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1 July 2010 Structural analysis of the antimalarial drug halofantrine by means of Raman spectroscopy and density functional theory calculations
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Abstract
The structure of the antimalarial drug halofantrine is analyzed by means of density functional theory (DFT) calculations, IR, and Raman spectroscopy. Strong, selective enhancements of the Raman bands of halofantrine at 1621 and 1590 cm−1 are discovered by means of UV resonance Raman spectroscopy with excitation wavelength exc=244 nm. These signal enhancements can be exploited for a localization of small concentrations of halofantrine in a biological environment. The Raman spectrum of halofantrine is calculated by means of DFT calculations [B3LYP/6-311+G(d,p)]. The calculation is very useful for a thorough mode assignment of the Raman bands of halofantrine. The strong bands at 1621 and 1590 cm−1 in the UV Raman spectrum are assigned to combined CC stretching vibrations in the phenanthrene ring of halofantrine. These bands are considered as putative marker bands for interactions with the biological target molecules. The calculation of the electron density demonstrates a strong distribution across the phenanthrene ring of halofantrine, besides the electron withdrawing effect of the Cl and CF3 substituents. This strong and even electron density distribution supports the hypothesis of stacking as a possible mode of action of halofantrine. Complementary IR spectroscopy is performed for an investigation of vibrations of polar functional groups of the halofantrine molecule.

1.

Introduction

Malaria is a major human parasitic disease and affects approximately 40% of the world population.1, 2, 3 The mortality and morbidity from the disease strongly increased in the recent decades. The disease has tremendous effects on the economic development primarily of sub-Saharan countries.1, 2 The control of malaria was declared as a global priority by the World Health Organization2 (WHO). The main reason for this disaster is a rise of worldwide resistances against existing drugs, most of all against chloroquine.1, 2, 3, 4 Chloroquine was the mainstay therapy for decades and is one of the most important drugs used against an infectious disease.5, 6 To address these problems with resistances, more effort is required in the development of novel antimalarials.3, 4 In particular, a better understanding of the molecular mode of action of the active agents is required for the design of structure-based drugs.

Halofantrine is a drug that is effective against chloroquine-resistant strains of Plasmodium falciparum 7 and Plasmodium vivax.8 Halofantrine was first synthesized by Colwell 9 in an army research program on malaria, within a large series of phenanthrenes, quinolines, and related arylaminoethanols. The drug is marketed by GlaxoSmithKline under the name Halfan. Halofantrine was also successfully applied in a pediatric study.10 The drug shows enhanced activity in combination with chloroquine11 and is especially prescribed against a high incidence of multidrug resistances. 12

However, the molecular mode of action of halofantrine is not well understood. The drug interferes with the detoxification process of hemoglobin digestion by-products in the 48-h cycle of the red blood cell state of the malaria parasite Plasmodium.13, 14, 15, 16, 17, 18 It was shown that protein mutations in P. falciparum confer resistances to halofantrine.19 Recently a crystal structure of a complex of halofantrine and ferriprotoporphyrin IX (FPPIX) was derived and it was found that ππ stacking of the phenanthrene ring of the drug and the porphyrin of the target structure may play a major role in the biological activity.20, 21 Halofantrine inhibits the formation of the malaria pigment hemozoin. Therefore, a selective binding to the small, active growing faces of the hemozoin crystallites may be considered as a mode of action.22, 23 Halofantrine showed essential chemical features in a pharmacophore model for FPPIX-targeting antimalarials.24

New diagnostic techniques are required to elucidate the molecular mode of action of halofantrine in vitro as well as in Plasmodium. Therefore, the fast, easy, and reliable localization of the drug in the biological environment is of high importance. Halofantrine was determined in rat25 and human12, 26 plasma by means of high-performance liquid chromatography (HPLC). However, a localization of halofantrine in life cells with high spatial resolution would be desired to address the specific binding sides of the drugs in Plasmodium. Novel photonic diagnostic techniques were recently developed and exciting results were achieved in biomedical research.27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37 The application of such biophotonic techniques has high potential to elucidate the molecular mode of action of halofantrine and to contribute in a structure-based design of new, effective drugs against malaria.

2.

Material and Methods

2.1.

Chemicals

Halofantrine hydrochloride was a kind gift of GlaxoSmithKline and was used without further purification. The systematic IUPAC name of halofantrine is 3-dibutylamino-1-[1,3-dichloro-6-(trifluoromethyl) phenanthren-9-yl]-propan-1-ol. The molecular structure of halofantrine is shown in Fig. 1 . The atomic numbering scheme and the partitioning of the phenanthrene ring in P1, P2 and P3 in Figure 1 is used in the context for the description of the mode assignment.

Fig. 1

Molecular structure of halofantrine. The atomic numbering scheme and the partitioning of the phenanthrene ring in P1, P2, and P3 is used in the context for the description of the mode assignment.

041516_1_005004jbo1.jpg

2.2.

Spectroscopy

The Fourier transform (FT) Raman spectrum of halofantrine hydrochloride was recorded with a Bruker FT Raman spectrometer (RFS 100/S) at the macroscopic mode with a spectral resolution of 2cm1 . The instrument was equipped with a Nd:YAG laser ( λexc=1064nm , estimated laser power at the samples P=50mW ) as the excitation source and a liquid-nitrogen-cooled germanium detector.

The complementary FT IR spectrum of halofantrine hydrochloride was measured as KBr pellet using a Bruker IFS 66 spectrometer equipped with a DTGS (doped triglycerinsulfate) detector and with 4-cm1 spectral resolution.

UV resonance Raman microspectroscopy was performed with an UV Raman setup (HR800 LabRam, Horiba/Jobin-Yvon, focal length of 800mm and a 2400linesmm grating) equipped with an Olympus BX41 microscope, UV sensitive video camera, and a liquid- N2 -cooled CCD detector. For UV microscopy, a UV achromatic fused silica/ CaF2 microspot objective [LMU- 40× -UVB, numerical aperture (NA)=0.5 ] with broad band UVB coating was chosen. Validation of the wave number axis was performed via the Raman signals from Teflon. The excitation wavelength 244nm was derived from an intracavity frequency-doubled argon-ion laser (Innova300-MotoFreD, Coherent Inc.). The laser power at the sample was estimated to be 1mW . The spectral resolution was 5cm1 . Because the laser wavelength was chosen to excite the sample in an electronic absorption band, it was necessary to carefully avoid any sample destruction. Furthermore, the sample rotation technique37, 38 was applied.

2.3.

Density Functional Theory Calculation

Density functional theory (DFT) calculations were performed with Gaussian 03 (revision D.01)39 with Becke’s three-parameter exchange functional40 (B3) as slightly modified by Stephens,41 coupled with the correlation functional of Lee 42 (B3LYP) and B3 in combination with the correlation part of the functional from Perdew and Wang43 and Perdew 44 (B3PW91). Double (6-31+G(d,p)) and triple (6-311+G(d,p)) split valence basis sets of contracted Gaussian functions with polarized and diffuse functions45, 46, 47 were applied. These hybrid exchange correlation functionals provide reliable estimates of experimental frequencies of organic molecules with small root mean square deviations.48, 49, 50, 51, 52, 53

The DFT calculated harmonic vibrational frequencies are typically too large compared with the experimentally observed ones, due to neglect of anharmonicity, incomplete incorporation of electron correlation, and the use of finite basis sets. Fortunately, this overestimation of the calculated harmonic vibrational frequencies is relatively uniform and can be circumvented by applying transferable scaling factors to the harmonic frequencies.48, 54 For the B3LYP6-311+G(d,p) calculations, values of 0.98 for modes below 1800cm1 and 0.96 above 1800cm1 were applied, in agreement with the literature.54, 55, 56, 57, 58, 59, 60

The calculated Raman activities were transformed into Raman intensities61 and were further convoluted with Gauss-Lorentz-weighted profiles to simulate Raman spectra with finite bandwidth.

3.

Results and Discussion

Raman spectroscopy has unique capabilities as diagnostic tool in biomedical research.27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37 The method is highly specific for slight environmental changes that have importance in the biological activity of molecules—such as changes in pH,55 water environment,56 or molecular interactions.35, 37 The measurement is fast and does not require further labeling or preparation steps. The combination of a Raman spectrometer with a confocal microscope enables targeting of biological molecules in subcellular compartments with a spatial resolution of approx. 0.5μm .33, 62 Living cells can be studied since water is a weak Raman scatterer and the technique is nondestructive. The sensitivity and selectivity of Raman spectroscopy can be further improved if the excitation wavelength is tuned into the electronic absorption of the molecules.30, 33, 63, 64, 65 Resonance Raman spectroscopy is an extremely capable technique and enables a selective enhancement of the Raman signals of chromophores in complex biological environments.33, 63

Furthermore, the application of UV laser excitation wavelengths causes very strong signal enhancements due to the intrinsic enhancement of the scattering effect [I(ω)ω4] and the resonance enhancement of vibrations that are coupled to the electronic transitions.55, 56, 58 Additionally, the Raman signals are spectrally shifted with respect to disturbing fluorescence and it is possible to derive Raman spectra with high contrast within an environment with fluorescing background.58, 66 UV resonance Raman microspectroscopy was applied for a localization of small amounts of antimalarial drugs in plant material.56, 58

3.1.

UV Resonance Raman Spectroscopy

The molecular structure of halofantrine (Fig. 1) shows a chromophoric phenanthrene structure that is highly polarizable and causes strong Raman signals [Fig. 2 ]. These Raman intensities can be enhanced by several magnitudes by exploitation of the described UV resonance effect. Halofantrine shows strong absorption bands in the deep UV at approx. 250nm .67 Therefore, the laser excitation wavelengths λexc=244nm was chosen for the investigation of halofantrine. A strong enhancement of two Raman bands at 1621 and 1590cm1 is seen in the UV resonance Raman spectrum [Fig. 2] in comparison with the nonresonant FT Raman spectrum (λexc=1064nm) [Fig. 2]. These strongly enhanced Raman signals of halofantrine can be used as marker bands to localize halofantrine in a complex biological environment. The intensities in the spectra in Fig. 2 are normalized to the strongest Raman peaks, respectively, for better illustration.

Fig. 2

(a) UV resonance Raman spectrum of halofantrine (λexc=244nm) and (b) FT-Raman spectrum of halofantrine (λexc=1064nm) . The wavenumber values of some prominent bands are given in graph (b). The band at 1621cm1 is very strong in the UV Raman spectrum (a) due to the resonance enhancement. The intensities in both graphs are normalized to the strongest peak in the spectra, respectively.

041516_1_005004jbo2.jpg

3.2.

DFT Calculation

3.2.1.

Raman spectra

To interpret the individual Raman bands in more detail, a thorough normal mode assignment is required. High-performance computing facilities nowadays enable a reliable calculation of vibrational spectra of medium-sized biomolecules.33, 58 The DFT calculation [B3LYP6-311+G(d,p)] of the Raman spectrum of halofantrine is shown in Fig. 3 . The individual Raman bands in the calculated stick spectrum [Fig. 3] were convoluted with line profiles [Fig. 3] for comparison with measured Raman spectra with finite bandwidth [Fig. 3]. One can see that often several normal modes contribute to an individual Raman band. However, the strongest peak in the nonresonant Raman spectrum of halofantrine [Fig. 3] is dominated by one distinct mode at 1347cm1 [Fig. 3]. Several weaker normal modes contribute to the wings of the line profile of this band. Two molecular vibrations at 1621cm1 and weaker at 1629cm1 [Fig. 3] contribute to the Raman peak at 1621cm1 [Fig. 3], which is strongly enhanced in the UV resonance Raman spectrum of halofantrine [Fig. 2]. The overall agreement of the calculated Raman spectrum of halofantrine [Fig. 3] and the measurement [Fig. 3] is very good, while the intensity of the band at 1431cm1 is underestimated in the calculation. DFT calculations with the hybrid functional B3PW91 were performed to address whether any improvement can be achieved (results are shown in Fig. 4 ). The calculation with B3PW91 shows an improvement of the intensity of the Raman band at 1431cm1 ; however, the intensity of the band at 1621cm1 is overestimated. The results of the DFT calculations with double [6-31+G(d,p)] [Figs. 4 and 4] and triple [6-311+G(d,p)] [Figs. 4 and 4] split valence basis sets are very similar and no significant basis size effect was found. The calculation with the model chemistry B3LYP6-311+G(d,p) was chosen for further consideration because of the very good agreement in the wavenumber region 1500to1650cm1 , where certain Raman modes gain strong UV resonance enhancement.

Fig. 3

Comparison of an experimental FT Raman spectrum (λexc=1064nm) of halofantrine (a) alongside a calculated [B3LYP6-311+G(d,p)] Raman profile (b). The individual Raman modes of the convoluted profile (b) are shown in the stick spectrum (c). The wave number values of some prominent bands are given in (a).

041516_1_005004jbo3.jpg

Fig. 4

Comparison of measured (e) and calculated Raman spectra (a–d) of halofantrine with different model chemistries. The convolution of the profiles was performed with FWHM 6 instead of FWHM 10 in Fig. 3. (a) DFT calculation of the Raman spectrum of halofantrine [B3PW916-311+G(d,p)] , (b) DFT calculation of the Raman spectrum of halofantrine [B3PW916-31+G(d,p)] , (c) DFT calculation of the Raman spectrum of halofantrine [B3LYP6-311+G(d,p)] , (d) DFT calculation of the Raman spectrum of halofantrine [B3LYP6-31+G(d,p)] , (e) Measured FT Raman spectrum of halofantrine (λexc=1064nm) .

041516_1_005004jbo4.jpg

3.2.2.

Electron density distribution

The electron density distribution of halofantrine was calculated and yields fundamental insight regarding the possible binding behavior of the halofantrine molecule. A ππ stacking of the phenanthrene ring of halofantrine with the porphyrin backbone of ferriprotoporphyrin IX (FPPIX) was found20, 21 in a recent crystal structure of an in vitro complex of halofantrine with FPPIX. One should consider whether such binding is possible, due to strongly electron withdrawing substituent’s at the phenanthrene ring of halofantrine. The electron density distribution of halofantrine (Fig. 5 ) highlights the electron withdrawing effect of the substituent’s [C(1)Cl(31) and C(3)Cl(32) as well as C(27)F(28)F(29)F(30)]. However, a strong and evenly distributed electron density remains across the phenanthrene ring. This result supports the hypothesis that ππ stacking may play an important role in the biological activity of halofantrine.

Fig. 5

Calculation of the electron density distribution of halofantrine (iso-value 0.25 electronsa03 ). An even electron distribution is seen in the phenanthrene ring of halofantrine, despite the electron withdrawing effect of substituents C(1)Cl(31) and C(3)Cl(32) as well as C(27)F(28)F(29)F(30).

041516_1_005004jbo5.jpg

3.3.

Mode Assignment and Atomic Displacements

The ultimate goal of the Raman spectroscopic investigation of halofantrine is a localization of the drug at the binding side of the biological target in life cells and an elucidation of changes in the Raman spectrum, caused by molecular interactions. To enable a discussion of such changes in the Raman bands of halofantrine, caused by changes in the environment, a detailed interpretation of the individual Raman bands is required. The DFT calculations can be used for a thorough band assignment. The calculations of the atomic displacements of the molecular normal modes provide much deeper understanding of the associated Raman bands.

The Raman spectrum of halofantrine consists of two well separated parts above 2800cm1 and below 1800cm1 (see Fig. 6 ). The range with high wave number values consist of two separated parts: a region with CH-stretching vibrations at the phenanthrene ring (3120to3060cm1) and an area with CH2 , CH3 stretching vibrations at the side chain (3010to2840cm1) of halofantrine. In addition, there is also one OH stretching vibration present at 3710cm1 . However, the fingerprint region of halofantrine (below 1800cm1 ) is of more importance in the further discussion. The graphical illustration of the atomic displacements of two prominent modes at 1621 and 1349cm1 are shown in Figs. 7 and 7 , respectively. The assignment of more Raman bands of halofantrine is summarized in Table 1 .

Fig. 6

Comparison of measured and calculated [B3LYP6-311+G(d,p)] vibrational spectra of halofantrine. Two scaling factors were applied: 0.98 for wave numbers smaller than 1800cm1 and 0.96 for those above. (a) DFT calculation of the IR stick spectrum of halofantrine, (b) DFT calculation and convolution of the IR spectrum of halofantrine, (c) measured FT IR spectrum of halofantrine, (d) measured FT Raman spectrum of halofantrine, (e) DFT calculation and convolution of the Raman spectrum of halofantrine, and (f) DFT calculation of the Raman stick spectrum of halofantrine. The IR and Raman spectra of halofantrine are complementary. Distinct normal modes show either strong IR or Raman activity, respectively. The wavenumber values of some prominent IR/Raman bands are given in (c) and (d), respectively.

041516_1_005004jbo6.jpg

Fig. 7

(a) Calculated [B3LYP6-311+G(d,p)] atomic displacements of the prominent normal mode of halofantrine at 1621cm1 . This Raman band is strongly enhanced with UV excitation λexc=244nm [see Fig. 2]. The normal mode is localized at the phenanthrene ring with very strong contributions from P1 (see Fig. 1) as described in the context. (b) Calculated [B3LYP6-311+G(d,p)] atomic displacements of the prominent normal mode of halofantrine at 1349cm1 . This Raman band is very strong in the FT-Raman spectrum λexc=1064nm [see Fig. 2]. The normal mode represents a highly symmetric stretching/narrowing vibration, localized at the phenanthrene ring, as described in the text.

041516_1_005004jbo7.jpg

Table 1

Mode assignment and wave number values of prominent Raman bands of halofantrine [Figs. 2, 3, 6].

ν̃(cm−1) Assignment
3120–3060Localized νCH at phenanthrene ring
3010–2840 νCH2 , νCH3 at side chain
1621Very strong CC stretching vibration at the phenanthrene ring; strongest contribution by νC(5)C(6) in P2 as well as C(6)C(15) at the connection of the side chain and δipC(5)H ; highly symmetric vibration across the phenanthrene ring: νC(1)C(14) , νC(3)C(4) , νC(13)C(12) , νC(5)C(6) , νC(11)C(10) , and νC(7)C(8) stretch in phase, with a parallel, out-of-phase movement of the atom pairs {C(4), C(7)} and {C(12)C(9)} toward each other, respectively; further bending: δipC(14)H , δipC(11)H and weaker δipC(9)H and δipC(8)H ; F(28), F(29), F(30), Cl(31), and Cl(32) are fixed in space; no contribution from the side chain
1590Strong CC stretching vibration at the phenanthrene ring; dominated by in-phase C(2)C(3) and C(14)C(13) stretching in P1, C(1)C(14) and C(3)C(4) stretch out-of-phase, C(31) and C(32) are very rigid and distort the symmetry of the mode; strong νCC in P2 and weaker in P3; further very strong δipC(2)H and δipC(14)H and weaker δipC(11)H , δipC(9)H , δipC(8)H , δipC(5)H , and δipC(15)H ; no further contribution from the side chain
1565Highly symmetric, accordion-like CC stretching vibration at the phenanthrene ring; C(1)C(2) , C(13)C(4) , and C(7)C(6) stretch in phase; the vibration is more disturbed in P3 due to the rigid position of C(27)F3 ; medium δipCH around the phenanthrene ring; no further contribution from the side chain
1525Strong CC stretching vibration in P3 of the phenanthrene ring with very strong δipC(11)H , δipC(9)H , and δipC(8)H ; also accordion-like CC stretching vibration in P1 and P2 the phenanthrene ring similar to the one at 1565cm1 but with less intensity; no further contribution from the side chain
1431Tentative assignment: strong CC stretching vibration as well as δipCH around the phenanthrene ring; δO(33)H , δC(15)H and δSC(16)H2 ; no further contribution from the side chain
1383Strong CC stretching vibration as well as δipCH around the phenanthrene ring; δO(33)H , δC(15)H and δSC(16)H2 ; no further contribution from the side chain
1349Very strong combined CC-stretching vibration at the phenanthrene ring; in-phase narrowing/breathing vibration (C(4)C(13), C(7)C(12)) of P1, P2, and P3 (combined with a deformation mode, see Fig. 7); νC(10)C(27) ; νC(27)F3 ; very strong δipC(9)H , δipC(8)H , δipC(11)H , δipC(14)H , medium δipC(2)H , δipC(5)H , δipC(15)H ; small ωCH2 contribution from the side chain
1279This normal mode is spread across the whole molecule; strong combined CC- stretching/breathing vibration at the phenanthrene ring; very strong δipC(11)H ; very strong νC(10)C(17) and δSC(27)F3 (while the F atoms are rigid); medium δipC(2)H , δipC(14)H , δipC(5)H , δipC(8)H , δipC(9)H ; medium ωCH2 , ωCH3 across the side chain
902This normal mode is delocalized across the whole molecule; strong asymmetric breathing vibrations at P1, P2, and P3 of the phenanthrene ring; medium δipC(2)H , δipC(14)H , δipC(5)H , δipC(8)H , δipC(9)H ; δipC(11)H ; strong ρCH2 , ρCH3 across the side chain
757Strong, highly symmetric breathing vibrations at P1, P2, and P3 of the phenanthrene ring; strong νC(1)Cl(31) , νC(10)C(27) , δSC(27)F3 ; δipC(15)H ; ρC(16)H2 , ρC(17)H2
The assignment was performed with help of DFT calculation [B3LYP∕6–311+G(d,p)] . The atomic numbering scheme, as shown in Fig. 1, is used for the assignment. The atomic displacements of the prominent mode at 1622cm−1 , which is strongly enhanced with UV excitation λexc=244nm [Fig. 2], is shown in Fig. 5. The atomic displacements of the strongest mode in the non-resonant Raman spectrum [Fig. 2] at 1349cm−1 is shown in Fig. 7.

The strongest peak in the UV resonance Raman spectrum (λexc=244nm) of halofantrine (Fig. 1) is found at wavenumber position 1621cm1 [Fig. 2]. This band is assigned to a very strong, combined CC -stretching vibration at the phenanthrene ring. The atomic displacement of the mode is displayed in Fig. 7. The strongest contribution to this normal mode arises from νC(5)C(6) in P2 (see Fig. 1 for numbering scheme) as well as from νC(6)C(15) at the connection of the side chain of the molecule and δipC(5)H . A highly symmetric vibration is seen across the phenanthrene ring, where νC(1)C(14) , νC(3)C(4) , νC(13)C(12) , νC(5)C(6) , νC(11)C(10) , and νC(7)C(8) stretch in phase, with a parallel, out-of-phase movement of the pairs {C(4), C(7)} and {C(12), C(9)} toward each other, respectively. Further bending contributions take place δipC(14)H , δipC(11)H and weaker δipC(9)H and δipC(8)H . The fluorine atoms F(28), F(29), and F(30) and chlorine atoms Cl(31) and Cl(32) are fixed in space. There are no contributions from the side chain. The very strong enhancement of this mode in the UV resonance Raman spectrum [Fig. 2] is very promising, since this combined CC -stretching vibration is likely to be influenced by ππ stacking of the phenanthrene ring to the porphyrin ring of biological target structures.20, 21 The calculation of the electron density distribution of halofantrine supports the hypothesis of such interaction. The monitoring of changes in this Raman band during the drug-target interaction might yield novel insight into proposed biological activities of halofantrine.

The strongest band in the nonresonant Raman spectrum is seen at wavenumber position 1349cm1 [Fig. 2]. Two different snapshots of the atomic displacement of this mode are illustrated in Fig. 7. The normal mode consists of a very strong, combined CC-stretching vibration at the phenanthrene ring, an in-phase breathing vibration at [C(4)C(13), C(7)C(12)] of P1, P2, and P3; combined with a deformation mode [see Fig. 7]. The symmetric narrowing of the ring system at C(4)C(13) and C(7)C(12) is nicely illustrated in the two snapshots of the molecular vibration in Fig. 7. More contributions to this mode arise from νC(10)C(27) and νC(27)F3 as well as very strong δipC(9)H , δipC(8)H , δipC(11)H , and δipC(14)H and medium δipC(2)H , δipC(5)H , and δipC(15)H vibrations. Small ωCH2 contributions are present at the side chain.

3.4.

IR Spectroscopy

The IR and Raman spectra of halofantrine yield complementary information. Highly symmetric CC -stretching vibrations in the phenanthrene ring [e.g., Figs. 7 and 7] posses strong changes in polarizability and cause intense bands in the Raman spectrum (Figs. 2 and 3). On the other hand, polar functional groups in the halofantrine molecule cause weak Raman signals, but very strong IR bands. All IR and Raman bands of halofantrine are shown in Fig. 6. The wave number values of some prominent IR and Raman bands are given in the Figs. 6 and 6, respectively. The strongest IR bands of halofantrine are seen at wave number values 2961, 1320, 1147, 1126, 1082, and 1049cm1 . The atomic displacement of the very strong IR mode at 2961cm1 is exemplarily shown in Fig. 8 . This mode represents a combined, asymmetric CH2CH3 -stretching vibration in the butyl part of the side chain of halofantrine.

Fig. 8

Calculated [B3LYP6-311+G(d,p)] atomic displacements of the prominent normal mode of halofantrine at 2961cm1 . This mode is very strong in the IR spectrum [Figs. 3, 3, 3] and represents a combined CH2CH3 -stretching vibration (polar functional group) in the side chain of halofantrine, as described in the text.

041516_1_005004jbo8.jpg

4.

Conclusion and Outlook

The vibrational spectra of halofantrine, an active agent against multidrug-resistant strains of the malaria parasite Plasmodium, were analyzed by means of nonresonant Raman, UV resonance Raman, and IR spectroscopy as well as DFT calculations. The comparison of the nonresonant Raman spectra and UV resonance Raman spectra (Fig. 2) showed that distinct Raman bands at 1621 and 1590cm1 can be enhanced with excitation wavelength λexc=244nm . UV resonance Raman spectra are many orders of magnitude stronger than conventional Raman spectra due to the ω4 dependency of the scattering effect and the selective resonance enhancement. The discovery of these signal enhancements is the basis for continuing experiments that will address a localization of small concentrations of halofantrine in a biological environment. Raman microspectroscopy has unique capabilities to derive molecular information with submicrometer spatial resolution in life cells. It was possible to identify strong, combined CC -stretching vibrations in the phenanthrene ring of halofantrine by means of a thorough mode assignment of the Raman spectra of halofantrine. These vibrations are considered as putative marker bands for ππ interactions with the porphyrin ring of biological target structures. The calculation of the electron density indicates a strong and even distribution across the phenanthrene ring, despite the electron withdrawing effect of the Cl and CF3 substituents. The electron density calculation supports the hypothesis that ππ interactions play an important role in the molecular mode of action of halofantrine.

Further experiments are underway to monitor changes of the putative Raman marker bands during the drug-target binding process. These results will shed more light on suggested models for the biological activity of halofantrine.20, 21 A better elucidation of the molecular mode of action of halofantrine will assist in structure-based design of new, effective antimalarials.

Acknowledgments

The authors gratefully acknowledge that computations were run at the computing center of the University Leipzig. Halofantrine was a kind gift of GlaxoSmithKline.

References

1. 

R. W. Snow, C. A. Guerra, A. M. Noor, H. Y. Myint, and S. I. Hay, “The global distribution of clinical episodes of Plasmodium falciparum malaria,” Nature, 434 (7030), 214 –217 (2005). https://doi.org/10.1038/nature03342 0028-0836 Google Scholar

2. 

, “Key facts,” Rollback Malaria, http://rbm.who.int Google Scholar

3. 

B. Witkowski, A. Berry, and F. Benoit-Vical, “Resistance to antimalarial compounds: Methods and applications,” Drug Resist Update, 12 42 –50 (2009). https://doi.org/10.1016/j.drup.2009.01.001 Google Scholar

4. 

R. G. Ridley, “Medical need, scientific opportunity and the drive for antimalarial drugs,” Nature, 415 686 –693 (2002). https://doi.org/10.1038/415686a 0028-0836 Google Scholar

5. 

I. M. Hastings, P. G. Bray, and S. A. Ward, “A requiem for chloroquine,” Science, 298 74 –74 (2002). https://doi.org/10.1126/science.1077573 0036-8075 Google Scholar

6. 

T. E. Wellems, “Plasmodium chloroquine resistance and the search for a replacement antimalarial drug,” Science, 298 124 –126 (2002). https://doi.org/10.1126/science.1078167 0036-8075 Google Scholar

7. 

T. Weinke, H. D. Notdurft, H. Kretschmer, K. Fleischer, T. Loscher, B. Braendli, K. Markwalder, T. Schlunk, R. Clemens, and H. L. Bock, “Halofantrine in the treatment of imported malaria in non-immune travellers,” Dtsch. Med. Wochenschr., 118 254 –259 (1993). https://doi.org/10.1055/s-2008-1059324 0012-0472 Google Scholar

8. 

J. K. Baird, H. Basri, B. Subianto, D. J. Fryauff, P. D. McElroy, B. Leksana, T. L. Richie, S. Masbar, F. S. Wignall, and S. L. Hoffman, “Treatment of chloroquine-resistant Plasmodium vivax with chloroquine and primaquine or halofantrine,” J. Infect. Dis., 171 1678 –1682 (1995). 0022-1899 Google Scholar

9. 

W. T. Colwell, V. Brown, P. Christie, J. Lange, C. Reece, and K. Yamamoto, “Antimalarial arylaminopropanols,” J. Med. Chem., 15 771 –775 (1972). https://doi.org/10.1021/jm00277a018 0022-2623 Google Scholar

10. 

P. Minodier, G. Noel, M. Salles, K. Retornaz, H. Walters, J. C. Combes, and J. M. Garnier, “Mefloquine versus halofantrine in children suffering from acute uncomplicated falciparum malaria,” Arch. Pediatr., 12 S67 –71 (2005). https://doi.org/10.1016/S0929-693X(05)80017-X 0929-693X Google Scholar

11. 

N. Singh and S. K. Puri, “Modulation of halofantrine resistance after coadministration of halofantrine with diverse pharmacological agents in a rodent malaria model,” Life Sci., 67 1345 –1354 (2000). https://doi.org/10.1016/S0024-3205(00)00728-1 0024-3205 Google Scholar

12. 

Y. T. Kolade, C. P. Babalola, and G. K. E. Scriba, “Analysis of the antimalarial drug halofantrine and its major metabolite N-desbutylhalofantrine in human plasma by high performance liquid chromatography,” J. Pharm. Biomed. Anal., 41 315 –319 (2006). https://doi.org/10.1016/j.jpba.2005.11.002 0731-7085 Google Scholar

13. 

M. Foley and L. Tilley, “Quinoline antimalarials: mechanisms of action and resistance and prospects for new agents,” Pharmacol. Ther., 79 (1), 55 –87 (1998). https://doi.org/10.1016/S0163-7258(98)00012-6 0163-7258 Google Scholar

14. 

S. E. Francis, D. J. Sullivan Jr., D. E. Goldberg, “Hemoglobin metabolism in the malaria parasite Plasmodium falciparum,” Annu. Rev. Microbiol., 51 97 –123 (1997). https://doi.org/10.1146/annurev.micro.51.1.97 0066-4227 Google Scholar

15. 

L. M. B. Ursos and P. D. Roepe, “Chloroquine resistance in the malaria parasite Plasmodium falciparum,” Med. Res. Rev., 22 (5), 465 –491 (2002). https://doi.org/10.1002/med.10016 0198-6325 Google Scholar

16. 

J. Wiesner, R. Ortmann, H. Jomaa, and M. Schlitzer, “Neue Antimalaria-Wirkstoffe,” Angew. Chem., 115 5432 –5451 (2003). https://doi.org/10.1002/ange.200200569 0044-8249 Google Scholar

17. 

H. Matile, R. G. Ridley, and D. E. Goldberg, “A common mechanism for blockade of heme polymerization by antimalarial quinolines,” J. Biol. Chem., 273 31103 –31107 (1998). https://doi.org/10.1074/jbc.273.47.31103 0021-9258 Google Scholar

18. 

I. Y. Gluzman, D. G. Russell, and D. E. Goldberg, “On the molecular mechanism of chlorquine's antimalarial action,” Proc. Natl. Acad. Sci. U.S.A., 93 11865 –11870 (1996). https://doi.org/10.1073/pnas.93.21.11865 0027-8424 Google Scholar

19. 

M. B. Reed, K. J. Sallba, S. R. Carusana, K. Kirk, and A. F. Cowman, “Pgh1 modulates sensitivity and resistance to multiple antimalarials in Plasmodium falciparum,” Nature, 403 906 –909 (2000). https://doi.org/10.1038/35002615 0028-0836 Google Scholar

20. 

K. A. de Villiers, H. M. Marques, and T. J. Egan, “The crystal structure of halofantrine-ferriprotoporphyrin IX and the mechanism of action of arylmethanol antimalarials,” J. Inorg. Biochem., 102 1660 –1667 (2008). https://doi.org/10.1016/j.jinorgbio.2008.04.001 0162-0134 Google Scholar

21. 

K. A. de Villiers and T. J. Egan, “Recent advances in the discovery of Haem-targeting drugs for malaria and schistosomiasis,” Molecules, 14 2868 –2887 (2009). https://doi.org/10.3390/molecules14082868 1420-3049 Google Scholar

22. 

S. Pagola, P. W. Stephens, D. S. Bohle, A. D. Kosar, and S. K. Madsen, “The structure of malaria pagment β-haematin,” Nature, 404 307 –310 (2000). https://doi.org/10.1038/35005132 0028-0836 Google Scholar

23. 

R. Buller, M. L. Peterson, Ö. Almarsson, and L. Leisirowitz, “Quinoline binding site on malaria pigment crystal: a rational pathway for antimalaria drug design,” Cryst. Growth Des., 2 553 –562 (2002). https://doi.org/10.1021/cg025550i 1528-7483 Google Scholar

24. 

B. N. Acharya and M. P. Kaushik, “Pharmacophore-based predictive model generation for potent antimalarials targeting haem detoxification pathway,” Med. Chem. Res., 16 213 –229 (2007). https://doi.org/10.1007/s00044-007-9025-8 Google Scholar

25. 

A. Amponsaa-Karikari, N. Kishikawa, K. Ohyama, K. Nakashima, and N. Kuroda, “Determination if halofantrine and its main metabolite desbutylhalofantrine in rat plasma by high-performance liquid chromatography with on-line UV irradiation and peroxyoxalate chemilumuinescence detection,” Biomed. Chromatogr., 23 101 –106 (2009). https://doi.org/10.1002/bmc.1094 0269-3879 Google Scholar

26. 

Y. T. Kolade, C. P. Babalola, A. A. Olaniyi, and G. K. E. Scriba, “Effect of kolanut on the pharmacokinetics of the antimalarial drug halofantrine,” Eur. J. Clin. Pharmacol., 64 77 –81 (2008). https://doi.org/10.1007/s00228-007-0387-0 0031-6970 Google Scholar

27. 

D. Graham and V. Deckert, “Editorial–a light diagnosis,” Analyst, 134 1027 –1028 (2009). https://doi.org/10.1039/b907715b 0003-2654 Google Scholar

28. 

C. Krafft, B. Dietzek, and J. Popp, “Raman and CARS microspectroscopy of cells and tissues,” Analyst, 134 1046 –1057 (2009). https://doi.org/10.1039/b822354h 0003-2654 Google Scholar

29. 

C. Kendall, M. Isabelle, F. Bazant-Hegemark, J. Hutchings, L. Orr, J. Babrah, R. Baker, and N. Stone, “Vibrational spectroscopy: a clinical tool for cancer diagnostics,” Analyst, 134 1029 –1045 (2009). https://doi.org/10.1039/b822130h 0003-2654 Google Scholar

30. 

T. Frosch, S. Koncarevic, K. Becker, and J. Popp, “Morphology-sensitive Raman modes of the malaria pigment hemozoin,” Analyst, 134 1126 –1132 (2009). https://doi.org/10.1039/b821705j 0003-2654 Google Scholar

31. 

Vibrational Spectroscopy for Medical Diagnosis, Wiley, New York (2008). Google Scholar

32. 

P. Lasch and J. Kneipp, Biomedical Vibrational Spectroscopy, Wiley-VCH, Hoboken, NJ (2008). Google Scholar

33. 

T. Frosch, S. Koncarevic, L. Zedler, M. Schmitt, K. Schenzel, K. Becker, and J. Popp, “In situ localization and structural analysis of the malaria pigment hemozoin,” J. Phys. Chem. B, 1 (37), 11047 –11056 (2007). https://doi.org/10.1021/jp071788b 1089-5647 Google Scholar

34. 

L. Puskar, R. Tuckermann, T. Frosch, J. Popp, V. Ly, D. McNaughton, and B. Wood, “Raman acoustic levitation spectroscopy of red blood cell and Plasmodium falciparum trophozoites,” Lab Chip, 7 1125 –1131 (2007). https://doi.org/10.1039/b706997a 1473-0197 Google Scholar

35. 

T. Frosch, B. Küstner, S. Schlücker, A. Szeghalmi, M. Schmitt, W. Kiefer, and J. Popp, “In vitro polarization-resolved resonance Raman studies of the interaction of hematin with the antimalarial drug chloroquine,” J. Raman Spectrosc., 35 819 –821 (2004). https://doi.org/10.1002/jrs.1252 0377-0486 Google Scholar

36. 

C. Krafft, P. Roesch, and J. Popp, Handbook of Molecular Biophysics, Wiley-VCH, Hoboken, NJ (2009). Google Scholar

37. 

T. Frosch, T. Meyer, M. Schmitt, and J. Popp, “Device for Raman difference spectroscopy,” Anal. Chem., 79 6159 –6166 (2007). https://doi.org/10.1021/ac070440+ 0003-2700 Google Scholar

38. 

W. Kiefer, “Raman difference spectroscopy with the rotating cell,” Appl. Spectrosc., 27 253 (1973). https://doi.org/10.1366/000370273774333452 0003-7028 Google Scholar

39. 

M. J. Frisch, G. W. Trucks, H. B. Schlegel, G. E. Scuseria, M. A. Robb, J. R. Cheeseman, J. A. Montgomery Jr., T. Vreven, K. N. Kudin, J. C. Burant, J. M. Millam, S. S. Iyengar, J. Tomasi, V. Barone, B. Mennucci, M. Cossi, G. Scalmani, N. Rega, G. A. Petersson, H. Nakatsuji, M. Hada, M. Ehara, K. Toyota, R. Fukuda, J. Hasegawa, M. Ishida, T. Nakajima, Y. Honda, O. Kitao, H. Nakai, M. Klene, X. Li, J. E. Knox, H. P. Hratchian, J. B. Cross, V. Bakken, C. Adamo, J. Jaramillo, R. Gomperts, R. E. Stratmann, O. Yazyev, A. J. Austin, R. Cammi, C. Pomelli, J. W. Ochterski, P. Y. Ayala, K. Morokuma, G. A. Voth, P. Salvador, J. J. Dannenberg, V. G. Zakrzewski, S. Dapprich, A. D. Daniels, M. C. Strain, O. Farkas, D. K. Malick, A. D. Rabuck, K. Raghavachari, J. B. Foresman, J. V. Ortiz, Q. Cui, A. G. Baboul, S. Clifford, J. Cioslowski, B. B. Stefanov, G. Liu, A. Liashenko, P. Piskorz, I. Komaromi, R. L. Martin, D. J. Fox, T. Keith, M. A. Al-Laham, C. Y. Peng, A. Nanayakkara, M. Challacombe, P. M. W. Gill, B. Johnson, W. Chen, M. W. Wong, C. Gonzalez, and J. A. Pople, Gaussian 03, Revision D.01, Gaussian, Inc., Wallingford, CT (2004). Google Scholar

40. 

A. D. Becke, “Density-functional thermochemistry. II. The effect of the Perdew-Wang generalized-gradient correlation correction,” J. Chem. Phys., 97 9173 –9177 (1992). https://doi.org/10.1063/1.463343 0021-9606 Google Scholar A. D. Becke, “Density-functional thermochemistry. III. The role of exact exchange,” J. Chem. Phys., 98 5648 –5652 (1993). https://doi.org/10.1063/1.464913 0021-9606 Google Scholar

41. 

P. J. Stephens, F. J. Devlin, C. F. Chabalowski, and M. J. Frisch, “Ab initio calculation of vibrational absorption and circular dichroism spectra using density functional force fields,” J. Phys. Chem., 98 11623 (1994). https://doi.org/10.1021/j100096a001 0022-3654 Google Scholar

42. 

C. Lee, W. Yang, and R. G. Parr, “Development of the Colle-Salvetti correlation-energy formula into a function of the electron density,” Phys. Rev. B, 37 785 (1998). https://doi.org/10.1103/PhysRevB.37.785 0163-1829 Google Scholar

43. 

J. P. Perdew and Y. Wang, “Accurate and simple analytic representation of the electron-gas correlation energy,” Phys. Rev. B, 45 13244 (1992). https://doi.org/10.1103/PhysRevB.45.13244 0163-1829 Google Scholar

44. 

J. P. Perdew, J. A. Chevary, S. H. Vosko, K. A. Jackson, M. R. Pederson, D. J. Singh, and C. Fiolhais, “Atoms, molecules, solidas, and surfaces: Applications of the generalized gradient approximation for exchange and correlation,” Phys. Rev. B, 46 6671 (1992). https://doi.org/10.1103/PhysRevB.46.6671 0163-1829 Google Scholar

45. 

J. A. Pople, H. B. Schlegel, R. Krishnan, D. J. Defrees, J. S. Binkley, M. J. Frisch, and R. A. Whitside, “Molecular orbital studies of vibrational frequencies,” Int. J. Quantum Chem., 15 269 –278 (1981). 0020-7608 Google Scholar

46. 

M. J. Frisch, J. A. Pople, and J. S. Binkley, “Self-consistent molecular orbital methods 25. Supplementary functions for Gaussian basis sets,” J. Chem. Phys., 80 (7), 3265 –3269 (1984). https://doi.org/10.1063/1.447079 0021-9606 Google Scholar

47. 

W. J. Hehre, R. F. Stewart, and J. A. Pople, “Self-consistent molecular-orbital methods. I. Use of Gaussian expansions of slater-type atomic orbitals,” J. Chem. Phys., 51 (6), 2657 –2664 (1969). https://doi.org/10.1063/1.1672392 0021-9606 Google Scholar

48. 

A. P. Scott and L. Radom, “Harmonic vibrational frequencies: An evaluation of Hartree-Fock, Møller-Plesset, quadratic configuration interaction, density functional theory, and semiempirical scale factors,” J. Phys. Chem., 100 16502 –16513 (1996). https://doi.org/10.1021/jp960976r 0022-3654 Google Scholar

49. 

A. A. El-Azhary and H. U. Suter, “Comparison between optimized geometries and vibration frequencies calculated by the DFT methods,” J. Phys. Chem., 100 15056 –15063 (1996). https://doi.org/10.1021/jp960618o 0022-3654 Google Scholar

50. 

J. Baker, A. A. Jarzecki, and P. Pulay, “Direct scaling of primitive valence force constants: An alternative approach to scaled quantum mechanical force fields,” J. Phys. Chem. A, 102 1412 –1424 (1998). https://doi.org/10.1021/jp980038m 1089-5639 Google Scholar

51. 

G. Rauhut and P. Pulay, “Transferable scaling factors for density functional derived vibrational force fields,” J. Phys. Chem., 99 3093 –3100 (1995). https://doi.org/10.1021/j100010a019 0022-3654 Google Scholar

52. 

M. D. Halls and B. Schlegel, “Comparison of the performance of local, gradient-corrected, and hybrid density functional models in predicting infrared intensities,” J. Chem. Phys., 109 (24), 10587 –10593 (1998). https://doi.org/10.1063/1.476518 0021-9606 Google Scholar

53. 

M. P. Andersson and P. Uvdal, “New scale factors for harmonic vibrational frequencies using the B3LYP density functional method with the triple-ζ basis set 6–311+G(d,p),” J. Phys. Chem. A, 109 2937 –2941 (2005). https://doi.org/10.1021/jp045733a 1089-5639 Google Scholar

54. 

C. W. Bauschlicher and S. R. Langhoff, “The calculation of accurate harmonic frequencies of large molecules: the polycyclic aromatic hydrocarbons, a case study,” Spectrochim. Acta, Part A, 53 1225 –1240 (1997). https://doi.org/10.1016/S1386-1425(97)00022-X 0584-8539 Google Scholar

55. 

T. Frosch, M. Schmitt, G. Bringmann, W. Kiefer, and J. Popp, “Structural analysis of the anti-malaria active agent chloroquine under physiological conditions,” J. Phys. Chem. B, 111 (7), 1815 (2007). https://doi.org/10.1021/jp065136j 1089-5647 Google Scholar

56. 

T. Frosch, M. Schmitt, and J. Popp, “In situ UV resonance Raman micro-spectroscopic localization of the antimalarial quinine in cinchona bark,” J. Phys. Chem. B, 111 (16), 4171 –4177 (2007). https://doi.org/10.1021/jp066999f 1089-5647 Google Scholar

57. 

T. Frosch, M. Schmitt, and J. Popp, “Raman spectroscopic investigation of the antimalarial agent mefloquine,” Anal. Bioanal. Chem., 387 1749 –1757 (2007). https://doi.org/10.1007/s00216-006-0754-1 1618-2642 Google Scholar

58. 

T. Frosch, M. Schmitt, T. Noll, G. Bringmann, K. Schenzel, and J. Popp, “Ultrasensitive in situ tracing of the alkaloid dioncophylline A in the tropical liana triphyophyllum peltatum by applying deep-UV resonance Raman microscopy,” Anal. Chem., 79 (3), 986 –993 (2007). https://doi.org/10.1021/ac061526q 0003-2700 Google Scholar

59. 

T. Frosch, M. Schmitt, K. Schenzel, J. H. Faber, G. Bringmann, W. Kiefer, and J. Popp, “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 functional theory calculations,” Biopolymers, 82 (4), 295 –300 (2006). https://doi.org/10.1002/bip.20459 0006-3525 Google Scholar

60. 

T. Frosch and J. Popp, “Relationship between molecular structure and Raman spectra of quinolines,” J. Mol. Struct., 924–926 301 –308 (2009). https://doi.org/10.1016/j.molstruc.2008.12.019 0022-2860 Google Scholar

61. 

D. A. Long, The Raman Effect, Wiley, New York (2002). Google Scholar

62. 

G. J. Puppels, F. F. M. De Mul, C. Otto, J. Greve, M. Robert-Nicoud, D. J. Arndt-Jovin, and T. M. Jovin, “Studying single living cells and chromoscomes by confocal Raman microspectroscopy,” Nature, 347 301 –303 (1990). https://doi.org/10.1038/347301a0 0028-0836 Google Scholar

63. 

Biological Applications of Raman Spectroscopy, 1–3 Wiley, New York (1988). Google Scholar

64. 

R. Petry, M. Schmitt, and J. Popp, “Raman spectroscopy: A prospective tool in the life sciences,” ChemPhysChem, 4 14 (2003). https://doi.org/10.1002/cphc.200390004 1439-4235 Google Scholar

65. 

Handbook of Vibrational Spectroscopy. Vol. 5. Applications in Life, Pharmaceutical and Natural Sciences, Wiley & Sons, Chichester, UK (2002). Google Scholar

66. 

T. Frosch, N. Tarcea, M. Schmitt, H. Thiele, F. Langenhorst, and J. Popp, “UV Raman imaging-a promising tool for astrobiology: comparative Raman studies with different excitation wavelengths on SNC martian meteorites,” Anal. Chem., 79 1101 –1108 (2007). https://doi.org/10.1021/ac0618977 0003-2700 Google Scholar

67. 

A. Taillardat-Bertschinger, C. S. Pery, A. Galland, R. J. Prankerd, and W. N. Charman, “Partitioning of halofantrine hydrochloride between water, Micellar solutions, and soybean oil: Effects on its apparent ionization constant,” J. Pharm. Sci., 92 2217 –2228 (2003). https://doi.org/10.1002/jps.10479 0022-3549 Google Scholar
©(2010) Society of Photo-Optical Instrumentation Engineers (SPIE)
Torsten Frosch and Jürgen Popp "Structural analysis of the antimalarial drug halofantrine by means of Raman spectroscopy and density functional theory calculations," Journal of Biomedical Optics 15(4), 041516 (1 July 2010). https://doi.org/10.1117/1.3432656
Published: 1 July 2010
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