Anal. Chem. 2007, 79, 986-993
Ultrasensitive in situ Tracing of the Alkaloid Dioncophylline A in the Tropical Liana Triphyophyllum peltatum by Applying Deep-UV Resonance Raman Microscopy Torsten Frosch,† Michael Schmitt,† Torsten Noll,‡ Gerhard Bringmann,‡ Karla Schenzel,§ and Ju 1 rgen Popp*,†,|
Institut fu¨r Physikalische Chemie, Friedrich-Schiller-Universita¨t Jena, Helmholtzweg 4, D-07743 Jena, Germany, Institut fu¨r Organische Chemie, Universita¨t Wu¨rzburg, Am Hubland, D-97074 Wu¨rzburg, Germany, Landwirtschaftliche Fakulta¨t, Martin-Luther-Universita¨t Halle-Wittenberg, Ludwig-Wucherer-Strasse 2, D-06108 Halle, Germany, and Institut fu¨r Physikalische Hochtechnologie e.V., Albert Einstein-Strasse 9, 07745 Jena, Germany
UV resonance Raman microspectroscopy was applied for a localization of the antiplasmodial naphthylisoquinoline alkaloid dioncophylline A in very low concentrations in different parts of the samples (e.g., in the roots) of the tropical liana Triphyophyllum peltatum. The application of resonance Raman microspectroscopy was characterized by a very high sensitivity and selectivity. It was possible to assign the resonance Raman spectra of dioncophylline A, dioncophylline C, and dioncopeltine A by means of a combination of NIR Raman spectroscopy and DFT calculations. The UV resonance Raman spectra of T. peltatum are very well resembled by the spectra of dioncophylline A, while they can be clearly distinguished from the spectra of dioncophylline C and dioncopeltine A. This distinction between the various naphthylisoquinolines was possible by the two modes at 1356 and 1613 cm-1. These two modes were assigned to CdC stretching and CH bending vibrations. The presented results of a highly sensitive and selective in situ localization of the active agent dioncophylline A in different parts of the plant material of T. peltatum are of high importance for the acquisition of new antimalarials and for plant science in general. Malaria is a re-emerging infectious disease with tremendous impact on the economical development primarily of sub-Saharan African countries.1,2 This is because of arising resistances against well-established drugs like chloroquine on a global scale. The design and acquisition of new active agents against malaria is therefore of utmost importance. Fortunately traditional medical plants, like the tropical liana Triphyophyllum peltatum from the Ivory Coast, have been used by natives to fight fever for centuries * To whom correspondence should be addressed. E-mail: juergen.popp@ uni-jena.de. † Friedrich-Schiller-Universita¨t Jena. ‡ Universita¨t Wu ¨ rzburg. § Martin-Luther-Universita¨t Halle-Wittenberg. | Institut fu ¨ r Physikalische Hochtechnologie e.V. (1) Sachs, J.; Malaney, P. Nature 2002, 680-685. (2) http://rbm.who.int/wmr2005.
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Figure 1. Chemical structures and numbering scheme of dioncophylline A (1), dioncophylline C (2), and dioncopeltine A (3).
and are therefore a source of new promising active agents. The naphthylisoquinoline alkaloids (NIQs) such as dioncophylline A, dioncophylline C, and dioncopeltine A (see Figure 1) are isolated from these tropical lianas and have been investigated because of their high activity against Plasmodium falciparum. These compounds are structurally related to the quinoline antimalarials chloroquine and quinine and yet are most different, possessing only one nitrogen (a secondary amino function) and exhibiting the phenomenon of axial chirality. The molecular mode of action of the leading drug chloroquine is yet not fully understood; however, an attachment to the small, active growing faces of the malaria pigment hemozoin is considered.3,4 According to this hypothesis, chloroquine might dock onto the hemozoin crystallite by π-π interactions with the porphyrin framework5,6 and prevent further uptake of hematin. In analogy, the NIQ alkaloids might (3) Pagola, S.; Stephens, P. W.; Bohle, D. S.; Kosar, A. D.; Madsen, S. K. Nature 2000, 404, 307-310. (4) Buller, R.; Peterson, M. L.; Almarsson, O ¨ .; Leisirowitz, L. Cryst. Growth Des. 2002, 2, 553-562. (5) Constantinidis, I.; Satterlee, J. D. J. Am. Chem. Soc. 1988, 110, 927-932. (6) Constantinidis, I.; Satterlee, J. D. J. Am. Chem. Soc. 1988, 110, 4391-4395. 10.1021/ac061526q CCC: $37.00
© 2007 American Chemical Society Published on Web 12/23/2006
bind with their isoquinoline part to the surface of hemozoin and block the crystal growth with their extended naphthyl group very efficiently. The interaction between chloroquine and hematin has been studied via in vitro experiments by analyzing the influence of the interaction onto the Raman vibrations of hematin, where significant changes in the depolarization ratio of specific modes were monitored.7 Raman spectroscopy is well suited for the investigation of biological systems where water is present.8-13 Recently we successfully applied NIR (λexc.) 1064 nm) Raman microspectroscopy to localize dioncophylline A in high concentrations inside inclusions of 10-µm size located in the cortex of the stem or the beginning of the leaves of the plant material.14 While Raman spectra of samples of T. peltatum excited in the visible spectral range are obscured by the occurrence of very intensive fluorescence, NIR Raman spectroscopy circumvents the appearance of fluorescence by tuning the Raman excitation into the longer wavelength region, where an excitation of electronic states is no more possible and therefore no fluorescence takes place. Unfortunately, this longer excitation wavelength reduces the intrinsically low Raman scattering cross section even more dramatically, for which reason it is very often not possible to detect very lowconcentration substances in biological samples. However, the advantage of resonantly enhancing the Raman scattering by tuning the excitation wavelength into an electronic absorption of the sample to identify and localize low-concentration samples within plants can be applied. In particular, deep-UV Raman microspectroscopy is characterized by a very high sensitivity and selectivity. Here we report about deep-UV resonance Raman microspectroscopic investigations on various parts of the plant material of T. peltatum. In order to assign the detected resonance Raman signals and interpret the differences in the experimental Raman spectra of the various naphthylisoquinolines, a combination of nonresonant Raman spectroscopy and density functional theory (DFT) calculation has been performed. MATERIALS AND METHODS Chemicals and Plant Material. The naphthylisoquinoline alkaloids dioncophylline A, dioncophylline C, and dioncopeltine A (Figure 1) were isolated as described earlier.15-17 Fresh plant (7) Frosch, T.; Ku ¨ stner, B.; Schlu ¨ cker, S.; Szeghalmi, A.; Schmitt, M.; Kiefer, W.; Popp, J. J. Raman Spectrosc. 2004, 35, 819-821. (8) Petry, R.; Schmitt, M.; Popp, J. ChemPhysChem 2003, 4, 14-30. and references cited therein. (9) Spiro, T. G., Ed. 1988, Biological Applications of Raman Spectroscopy Vol. 1-3, Wiley and references cited therein. (10) Urlaub, E.; Popp, J.; Kiefer, W.; Bringmann, G.; Koppler, D.; Zimmermann, U.; Schneider, H.; Schrader, B. Biospectroscopy 1998, 4, 113-120. (11) Frosch, T.; Schmitt, M.; Bringmann, G.; Kiefer, W.; Popp, J. J. Phys. Chem. B 2006, accepted for publication. (12) Frosch, T.; Schmitt, M.; Popp, J. Anal. Bioanal. Chem. 2006, published online. (13) Frosch, T.; Schmitt, M.; Popp, J. J. Phys. Chem. B 2006, submitted for publication. (14) Frosch, T.; Schmitt, M.; Schenzel, K.; Faber, J. H.; Bringmann, G.; Kiefer, W.; Popp, J. Biopolymers 2006, 82, 295-300. (15) Bringmann, G.; Ru ¨ benacker, M.; Jansen, J. R.; Scheutzow, D. Tetrahedron Lett. 1990, 31, 639-642. (16) Bringmann, G.; Ru ¨ benacker, M.; Weirich, R.; Ake´ Assi, L. Phytochemistry 1992, 31, 4019-4021. (17) Bringmann, G.; Ru ¨ benacker, M.; Vogt, P.; Busse, H.; Ake´ Assi, L.; Peters, K.; von Schnering, H. G. Phytochemistry 1991, 30, 1691-1696.
material of T. peltatum was taken from sterile, whole plants cultivated on a modified Anderson rhododendron medium.18 Thin cross sections (200 µm) of different parts of the plant (leave, steam, roots) were cut manually with a microtome from Leica Microsystems and were used without any further sample preparation. Dioncophylline A is concentrated with 60 ppm in the roots of T. peltatum. Spectroscopy. The nonresonant Raman spectra of the pure isolated naphthylisoquinoline alkaloids were recorded with a Bruker FT Raman spectrometer (RFS 100/S) operated at the macroscopic mode with a spectral resolution of 2 cm-1. A Nd: YAG laser operating at its fundamental wavelength of 1064 nm with an estimated laser power at the samples of 100 mW was used as the Raman excitation source. The Raman scattered light was collected by means of a nitrogen-cooled Ge detector. UV/visible absorption and fluorescence spectra were recorded on a Perkin-Elmer Lambda 16 and a Perkin-Elmer LS50 spectrometer. UV resonance Raman microspectroscopy was performed with an UV Raman setup (HR800 LabRam, Horiba/Jobin-Yvon, focal length of 800 mm and a 2400 lines/mm grating) equipped with an Olympus BX41 microscope, UV-sensitive video camera, and a liquid N2-cooled CCD detector. For UV Raman microscopy, an UV achromatic fused-silica-CaF2 microspot objective (LMU-40×UVB, NA ) 0.5) with broadband UVB coating was chosen. Validation of the wavenumber axis was performed via the wellknown Raman signals of polytetrafluoroethylene. The excitation wavelengths 244 and 257 nm were derived from an intracavity frequency doubled argon ion laser (Innova300-MotoFreD, Coherent Inc.). The laser power at the samples was estimated to be 1 mW. The spectral resolution was 5 cm-1. A handy and robust sample handling (with field practicability) was developed to avoid photodegradation in the UV-Raman experiments. Therefore, a wobbling of the sample material relative to the laser beam across a spot size of ∼5 µm2 and a shutter sequence (control of the laser exposure and the relaxation times) for the laser have been performed. In doing so, a continuous laser exposure of the material has been avoided, while the spatial resolution is still ∼5 µm. Also cooling of the sample material was successfully tried out with the help of a liquid nitrogen-cooled Linkam freezing state, but finally not applied because it was difficult to be combined with the sample movement. The acquisition times for measuring the shown average spectra across a sample spot size of 5 µm2 were 10 min. Density Functional Theory Calculation. DFT calculations were performed with Gaussian 0319 employing the hybrid exchange correlation functionals B3LYP20-23 and B3PW9120,24,25 and the split valence basis set 6-311++G(d,p).26-28 B3LYP and B3PW91 proved to provide reliable estimates of experimental vibrational frequencies of organic molecules showing the lowest root-mean-square deviations.29-35 For all presented results, a more precise integration grid (keyword tight) was applied, being essential for DFT calculations. The DFT calculated harmonic vibrational frequencies are typically too large to be compared with the experimentally observed ones due to the neglect of anharmonicity, incomplete incorporation of electron correlation, and application of finite basis sets. Fortunately, the overestimation of (18) Bringmann, G.; Rischer, H. Plant Cell Rep. 2001, 20, 591-595.
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the calculated harmonic vibrational frequencies is relatively uniform and this problem can be circumvented by applying transferable scaling factors directly to the harmonic frequencies (especially in case of B3PW91 and B3LYP).30,35,36 For the chosen model chemistry B3LYP/6-311++G(d,p), values of 0.979 for modes below 1800 cm-1 and 0.96 above 1800 cm-1 were applied as recommended in the literature.37,38 Since the DFT calculations with B3PW91/6-311++G(d,p) showed results very similar to the calculations performed with B3LYP/6-311++G(d,p), the results of the calculations with B3PW91 are not shown. Based on the DFT calculated Raman scattering activities, the Raman intensities were calculated and further convoluted with Gauss-Lorentz weighted profiles to simulate the experimental Raman spectra with a finite bandwidth. RESULTS AND DISCUSSION UV Resonance Raman Spectroscopy. Recently we have reported about a NIR FT Raman microspectroscopic localization of dioncophylline A (Figure 1) present in high concentrations in inclusions located in the cortex of the stem material of T. peltatum.14 Outside of these spots, however, it was not possible to detect even traces of dioncophylline A due to the low sensitivity of NIR Raman spectroscopy and because the laser power needed to obtain Raman spectra of the bright inclusions was too high for the plant material outside, which was destroyed under these conditions. Furthermore, it was not possible to investigate the (19) Gaussian 03, Revision C.02, Frisch, M. J.; Trucks, G. W.; Schlegel, H. B.; Scuseria, G. E.; Robb, M. A.; Cheeseman, J. R.; Montgomery, J. A., Jr.; Vreven, T.; Kudin, K. N.; Burant, J. C.; Millam, J. M.; Iyengar, S. S.; Tomasi, J.; Barone, V.; Mennucci, B.; Cossi, M.; Scalmani, G.; Rega, N.; Petersson, G. A.; Nakatsuji, H.; Hada, M.; Ehara, M.; Toyota, K.; Fukuda, R.; Hasegawa, J.; Ishida, M.; Nakajima, T.; Honda, Y.; Kitao, O.; Nakai, H.; Klene, M.; Li, X.; Knox, J. E.; Hratchian, H. P.; Cross, J. B.; Adamo, C.; Jaramillo, J.; Gomperts, R.; Stratmann, R. E.; Yazyev, O.; Austin, A. J.; Cammi, R.; Pomelli, C.; Ochterski, J. W.; Ayala, P. Y.; Morokuma, K.; Voth, G. A.; Salvador, P.; Dannenberg, J. J.; Zakrzewski, V. G.; Dapprich, S.; Daniels, A. D.; Strain, M. C.; Farkas, O.; Malick, D. K.; Rabuck, A. D.; Raghavachari, K.; Foresman, J. B.; Ortiz, J. V.; Cui, Q.; Baboul, A. G.; Clifford, S.; Cioslowski, J.; Stefanov, B. B.; Liu, G.; Liashenko, A.; Piskorz, P.; Komaromi, I.; Martin, R. L.; Fox, D. J.; Keith, T.; Al-Laham, M. A.; Peng, C. Y.; Nanayakkara, A.; Challacombe, M.; Gill, P. M. W.; Johnson, B.; Chen, W.; Wong, M. W.; Gonzalez, C.; Pople, J. A. Gaussian, Inc., Wallingford CT, 2004. (20) Becke, A. D. J. Chem. Phys. 1992, 97, 9173 (21) Becke, A. D. J. Chem. Phys. 1993, 98, 5648 (22) Stephens, P. J.; Devlin, F. J.; Chabalowski, C. F.; Frisch, M. J. J. Phys. Chem. 1994, 98, 11623-11627. (23) Lee, C.; Yang, W.; Parr, R. G. Phys. Rev. 1988, B37, 785-789. (24) Perdew, J. P.; Wang Y. Phys. Rev. 1992, B45, 13244-13249. (25) Perdew, J. P.; Chevary, J. A.; Vosko, S. H.; Jackson, K. A.; Pederson, M. R.; Singh, D. J.; Fiolhais, C. Phys. Rev. B 1992, 46, 6671-6687. (26) Pople, J. A.; Schlegel, H. B.; Krishnan, R.; Defrees, D. J.; Binkley, J. S.; Frisch, M. J.; Whitside, R. A. Int. J. Quantum Chem.: Quantum Chem. Symp. 1981, 15, 269-278. (27) Frisch, M. J.; Pople, J. A.; Binkley, J. S. J. Chem. Phys. 1984, 80, 32653269, and references therein. (28) Hehre, W. J.; Stewart, R. F.; Pople, J. A. J. Chem. Phys. 1969, 51, 26572664, and references therein (29) Neugebauer, J.; Hess, B. A. J. Chem. Phys. 2003, 118, 7215-7225. (30) Scott, A. P.; Radom, L. J. Phys. Chem. 1996, 100, 16502-16513. (31) El-Azhary, A. A.; Suter, H. U. J. Phys. Chem. 1996, 100, 15056-15063. (32) Baker, J.; Jarzecki, A. A.; Pulay, P. J. Phys. Chem. A 1998, 102, 1412-1424. (33) Rauhut, Guntram; Pulay, P. J. Phys. Chem. 1995, 99, 3093-3100. (34) Halls, M. D.; Schlegel, B. J. Chem. Phys. 1998, 109, 10587-10593. (35) Andersson, M. P.; Uvdal, P. J. Phys. Chem. A 2005, 109, 2937-2941. (36) Wong, M. W. Chem. Phys. Lett. 1996, 256, 391-399. (37) Bauschlicher, C. W.; Langhoff, S. R. Spectrochim. Acta. Part A 1997, 53, 1225-1240. (38) Halls, M. D.; Velovski, J.; Schlegel, H. B. Theor. Chem. Acta 2001, 105, 413-421.
988 Analytical Chemistry, Vol. 79, No. 3, February 1, 2007
Figure 2. Absorption (A) and fluorescence (F) spectra of 1. Spectra of 2 and 3 are very similar. Positions of the UV excitation wavelengths and wavenumber ranges of 4000 cm-1 (typical Raman spectra) are marked by arrows.
dark-colored root material at all. Therefore, in this contribution, we take advantage of the fact that the Raman scattering is significantly enhanced if Raman excitation wavelengths in the deep-UV spectral range are applied due to the ω4-dependency and an electronic resonance enhancement of the Raman scattering intensity. For the investigation of biological samples, resonance Raman spectroscopy offers several advantages over nonresonant Raman spectroscopy: (I) The Raman scattered light is enhanced up to a factor of 108 if an electronic eigenstate of the molecule is involved in the scattering process (resonance condition). (II) By selectively tuning the excitation wavelength through the electronic absorptions, it is possible to enhance specific molecular vibrations coupled to the resonant electronic transition and therefore to detect exclusively the chromophore of interest (in our case the active agent) and to suppress background signals due to other ingredients (in our case of the surrounding plant material). For our Raman studies, we applied the two deep-UV excitation wavelengths of 244 and 257 nm. Both wavelengths are marked by vertical arrows in the absorption spectrum of dioncophylline A (Figure 2). The electronic absorption spectra of the two other naphthylisoquinolines investigated, dioncophylline C and dioncopeltine A, are very similar and are therefore not shown explicitly. Figure 2 shows that an excitation wavelength of 244 nm should lead to resonance Raman spectra while an excitation wavelength of 257 nm would be expected to give more structured preresonance Raman spectra. The fluorescence spectrum of dioncophylline A obtained for an excitation into the maximum of the absorption spectra at 228 nm is also depicted in Figure 2 and shows one of the major advantages of deep-UV resonance Raman spectroscopy, namely, that the respective spectra are virtually never obscured by fluorescence signals. This great advantage of deep-UV resonance Raman spectroscopy is nicely demonstrated in Figure 2. While a relative Raman shift of ∼4000 cm-1 excited with the deep-UV wavelengths 244 and 257 nm corresponds to ranges of 244-270 and 257-286 nm, respectively, no fluorescence emission occurs below 320 nm. This is because deep-UV light leads to an excitation of highly excited electronic states while fluorescence typically takes place from the first electronically
Figure 3. UV resonance Raman spectra (λexc.) 244 nm) of fresh root sample material of T. peltatum and spectra of pure 1, 2, and 3. Intensities of spectra of solutions of 1-3 are stretched by a factor of 5 from 500 to 1300 cm-1 for better illustration, and the spectrum of T. peltatum is stretched by a factor of 10 from 500 to 1300 cm-1. (A) UV resonance Raman spectrum of extracted 3; (B) UV resonance Raman spectrum of extracted 2; (C) UV resonance Raman spectrum of extracted 1; (D) UV resonance Raman spectrum of fresh plant material of T. peltatum.
excited state. Therefore, Raman spectra and fluorescence spectra are well separated from each other and Figure 2 clearly shows that the UV resonance Raman spectra of dioncophylline A excited at wavelengths of 244 and 257 nm are definitely free of any fluorescence signal. Root material of T. peltatum was inaccessible by NIR Raman spectroscopy in an earlier study,14 because the high irradiation needed caused damage to the samples. The increased sensitivity due to the resonance enhancement by application of UV excitation wavelengths, however, made it possible to acquire signals with high signal-to-noise ratio from different (random) parts of the roots. Also, the same spectra have been measured from other parts of the plant, e.g., the stem. The UV resonance Raman spectra of root samples of T. peltatum (as representative examples for the plant material) as well as of the pure NIQs dioncophylline A, dioncophylline C, and dioncopeltine A excited at 244 and 257 nm are shown in Figures 3 and 4. The resonance Raman spectra for both excitation wavelengths show a strong enhancement of vibrations within the wavenumber range between 1350 and 1650
Figure 4. UV resonance Raman spectra (λexc.) 257 nm) of fresh root sample material of T. peltatum and spectra of pure 1, 2, and 3. Intensities of spectra of solutions of 2 and 3 are stretched by a factor of 5 from 500 to 1320 cm-1 for better illustration, and the spectra of 1 and T. peltatum are stretched by a factor of 10 from 500 to 1320 cm-1. (A) UV resonance Raman spectrum of extracted3; (B) UV resonance Raman spectrum of extracted 2; (C) UV resonance Raman spectrum of extracted 1; (D) UV resonance Raman spectrum of fresh plant material of T. peltatum.
cm-1. However, the resonance Raman spectra recorded under preresonance conditions (λexc ) 257 nm; Figure 4) show more Raman bands; the selectivity is thus less rigorous than for a fully resonant excitation (see Figure 3; λexc ) 244 nm). Figures 3 and 4 show that the resonance Raman spectrum of T. peltatum (spectrum D) well resembles the spectrum of dioncophylline A (spectrum C). The resonance Raman spectra of dioncophylline C (spectrum B) and dioncopeltine A (spectrum A) can be clearly distinguished from the resonance Raman spectra of T. peltatum (spectrum D) and dioncophylline A (spectrum C) by the prominent mode at 1355 cm-1 for T. peltatum and dioncophylline A, which is shifted to 1367 cm-1 in the case of dioncophylline C and to 1370 cm-1 in the case of dioncopeltine A. Furthermore, the mode at 1613 cm-1 for T. peltatum and dioncophylline A is shifted to 1624 cm-1 in the case of dioncophylline C and dioncopeltine A. These experiments nicely show that even very low amounts of only ∼60 ppm by weight of dioncophylline A located in the roots of T. peltatum can be in situ detected by means of deep-UV resonance Raman microspectroscopy. No fluorescence signal Analytical Chemistry, Vol. 79, No. 3, February 1, 2007
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obscures the Raman spectrum, nor does any Raman signal from the surrounding plant material play an important role because of the selective resonance condition. Compared to the previously reported NIR FT Raman microscopic studies, the application of UV resonance Raman spectroscopy allows for a straightforward detection of low concentrations of the active agent in plant regions where no Raman information at all had previously been retrieved by NIR Raman excitation. Furthermore, UV resonance Raman spectroscopy is capable of an in situ differentiation between the structurally very similar active agents dioncophylline A, dioncophylline C, and dioncopeltine A (Figure 1). NIR FT Raman Spectroscopy and DFT Calculation of the Raman Spectra. In order to assign the resonantly enhanced Raman bands of the naphthylisoquinolines, quantum chemical calculations of the Raman spectra of the three active agents were performed. In combination with experimental nonresonant Raman spectra of these molecules, the differences in the resonance Raman spectra were fully interpretable. Figure 5 presents a Raman spectroscopic characterization of dioncophylline A (Figure 5C), dioncophylline C (Figure 5B), and dioncopeltine A (Figure 5A). The experimental nonresonant (λexc.) 1064 nm) Raman spectra are shown in the upper parts (A1, B1, C1) of Figure 5 together with the simulated Raman spectra with finite bandwidth in the central parts (A2, B2, C2) and the calculated (B3LYP/6-311++G(d,p)) Raman stick spectra in the lower parts (A3, B3, C3). The experimental nonresonant (λexc ) 1064 nm) Raman spectrum (Figure 5 C1) and the simulated Raman spectrum (Figure 5 C2) of dioncophylline A agree well, thus enabling a detailed mode assignment of the Raman bands shown in Figure 5 C1. The calculated stick spectrum of dioncophylline A (Figure 5 C3) demonstrates that some of the Raman bands in the low-wavenumber region have contributions from different molecular normal modes. However, the intense Raman signals of dioncophylline A around 1355 and 1613 cm-1 can be unambiguously assigned to one particular normal mode. Figures 5B and A display the results of a Raman spectroscopic characterization of dioncophylline C and dioncopeltine A. The three structurally very similar active agents can nicely be distinguished from each other by means of their NIR Raman spectra (Figures 5 A1, B1, C1). These experimental vibrational spectra of all three compounds resemble well the calculated ones (Figures 5 A2, B2, C2) permitting one to explain the observed differences in the experimental nonresonant and resonant Raman spectra of these three naphthylisoquinolines. The calculated Raman spectra of dioncophylline A (Figure 5 C2), dioncophylline C, and dioncopeltine A (Figure 5 B2, A2) also show the significant mode shifts in respect to each other within the wavenumber region around 1360 and 1610 cm-1 observed in the UV resonance Raman spectra (see Figures 2 and 3) used to differentiate these three NIQs. Furthermore, some Raman bands above 1615 cm-1, which cannot be resolved clearly in the experimental nonresonant Raman spectra of dioncophylline C and dioncopeltine A (Figure 5 A1, A2 and B1, B2), have their origin in the calculated stick spectra of dioncophylline C and dioncopeltine A (Figure 5 A3, B3) and are very intense in the UV Raman spectra (e.g., band position 1624 cm-1 in Figures 3 and 4) due to a strong resonance enhancement. Utilizing the good agreement between the calculated Raman spectra and the experimental ones of the investigated naphthyl990 Analytical Chemistry, Vol. 79, No. 3, February 1, 2007
Figure 5. Raman spectroscopic characterization of dioncophylline A (C1-C3), dioncophylline C (B1-B3), and dioncopeltine A (A1A3). (A1) Experimental FT Raman spectrum of dioncopeltine A. (A2) Simulated Raman spectrum of dioncopeltine A. (A3) DFT calculation of the Raman scattering activity of dioncopeltine A (B3LYP/6311++G(d,p)). (B1) Experimental FT Raman spectrum of dioncophylline C. (B2) Simulated Raman spectrum of dioncophylline C. (B3) DFT calculation of the Raman scattering activity of dioncophylline C (B3LYP/6-311++G(d,p)). (C1) Experimental FT Raman spectrum of dioncophylline A. (C2) Simulated Raman spectrum of dioncophylline A. (C3) DFT calculation of the Raman scattering activity of dioncophylline A (B3LYP/6-311++G(d,p)).
isoquinolines, the most prominent Raman peaks of dioncophylline A are discussed in detail in the following section. To get more
Figure 6. Atomic displacements of calculated prominent vibrational modes of dioncophylline A.
insight into the vibrations, the atomic displacements of the most prominent normal modes of dioncophylline A are shown in Figure 6. An understanding of the atomic displacements is of particular importance for an interpretation of upcoming experiments where the docking process of the active agents to the biological target hemozoin is studied in terms of monitoring the changes of the sensitive Raman peaks. Assignment of the Raman Spectra and Discussion of the Atomic Displacements. The most intense Raman peak in the UV resonance Raman spectra of T. peltatum (Figures 3D, 4D) is located at 1613 cm-1 and is reproduced in the NIR Raman spectrum of dioncophylline A at 1611 cm-1 (Figure 5 C1). This molecular vibration can be assigned to a CdC stretch vibration
in the Q1 part of the isoquinoline unit, where νC(8)dC(9) and νC(6)dC(5) stretch in phase, to a stretching of the biaryl bridge bond νC(7)sC(1′) and to bendings vibrations (Figure 6). The vibration at 1580 cm-1 in the NIR Raman spectrum of dioncophylline A (Figure 5 C1) also present in the UV resonance Raman spectra of T. peltatum (Figures 3D and 4D) and of all three naphthylisoquinolines (Figures 3A, B, C and 4A, B, C) is assigned to a very symmetric stretching mode in the naphthalene unit, where νC(3′)dC(2′), νC(10′)dC(9′), and νC(6′)dC(7′) compress in phase, while the atoms C(4′), C(5′), C(1′), and C(8′) move outward at the same time (Figure 6). There are also CH-bending motions as well as smaller νCdC and δCH motions within the isoquinoline unit contributing to this mode. Analytical Chemistry, Vol. 79, No. 3, February 1, 2007
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Table 1. Wavenumbers and Vibrational Assignment of the Raman Modes of Dioncophylline A (1)a ˜ν/cm-1
NIQ
vibrational assignment
3295
(1)
νN(2)H
3200-2700
(1)
localized νsCH, νasCH isoquinoline and naphthalene (2734 cm-1 : νC(1)H)
1624
(2), (3)
νCdC, δipCH, δsCH3 and δsOCH3 in naphthalene
1611
(1)
stretching mode in isoquinoline ring mostly Q1 where νC(8)dC(9) and νC(6)dC(5) stretch in phase; δipC(6)H, δipC(5)H, δ(13)H; νC(7)-C(1′) bridge bond
1604
(2)
νCdC in isoquinoline (Q1); δipCH, dO(13)H, nC(7)-C(1') bridge bond
1580
(1)
very symmetric stretching mode in naphthalene where νC(3′)dC(2′), νC(10′)dC(9′) and νC(6′)dC(7′) compress in phase, while atoms C(4′), C(5′), C(1′) and C(8′) move outward at the same time; CH bendings: δipC(6′)H, δipC(7′)H, δipC(3′)H, δsOC(13′)H3, δsOC(15′)H3, ωC(11′)H3; smaller νCdC and δCH in isoquinoline
1512
(1)
νCdC in naphthalene; δipC(6′)H, δipC(7′)H, δipC(8′)H, δsOC(13′)H3, δsOC(15′)H3, ωC(11′)H3
1460
(1)
ωCH3 in naphthalene
1430
(1)
δsCH3, δipCH in naphthalene and isoquinoline
1395
(3)
strong νCdC in naphthalene; νC(5′)-O(14′), νC(7)-C(1′) bridge bond, δipCH, δsOC(14′)H3, ωC(11′)H2, δipO(12')H
1381
(1)
very strong δsC(11′)H3; δipCH, δsOCH3 and νCdC in naphthalene; δsC(11)H3
1355
(1)
very strong ωC(4)H2 and synchronic δC(3)H; δsC(12)H3 in isoquinoline; νCdC in naphthalene N1 (νC(3′)dC(4′), νC(1′)dC(9′), νC(4′)dC(10′), νC(10′)dC(9′), νC(1′)dC(2′)); δipC(8′)H, δipC(6′)H, δsC(11′)H3, δsOC(13′)H3, νC(4′)-O(12′)
1338
(1)
symmetric νCdC in naphthalene where νC(3′)dC(2′), νC(6′)dC(7′), νC(4′)dC(10′), νC(1′)dC(9′), νC(10′)dC(5′) and νC(9′)dC(8′) stretch in phase while νC(10′)dC(9′), νC(5′)dC(6′) and νC(8′)dC(7′) compress at the same time; νC(4′)-O(12′); δsC(11′)H3, δsC(11′)H3, smaller δCH in isoquinoline
1315
(1)
νCdC in isoquinoline Q1; δCH, δOH, δNH
1295
(1)
ring breathing in isoquinoline (Q1) and naphthalene; νC(4′)O(12′), νC(5′)O(14′); symmetric δipC(6′)H, δipC(7′)H, δipC(8′)H
1240
(1)
ring breathing in isoquinoline (Q1); δipC(5)H, δipC(6)H, δipC(1)H, δipC(3)H; δ(2)H, δ(13)H, small asymmetric ring breathing in naphthalene
1174
(1)
small ring breathing in naphthalene; asymmetric δipC(7′)H and δipC(8′)H; ωC(15′)H3
1115
(1)
ring breathing in isoquinoline; wagging ωC(11)H3, ωC(13′)H3, ωC(15′)H3; νC(4′)O(12′), νC(5′)O(14′), νC(8)O(13); δipCH
966
(1)
ring breathing in naphthalene and in isoquinoline; δoopC(5)H, δoopC(6)H; νC-C in naphthalene; νC(4′)O(12′),νC(5′)O(14′); rocking FC(11′)H3, FC(11)H3, FC(12)H3
788
(1)
δoopC(5)H, δoopC(6)H; δoopN(2)H; FC(4)H2, FC(11)H3, FC(12)H3
729
(1)
asynchronous ring breathing in naphthalene and isoquinoline (Q1); νCC and νCN in isoquinoline; δN(2)H, FC(4)H2, FC(11)H3, FC(13′)H3, FC(15′)H3, νC(5′)O(14′)
690
(1)
out-of-plane bending in naphthalene and isoquinoline rings; δoopCH, δoopOH, FCH3, FCH3
636
(1)
νCC in naphthalene; δCOC at OMe in naphthalene; ring breathing in isoquinoline
595
(1)
out-of-plane bending in naphthalene and isoquinoline; δCO(14′)C; δoopN(2)H; δoopCH in isoquinoline
561
(1)
out of plane bending in naphthalene and isoquinoline; δCOC at OMe in naphthalene
415
(1)
out of plane bending in naphthalene and isoquinoline; δCOC at OMe in naphthalene; δoopN(2)H
358
(1)
out of plane bending “butterfly” in isoquinoline; δCO(14′)C
286
(1)
twisting τOC(Me) in naphthalene; δoopO(13)H
a The atomic numbering scheme is used as shown in Figure 1. Some of the most important modes of dioncophylline C (2) and diocopeltine A (3) are also included. The most prominent modes in the Raman spectra are marked in boldface type and their atomic displacements are shown in Figure 6.
The band at 1381 cm-1 in the Raman spectra of dioncophylline A (Figures 3C, 4C, 5 C1), which is also present in the spectra of dioncophylline C (Figures 3B, 4B, 5 B1), but not in the spectra of dioncopeltine A (Figures 3A, 4A, 5 A1), can be assigned to a very strong scissoring, motion, to bending and smaller νCdC motions in the naphthalene unit as well as to a scissoring vibration in Q2 (Figure 6). The most intense vibration in the Raman spectra of dioncophylline A (Figures 3C, 4C, 5 C1), located at 1355 cm-1, is also seen very well in the resonance Raman spectra of T. peltatum 992
Analytical Chemistry, Vol. 79, No. 3, February 1, 2007
(Figures 3D, 4D) but is shifted in the spectra of dioncophylline C (Figures 3B, 4B, 5 B1) and dioncopeltine A (Figures 3A, 4A, 5 A1) to 1367 and 1370 cm-1, respectively. This mode has been previously used in NIR Raman microscopy8 experiments to distinguish between the various naphthylisoquinolines and was assigned to very strong wagging and synchronic bending motions in the isoquinoline unit of dioncophylline A as well as to CdC and CsO stretching and bending vibrations within the naphthalene ring (see Figure 6).
The intense mode at 1338 cm-1 in the Raman spectra of dioncophylline A (Figures 3C, 4C, 5 A1) is assigned to a symmetric νCdC vibration in the naphthalene ring (Figure 6), where νC(3′)dC(2′), νC(6′)dC(7′), νC(4′)dC(10′), νC(1′)dC(9′), νC(10′)d C(5′), and νC(9′)dC(8′) stretch in phase, while νC(10′)dC(9′), νC(5′)dC(6′), and νC(8′)dC(7′) compress at the same time, and to bending motions. The low-wavenumber mode at 729 cm-1 in the resonance Raman spectra of dioncophylline A (Figures 3C, 4C) can also be applied to distinguish diocophylline A (Figures 3D, 4D) from dioncophylline C (Figures 3B, 4B) and dioncopeltine A (Figures 3A, 4A) in T. peltatum. This mode is due to an asynchronous ringbreathing motion in the naphthalene and isoquinoline units (Q1), to νCC and νCN vibrations in isoquinoline, and to various bending vibrations (Figure 6). The out-of-plane bending mode in dioncophylline A (Figure 6) at 690 cm-1 is found in the Raman spectra of dioncophylline A (Figures 3D, 4D, 5 C1), dioncopeltine A (Figures 3A, 4A, 5 A1) and T. peltatum (Figures 3D, 4D), but not in the spectra of dioncophylline C (Figures 3B, 4B, 5 B1). The detailed mode assignment is summarized in Table 1. CONCLUSION AND OUTLOOK This contribution reports about the application of deep-UV resonance Raman spectroscopy for a very sensitive and selective in situ localization of low concentrations of dioncophylline A in different parts of the tropical liana T. peltatum. This sensitive localization was possible by using the advantages of the resonance Raman effect. Because the naphthylisoquinoline alkaloids exhibit strong electronic absorptions in the deep-UV spectral range, the Raman excitation wavelengths of 244 and 257 nm were successfully applied to resonantly enhance the Raman signals of selective vibrations of dioncophylline A in situ in T. peltatum. This resonance enhancement made it possible to record signals with very high signal-to-noise ratio by application of very low laser powers. Furthermore, the UV Raman spectra of T. peltatum are
free of any fluorescence from the surrounding plant material. By comparison, the UV resonance Raman spectra of T. peltatum with the spectra of the pure active agents dioncophylline A, dioncophylline C, and dioncopeltine A is was possible to unambiguously localize dioncophylline A in different parts of T. peltatum and to distinguish the Raman spectra of the tropical liana from the spectra of the alkaloids dioncophylline C and dioncopeltine A. These differences within the resonance Raman spectra were assigned to CdC stretching and CH bending vibrations by means of DFT calculations of the Raman spectra of the three naphthylsioquinoline alkaloids. The presented results demonstrate the great potential of UV resonance Raman microspectroscopy for acquiring new antimalaria active agents and for investigating plants in general. By selectively enhancing vibrational modes of the active agents that are expected to be sensitive to possible π-π interactions to the biological target hemozoin in upcoming in vitro and in vivo (inside the food vacuole of P. falciparum) Raman experiments, it seems to be promising to monitor influences onto these molecular vibrations due to the docking process. Thus, it will hopefully be possible to understand the restrictions onto the molecular design of the antimalarials and help in the fight against malaria. Investigations toward this goal are currently underway in our laboratory. ACKNOWLEDGMENT The authors gratefully acknowledge the financial support from the Fonds der Chemischen Industrie and the Deutsche Forschungsgemeinschaft (Sonderforschungsbereich 630 “Recognition, Preparation, and Functional Analysis of Agents Against Infectious Diseases”, projects C1 and A2).
Received for review August 16, 2006. Accepted November 16, 2006. AC061526Q
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