; Baranska, M.; Schulz, H.; Heiselmayer, P.; Musso, M. Discrimination of carotenoid and flavonoid content in petals of pansy cultivars (Viola x wittrockiana) by FT-Raman spectroscopy. J. Raman Spectrosc., 2011, 42, 1240-1247.

[90] Baranska, M.; Baranski, R.; Grzebelus, E.; Roman, M. In situ detection of a single carotenoid crystal in a plant cell using Raman microspectroscopy. Vib. Spectr., 2011, 56, 166-169.

[91] Brackmann, C.; Bengtsson, A.; Alminger, M.L.; Svanberg, U.; Enejder, A. Visualization of -carotene and starch granules in plant cells using CARS and SHG microscopy. J. Raman Spectr., 2011, 42, 586-592.

[92] da Silva, C.E.; Vandenabeele, P.; Edwards, H.G.M.; Cappa de Oliveira, L.F. NIR-FT-Raman spectroscopic analytical characteri-zation of the fruits, seeds, and phytotherapeutic oils from rosehips. Anal. Bioanal. Chem., 2008, 392, 1489-1496.

[93] de Oliveira, V.E.; Castro, H.V.; Edwards, H.G.M.; de Oliveira, L.F.C. Carotenes and carotenoids in natural biological samples: a Raman spectroscopic analysis. J. Raman Spectrosc., 2010, 41, 642-650.

[94] López-Sánchez, M.; Ayora-Cañada, M.J.; Molina-Díaz, A. Olive Fruit Growth and Ripening as Seen by Vibrational Spectroscopy. J. Agric. Food Chem., 2010, 58, 82-87.

[95] Ivleva, N.P.; Niessner, R.; Panne, U. Characterization and dis-crimination of pollen by Raman microscopy. Anal. Bioanal. Chem., 2005, 381, 261-267.

[96] Schulte, F.; Lingott, J.; Panne, U.; Kneipp, J. Chemical Characteri-zation and Classification of Pollen. Anal. Chem., 2008, 80, 9551-9556.

[97] Schulte, F.; Mäder, J.; Kroh, L.W.; Panne, U.; Kneipp, J. Charac-terization of Pollen Carotenoids with in situ and High-Performance Thin-Layer Chromatography Supported Resonant Raman Spec-troscopy. Anal. Chem., 2009, 81, 8426-8433.

[98] Joseph, V.; Schulte, F.; Rooch, H.; Feldmann, I.; Dörfel, I.; Österle, W.; Panne, U.; Kneipp, J. Surface-enhanced Raman scat-tering with silver nanostructures generated in situ in a sporopollen in biopolymer matrix. Chem. Commun., 2011, 47, 3236-3238.

[99] Sánchez-Millán, J.L; Serrato-Cruz, M.A.; Del Villar-Martínez, A.A.; Venegas-Espinosa, P.E.; Evangelista-Lozano, S.; Jiménez-Aparicio, A.; Quintero-Gutiérrez, A. G.; García-Jiménez, F.A. Es-pectroscopia Raman: Herramienta Rápida Y No Invasiva Para De-terminar Carotenoides En Inflorescencias De Tagetes Erecta L. Raman Spectroscopy: A Rapid, Non-Invasive Tool For Determin-

Recent Advances in Raman Analysis of Plants Current Analytical Chemistry, 2013, Vol. 9, No. 1 125

ing Carotenoids In Inflorescences of Tagetes erecta L. Agrociencia, 2007, 41, 863-871.

[100] Maoka, T. Sterically hindered carotenoids with 3Z, 5Z configura-tion from the seeds of oriental bitter sweet, Celastrus orbiculatus. Phytochem., 2009, 70, 920-923.

[101] Kaczor, A.J.; Turnau, K.; Baranska, M. In situ Raman imaging of astaxanthin in a single microalgal cell. Analyst., 2011, 136, 1109-1112.

[102] Kaczor, A.J.; Baranska, M. Structural changes of carotenoid astax-anthin in a single algal cell monitored in situ by Raman spectros-copy. Anal. Chem., 2011, 83, 7763-7770.

[103] Abbas, A.; Josefson, M.; Abrahamsson, K. Characterization and mapping of carotenoids in the algae Dunaliella and Phaeodactylum using Raman and target orthogonal partial least squares. Chemom. Intell. Lab. Syst., 2011, 107, 174-177.

[104] Collins, A.M.; Jones, H.D.T.; Han, D.; Hu, Q.; Beechem, T.E.; Timlin, J.A. Carotenoid distribution in living cells of Haematococ-cus pluvialis (Chlorophyceae). PLoS ONE, 2011, 6, e24302.

[105] Sicard, C.; Brayner, R.; Margueritat, J.; Hémadi, M.; Couté, A.; Yéprémian, C.; Djediat, C.; Aubard, J.; Fiévet, F.; Livage, J.; Co-radin, T. Nano-goldbiosynthesis by silica-encapsulated micro-algae: a “living” bio-hybrid material. J. Mater. Chem., 2010, 20, 9342-9347.

[106] Premvardhan, L.; Roberta, B.; Beerb, A.; Büchel, C. Pigment orga-nization in fucoxanthin chlorophyll a/c2 proteins (FCP) based on resonance Raman spectroscopy and sequence analysis. Bioch. Bioph. Acta, 2010, 1797, 1647-1656.

[107] Heraud, P.; Wood, B.R.; Beardall, J.; McNaughton, D. Effects of pre-processing of Raman spectra on in vivo classification of nutri-ent status of microalgal cells. J. Chemometrics, 2006, 20, 193-197.

[108] Huang, Y.Y.; Beal, C.M.; Cai, W.W.; Ruoff, R.S.; Terentjev, E.M. Micro-Raman Spectroscopy of Algae: Composition Analysis and Fluorescence Background Behavior. Biotech. Bioeng., 2010, 105, 889-898.

[109] Heraud, P.; Beardall, J.; McNaughton, D.; Wood, B.R. In vivo prediction of the nutrient status of individual microalgal cells using Raman microspectroscopy. FEMS Microbiol Lett., 2007, 275, 24-30.

[110] Papaioannou, E.H.; Liakopoulou-Kyriakides, M.; Christofilos, D.; Arvanitidis, I.; Kourouklis, G. Raman spectroscopy for intracellu-lar monitoring of carotenoid in Blakeslea trispora. Appl. Biochem. Biotechn., 2009, 159, 478-487.

[111] Jorge-Villar, S.E.; Edwards, H.G.M. Lichen colonization of an active volcanic environment: a Raman spectroscopic study of ex-tremophile biomolecular protective strategies. J. Raman Spectrosc., 2010, 41, 63-67.

[112] Xu, X.; Yuan, Y.; Wang, X.; Li, Y. Monitoring and rapid quantifi-cation of total carotenoids in Rhodotorula glutinis cells using laser tweezers Raman spectroscopy. FEMS Microbiol. Lett., 2011, 314, 42-48.

[113] Bohlmann, F.; Bronowski, H.; Arndt, Ch. Natürlich vorkommende Acetylenverbindungen. Fortschr. Chem. Forsch., 1962, 4, 138-272.

[114] Bohlmann, F. Natürlich vorkommende Acetylenverbindungen. Fortschr. Chem. Forsch., 1965, 6, 65-100.

[115] Hansen, L.; Boll, P.M. Polyacetylenes in Araliaceae: Their chemis-try, biosynthesis and biological significance. Phytochemistry, 1986, 25, 285-293.

[116] Minto, R.E.; Blacklock, B.J. Biosynthesis and function of polyace-tylenes and allied natural products. Prog. Lipid Res., 2008, 47, 233-306.

[117] Arnaud, A.M. Sur un nouvel acide gras non saturé de la série CnH2n-4O2. Bull Soc Chim France., 1892, 7, 233-234.

[118] Arnaud, M.A. Sur la constitution de l’acide taririque. Bull. Soc. Chim. France, 1902, 27, 489-496.

[119] Glaser, C. Beiträge zur Kenntniss des Acetenylbenzols. Ber. Deutsch. Chem. Ges., 1869, 2, 422-424.

[120] Bayer, A. Über Polyacetylenverbindungen. Ber. Deutsch. Chem. Ges., 1885, 18, 2269-2281.

[121] Chodkiewicz, W. Synthesis of acetylenic compounds. Ann. Chim. (Paris), 1957, 2, 819-869.

[122] Cadiot, P.; Chodkiewicz, W. In Chemistry of Acetylenes, Viehe, H.G., Ed.; Marcel Dekker: New York, 1969; pp. 597-647.

[123] Marino, J.P.; Nguyen, H.N. Bulky Trialkylsilyl Acetylenes in the Cadiot-Chodkiewicz Cross-Coupling Reaction. J. Org. Chem., 2002, 67, 6841-6844.

[124] Gibtner, T.; Hampel, F.; Gisselbrecht, J.-P.; Hirsch, A. End-Cap Stabilized Oligoynes: Model Compounds for the Linear sp Carbon Allotrope Carbyne. Chem. Eur. J., 2002, 68, 408-432.

[125] Armitage, J.B.; Jones, E.R.H.; Whiting, M.C. Researches on acety-lenic compounds. Part XXVIII. A new route to diacetylene and its symmetrical derivatives. J. Chem. Soc., 1951, 44-47.

[126] Cataldo, F. Structural relationship between dicopper diacetylide (Cu-C C-C C-Cu) and dicopper acetylide (Cu-C C-Cu). Eur. J. Solid. State. Inorg. Chem., 1998, 35, 281-291.

[127] Cataldo, F. From dicopper diacetylide (Cu-C C-C C-Cu) to car-bine. Eur. J. Solid. State. Inorg. Chem., 1998, 35, 293-304.

[128] Sun, Y.; Taylor, N.J.; Carty, A.J. Models for Linear Acetylide Polymers: Synthesis and Single-Crystal X-ray Structures of the Ru-thenium(II) Acetylide and Dlacetylldes Ru(CO)2(PEt3)2 [(C=C)nR]2, n= 1, R =Me,Si; n = 2, R = Me,Si, H). Organometal-

lics, 1992, 11, 4293-4300. [129] Barbu, E.; Tsibouklis, J. Unsymmetrically substituted aliphatic

diacetylenes. Tetrah. Lett., 1996, 37, 5023-5026. [130] Volkov, A.N.; Volkova, K.A. -Haloacetylene and diacetylene

alcohols. Rus. J. Org. Chem., 2007, 43, 161-169. [131] Pattison, F.L.M.; Dear, R.E.A. Toxic Fluorine Compounds XIX.

O-Fluoroalkynes. Can. J. Chem., 1960, 41, 2600-2606. [132] Blau, W.J.; Byrne, H.J.; Cardinb, D.J.; Davey, A.P. Non-linear

optical properties of group 10 metal alkynyls and their polymers. J. Mater. Chem., 1991, 1, 245-249.

[133] Keller, T.M.; Homrighousen, C. High temperature ceramics de-rived from linear carborane-(siloxane or silane)-acetylene copoly-mers. U.S. Appl., 2000, No. 09/625,271, 30.

[134] Bohlmann, F.; Burkhardt, T.; Zdero, C. In Naturally Occurring Acetylenes, Bohlmann, F.; Burkhardt, T.; Zdero, C., Eds.; Aca-demic Press: London, 1973.

[135] Baranski, R.; Baranska, M.; Schulz, H.; Simon, P.W.; Nothnagel, T. Single seed Raman measurements allow taxonomical discrimi-nation of apiaceae Accessions collected in gene banks. Bio-polymers, 2006, 81, 497-505.

[136] Czepa, A.; Hofmann, T. Structural and sensory characterization of compounds contributing to the bitter off-taste of carrots (Daucus carota L.) and carrot puree. J. Agric. Food Chem., 2003, 51, 3865-3873.

[137] Baranska, M.; Schulz, H. Spatial tissue distribution of polyacety-lenes in carrot root. Analyst, 2005, 130, 855-859.

[138] Zidorn, C.; Jöhrer, K.; Ganzera, M.; Schubert, B.; Sigmund, E.M.; Mader, J.; Greil, R.; Ellmerer, E.P.; Stuppner, H. Polyacetylenes from the Apiaceae vegetables carrot, celery, fennel, parsley, and parsnip and their cytotoxic activities. J. Agric. Food Chem., 2005, 53, 2518-2523.

[139] Menelaou, M.A.; Foroozesh, M.; Williamson, G.B.; Fronczek, F.R.; Fischer, H.D.; Fischer, N.H. Polyacetylenes from Chrysoma pauciflosculosa: effects on Florida sandhill species. Phytochem., 1992, 31, 3769-3771.

[140] Pellatia, F.; Calòa, S.; Benvenutia, S.; Adinolfib, B.; Nierib, P.; Melegaria, M. Isolation and structure elucidation of cytotoxic poly-acetylenes and polyenes from Echinacea pallida. Phytochem., 2006, 67, 1359-1364.

[141] Chicca, A.; Pellati, F.; Adinolfi, B.; Matthias, A.; Massarelli, I. Cytotoxic activity of polyacetylenes and polyenes isolated from roots of Echinacea pallida. Br. J. Pharmacol., 2008, 153, 879-885.

[142] Quintana, N.; Weir, T.L.; Du, J.; Broeckling, C.D.; Rieder, J.P.; Stermitz, F.R.; Paschke, M.W.; Vivanco, J.M. Phytotoxic polyace-tylenes from roots of Russian knapweed (Acroptilon repens (L.) DC.). Phytochem., 2008, 69, 2572-2578.

[143] Baranska, M.; Schulz, H.; Christensen, L.P. Structural changes of polyacetylenes in American ginseng root can be observed in situ by using Raman spectroscopy. J. Agric. Food Chem., 2006, 54, 3629-3635.

[144] Christensen, L.P.; Jensen, M.; Kidmose, U. Simultaneous determi-nation of ginsenosides and polyacetylenes in American ginseng root (Panax quinquefolium L.) by high-performance liquid chroma-tography. J. Agric. Food Chem., 2006, 54, 8995-9003.

[145] Liu, J.-H.; Lee, C.-S.; Leung, K.-M.; Yan, Z.-K.; Shen, B.-H.; Zhao, Z.-Z.; Jiang, Z.-H. Quantification of two polyacetylenes in Radix Ginseng and roots of related Panax species using a gas chromatography-mass spectrometric method. J. Agricul. Food Chem., 2007, 55, 8830-8835.

[146] Xu, L.-L.; Han, T.; Wu, J.-Z.; Zhang, Q.-Y.; Zhang, H.; Huang, B.-K.; Rahman, K.; Qin, L.-P. Comparative research of chemical con-

126 Current Analytical Chemistry, 2013, Vol. 9, No. 1 Baranska et al.

stituents, antifungal and antitumor properties of ether extracts of Panax ginseng and its endophytic fungus. Phytomedicine, 2009, 16, 609-616.

[147] Christensen, L.P.; Jensen, M. Biomass and content of ginsenosides and polyacetylenes in American ginseng roots can be increased without affecting the profile of bioactive compounds. J. Nat. Med., 2009, 63, 159-168.

[148] Choi, S.Y.; Ahn, E.-M.; Song, M.-C.; Kim, D.W.; Kang, J.H.; Kwon, O.-S.; Kang, T.-C.; Baek, N.-I. In vitro GABA-transaminase inhibitory compounds from the root of Angelica da-hurica. Phytother. Res., 2005, 19, 839-845.

[149] Lechner, D.; Stavri, M.; Oluwatuyi, M.; Pereda-Miranda, R.; Gib-bons, S. The anti-staphylococcal activity of Angelica dahurica (Bai Zhi). Phytochem., 2004, 65, 331-335.

[150] Randriamampionona, D.; Diallo, B.; Rakotoniriana, F.; Rabeman-antsoa, C.; Cheuk, K.; Corbisier, A.-M.; Mahillon, J.; Ratsi-mamanga, S.; El Jaziri, M. Comparative analysis of active con-stituents in Centella asiatica samples from Madagascar: Applica-tion for ex situ conservation and clonal propagation. Fitoterapia, 2007, 78, 482-489.

[151] De Wit, P.J.G.M.; Kodde, E. Induction of polyacetylenic phytoa-lexins in Lycopersicon esculentum after inoculation with Cla-dosporium fulvum (syn Fulvia fulva). Physiol. Plant Pathol., 1981, 18, 143-148.

[152] Elgersma, D.M.; Weijman, A.C.M.; Roeymans, H.J.; van Eijk, G.W. Occurrence of falcarinol and falcarindiol in tomato plants af-ter infection with Verticillium albo-atrum and characterization of four phytoalexins by capillary gas chromatography-mas spectrome-try. Phytopathol. Z., 1984, 109, 237-240.

[153] Imoto, S.; Ohta, Y. Elicitation of diacetylenic compounds in sus-pension cultured cells of eggplant. Plant Physiol., 1988, 86, 176-181.

[154] Metzger, B.T.; Barnes, D.M.; Reed, J.D. Purple carrot (Daucus carota L.) polyacetylenes decrease lipopolysaccharide-induced ex-pression of inflammatory proteins in macrophage and endothelial cells. J. Agric. Food Chem., 2008, 56, 3554-3560.

[155] Czepa, A.; Hofmann, T. Quantitative studies and sensory analyses on the influence of cultivar, spatial tissue distribution, and indus-trial processing on the bitter off-taste of carrots (Daucus carota L.) and carrot products. J. Agric. Food Chem., 2004, 52, 4508-4514.

[156] Matsunaga, H.; Katano, M.; Yamamoto, H.; Mori, M.; Takata, K. Studies on the panaxytriol of Panax ginseng C. A. Meyer. Isola-tion, determination and antitumor activity. Chem. Pharm. Bull., 1989, 37, 1279-1281.

[157] Fujimoto, Y.; Satoh, M. A new cytotoxic chlorine-containing poly-acetylene from the callus of Panax ginseng. Chem. Pharm. Bull., 1988, 36, 4206-4208.

[158] Wang C.-N.; Shiao Y.-J.; Kuo Y.-H., Chen C.-C.; Lin Y.-L. Induc-ible nitric oxidase synthase inhibitors from SaposhnikoVa diVari-cata and Panax quinquefolium. Planta Med., 2000, 66, 644-647.

[159] Kitagawa, I.; Yoshikawa, M.; Hayashi, T.; Taniyama, T. Chemical studies on crude drug precession. I. On the constituents of ginseng Radix rubra. Yakugaku Zasshi, 1983, 103, 612-622.

[160] Ziurys, L.M. The chemistry in circumstellar envelopes of evolved stars: Following the origin of the elements to the origin of life. Proc. Nat. Acad. Sci., 2006, 103, 12274-12279.

[161] Cernicharo, J.; Heras, A.M.; Tielens, A.G.G.M.; Pardo, J.R.; Her-pin, F.; Guélin, M.; Waters, L.B.F.M. Infrared space observatory's discovery of C4H2, C6H2, and benzene in CRL 618. Astrophys. J. Lett., 1997, 546, L123-L126.

[162] Remijan, A.J.; Hollis, J.M.; Snyder, L.E.; Jewell, P.R.; Lovas, F.J. Methyltriacetylene (CH3C6H) toward TMC-1: The largest detected symmetric top. Astrophys. J., 2006, 643, L37-L40.

[163] Johansson, L.E.B.; Andersson, C.; Ellder, J.; Friberg, P.; Hjalmar-son, A.; Hoglund, B.; Irvine, W.M.; Olofsson, H.; Rydbeck, G. Spectral scan of Orion A and IRC+10216 from 72 to 91 GHz. As-tron. Astrophys., 1984, 130, 227-256.

[164] Walmsley, C.M.; Winnewisser, G.; Toelle, F. Cyanoacetylene and cyanodiacetylene in interstellar clouds. Astronomy and Astrophys-

ics, 1990, 81, 245-250. [165] Remijan, A.J.; Hollis, J.M.; Lovas, F.J.; Plusquellic, D.F.; Jewell,

P.R. Interstellar isomers: The importance of bonding energy differ-ences. Astrophys. J., 2005, 632, 333-339.

[166] Coupeaud, A.; Turowski, M.; Gronowski, M.; Pietri, N.; Couturier-Tamburelli, I.; Kolos, R.; Aycard, J.P. Spectroscopy of cyanodiace-

tylene in solid argon and the generation of isocyano diacetylene. J. Chem. Phys., 2007, 126, 164301-164308.

[167] Miller, F.A.; Lemmon, D.H. The infrared and Raman spectra of dicyanodiacetylene, NCCCCCCN. Spectrochim. Acta, 1967, 23A, 1415-1423.

[168] Kolos, R. Photolysis of dicyanodiacetylene in argon matrices. Chem. Phys. Lett., 1999, 299, 247-251.

[169] Kolos, R. Evidence for the formation of diisocyanodiacetylene in argon matrices. Pol. J. Chem., 2000, 74, 1723-1730.

[170] Strelnikov, D.; Reusch, R.; Krätschmer, W. Infrared and ultraviolet absorptions of matrix isolated C6O2. J. Phys. Chem., 2006, 110A, 12395-12399.

[171] Wegner, G. Topochemische Reaktionen von Monomeren mit kon-jugierten Dreifachbindungen. Z. Naturforsch., 1969, 24B, 824-832.

[172] Baughman, R.H.J. Solid-state synthesis of large polymer single crystals. J. Polym. Sci, Polym. Phys. Ed., 1974, 12, 1511-1535.

[173] Wegner, G. Topochemical reactions of monomers with conjugated triple-bonds. IV. Polymerization of bis-(p-toluene sulfonate) of 2,4-hexadiin-1,6-diol. Makromol. Chem., 1971, 145, 85-94.

[174] Chance, R.R.; Patel, G.N.; Turi, E.A.; Khanna, Y.P. Energetics of the thermal polymerization of a diacetylene crystal. J. Am. Chem. Soc., 1978, 100, 1307-1309.

[175] Bässler, H. Photopolymerization of diacetylene. Adv. Polym. Sci., 1984, 63, 1-48.

[176] Prock, A.; Shand, M.L.; Chance, R.R. Solid-state photopolymeriza-tion of diacetylenes. Macromol., 1982, 15, 238-241.

[177] Shirakawa, H.; Louis, E.J.; MacDiarmid, A.G.; Chiang, C.K.; Heeger, A.J. Synthesis of electrically conducting organic polymers: halogen derivatives of polyacetylene, (CH)x. J. Chem. Soc. Chem. Commun., 1977, 16, 578-580.

[178] Hall, N. Twenty-five years of conducting polymers. Chem. Com-

mun., 2003, 9, 1-4. [179] Sun, A.; Lauher, J.W.; Goroff, N.S. Preparation of poly(diiododi-

acetylene), an ordered conjugated polymer of carbon and iodine. Sci., 2006, 312, 1030-1034.

[180] Schrader, B.; Schulz, H.; Baranska, M.; Andreev, G.N.; Lehner, C.; Sawatzki, J. Non-destructive Raman analyses - polyacetylenes in plants. Spectrochim. Acta, 2005, 61A, 1395-1401.

[181] Pfau, A.S.; Pictet, J.; Plattner, P.; Susz, B. Etudes sur les matières végétales volatiles III. Constitution et synthèse du Carlinoxyde. Helv. Chim. Acta., 1935, 18, 935-951.

[182] Schulz, H.; Baranska, M. Identification and quantification of valu-able plant substances by IR and Raman spectroscopy. Vib. Spec-

trosc., 2007, 43, 13-25. [183] Even, J.; Bertault, M.; Girard, A.; Delugeard, Y.; Fave, J.-L. Dy-

namical study by Raman scattering of the ferroelectric phase transi-tion of the disubstituted diacetylene 1,6-bis (2,4-dinitrophenoxy)-2,4-hexadiyne (DNP). Chem. Phys., 1994, 188, 235-246.

[184] Roman, M.; Dobrowolski, J.Cz.; Baranska, M. Theoretical model-ing of molecular spectra parameters of disubstituted diacetylenes. J. Chem. Inf. Model., 2011, 51, 283-295.

[185] Garrod, B.; Lewis, B.G. Location of the antifungal compound falcarindiol in carrot root tissue. Trans. Br. Mycol. Soc., 1979, 72, 515-517.

[186] Roman, M.; Dobrowolski, J.Cz.; Baranska, M.; Baranski, R. Spec-troscopic studies on bioactive polyacetylenes and other plant com-ponents in wild carrot root. J. Nat. Prod., 2011, 74, 1757-1763.

[187] Metzger, B.T.; Barnes, D.M.J. Polyacetylene diversity and bioac-tivity in orange market and locally grown colored carrots (Daucus carota L). Agric. Food Chem., 2009, 57, 11134-11139.

[188] Frese, L. Resistance of the wild carrot Daucus carota ssp. hispani-cus to the root-knot nematode Meloidogyne-hapla. Gartenbauwis-senschaft, 1983, 48, 259-265.

[189] Roman, M.; Baranski, R.; Baranska, M. Nondestructive Raman analysis of polyacetylenes in Apiaceae vegetables. J. Agric. Food Chem., 2011, 59, 7647-7653.

[190] Jones, H.; Safe, S.; Thaller, V. Natural acetylenes. Part XXIII. A C18 polyacetylenic keto-aldehyde related to falcarinone from an umbellifer (Pastinaca sativa L.). J. Chem. Soc. C, 1966, 52, 1220-1221.

[191] Alvarez, L.; Marquina, S.; Villarreal, M.L.; Alonso, D.; Aranda, E.; Delgado, G. Bioactive polyacetylenes from Bidens pilosa. Planta Med., 1996, 62, 355-357.

[192] Redl, K.; Breu, W.; Davis, B.; Bauer, R. Anti-inflammatory active polyacetylenes from Bidens campylotheca. Planta Med., 1994, 60, 58-62.

Recent Advances in Raman Analysis of Plants Current Analytical Chemistry, 2013, Vol. 9, No. 1 127

[193] Chang, M.-H.; Wang, G.-J.; Kuo, Y.-H.; Lee, C.-K. The low polar constituents from Bidens pilosa L. var. minor (Blume) Sherff. J. Chin. Chem. Soc., 2000, 47, 1131-1136.

[194] Baranska, M.; Schulz, H.; Siuda, R.; Strehle, M.A.; Rösch, P.; Popp, J.; Joubert, E.; Manley, M. Quality control of Harpagophy-tum procumbens and its related phytopharmaceutical products by

means of NIR-FT-Raman spectroscopy. Biopolymers, 2005, 77, 1-8.

[195] Strehle, M.A.; Rösch, P.; Baranska, M.; Schulz, H.; Popp, J. On the way to a quality control of the essential oil of fennel by means of Raman spectroscopy. Biopolymers, 2005, 77, 44-52.

Received: October 05, 2011 Revised: November 28, 2011 Accepted: November 29, 2011