Novel cholinesterase modulators and their ability to interact with DNA

Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy 115 (2013) 364–369

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Novel cholinesterase modulators and their ability to interact with DNA Jana Janockova a, Zuzana Gulasova a, Kamil Musilek b,c, Kamil Kuca c, Maria Kozurkova a,⇑ a

Institute of Chemistry, Department of Biochemistry, P.J. Šafárik University, Faculty of Science, Moyzesova 11, 04001 Kosice, Slovak Republic University of Hradec Kralove, Faculty of Science, Department of Chemistry, Rokitanskeho 62, 500 03 Hradec Kralove, Czech Republic c University Hospital, Sokolska 581, 500 05 Hradec Kralove, Czech Republic b

g r a p h i c a l a b s t r a c t

 We studied selected cholinesterase

600

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modulators – oximes with calf thymus DNA.  The binding constants were determinate from titrations (K = 3.5  104 to 1.4  105 M1).  Electrophoretic techniques proved that ligand 2 inhibited topoisomerase I (5 lM).

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Article history: Received 26 February 2013 Received in revised form 23 May 2013 Accepted 4 June 2013 Available online 17 June 2013 Keywords: Cholinesterase modulators DNA Topoisomerase I

a b s t r a c t In the present work, an interaction of four cholinesterase modulators (1–4) with calf thymus DNA was studied via spectroscopic techniques (UV–Vis, fluorescent spectroscopy and circular dichroism). From UV–Vis spectroscopic analysis, the binding constants for DNA-pyridinium oximes complexes were calculated (K = 3.5  104 to 1.4  105 M1). All these measurements indicated that the compounds behave as effective DNA-interacting agents. Electrophoretic techniques proved that ligand 2 inhibited topoisomerase I at a concentration 5 lM. Crown Copyright Ó 2013 Published by Elsevier B.V. All rights reserved.

Introduction The development of compounds which modify nucleic acids structure is an actual research topic and might have a significant contribution to antitumor and antiviral chemotherapy [1,2]. The search for new pharmaceutically active drugs has led to the discovery of many DNA binding drugs [3–6]. Binding of various molecules to DNA regulates its function, protein–DNA interactions form the basis of various molecular–biological processes required for cells physiology [7].

⇑ Corresponding author. E-mail address: [email protected] (M. Kozurkova).

Much effort has also been directed to study DNA-small molecule interactions, such as the interaction with inorganic complexes [5,8,9], fluorescent dyes [10,11] and most importantly organic molecules [12–17]. These studies help to understand the molecular basis of complex diseases and may provide potential drug candidates. Schwartsmann [18] reported that marine organisms are a rich source of chemically novel products with a broad spectrum of bioactivity. Along the biologically active compounds that have been isolated from marine sponges, there are several pyridinium derivatives [19]. Biologically active pyridinium derivatives, so far isolated from marine sponges, exert different biological activities, including cytotoxicity, hemolysis, inhibition of acetylcholinesterase enzyme activity and binding to different receptors, antibacterial, antifungal and insecticidal activity [20]. The ability to inhibit

1386-1425/$ - see front matter Crown Copyright Ó 2013 Published by Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.saa.2013.06.008

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J. Janockova et al. / Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy 115 (2013) 364–369 Table 1 The chemical structure of compounds (1–4) and their chemical characteristics.

MW (g mol1)

Sign

Name

1

Trimedoxime dibromide

443.98

2

(E)-1-(4-carbamoylpyridinium)-4-(4-hydroxyiminomethylpyridinium)but-2-ene dibromide

458.15

3

4,40 -Bis(hydroxyiminomethyl)-1,10 -(1,3-phenylenedimethyl)bispyridinium dibromide

508.21

4

Oct-1,8-diyl-1,8-bis(isoquinolinium) dibromide

530.34

structural factors (e.g., condensed or closely connected aromatic rings). In this paper, the biochemical and biological activities of selected cholinesterase modulators are described and tested as DNA-binding compounds. Their capacity to bind to DNA and to interfere with human topoisomerase I was examined.

Materials and methods Materials Studied pyridinium compounds 1–4 were prepared at the Department of Chemistry, Faculty of Science, University of Hradec Kralove [31–34]. The compounds were dissolved in dimethyl sulfoxide (SERVA) to a concentration of 2.0  104 lM. The chemical characteristics of used compounds are depicted in Table 1. UV–Vis measurements UV–Vis spectra were measured on a Varian Cary 100 UV–Vis spectrophotometer. The compounds 1–4 were all dissolved in DMSO, from which diluted working solutions were prepared using a 0.01 M Tris buffer (pH = 7.4) to concentration 25 lM. The concentration of calf thymus DNA (ctDNA, purchased from Sigma) ranged from 0 to 61.6  102 lM. The DNA concentration was determined 0,16

0,12

Absorbance

acetylcholinesterase activity has been focused by several authors, for its possible regulatory effects on cancer progression. The studies Hyatt et al. [21] indicate that CPT-11 (irinotecan, 7-ethyl-10-[4(1-piperidino)-1-piperidino]carbonyloxycamptothecin) is a potent inhibitor of AChE, and this probably accounts for the cholinergic syndrome that is observed in cancer patients. The cholinesterase modulators are chemical compounds with various applications in the pharmaceutical industry. Some of them are classified as oximes. Oximes possess considerable biological activities [22]. Pyridinium oximes are used mainly as antidotes for poisoning by organophosphorus compounds [23]. Other oximes can also act as agents with positive effect on the cardiovascular system [24] or for their an anti-inflammatory [25], anti-viral [26,27], antibacterial and antifungal activity [28,29]. The cholinesterase modulators have been only little studied as cytotoxic agents. Recently, it was shown that silicon-containing oximes have high cytotoxicity towards HT-1080 (human fibrosarcoma) and MG22A (mouse hepatoma) strains [30]. All presented cholinesterase modulators were previously prepared and evaluated on cholinesterase targets. Compound 1 (trimedoxime) is standard acetylcholinesterase (AChE) reactivator that was and still is used for decades to counteract organophosphorus (OP) intoxication [31]. Compounds 2 and 3 were originally designed as novel acetylcholinesterase reactivators against OP intoxication with main aim to treat nerve agent poisoning [32,33]. Both molecules have at least one oxime (hydroxyiminomethyl) functional group that is essential for their reactivation ability towards OP-inhibited AChE. The compounds 3 was found to be effective reactivator of pesticide inhibited AChE in vitro [32]. However, its reactivation potency was found limited due to its increased toxicity in vivo [34]. On the other hand, the AChE reactivator 2 was found among most potent compounds against tabun intoxication in vitro and only intermediate toxic compound [33,35,36]. Its reactivation results were further confirmed in vivo and it is very promising candidate for next studies related to human use in case of OP intoxication [36,37]. Compound 4 was designed as cholinesterase inhibitor [38]; introduced as peripherally acting with implications to Myasthenia gravis treatment. The in vitro screening confirmed its inhibitory ability on nM scale and the related in vivo studies are in progress [39]. The rational for their DNA interaction screening lie in their mice toxicity that they previously presented in vivo. Thus, the interaction of presented compounds with DNA was supposed due to their

Structure

0,08

0,04

0 320

340

360

380

400

Wavelength [nm] Fig. 1. UV–Vis spectrophotometric titration of derivatives 2 (25 lM) in 0.01 M Trisbuffer (pH = 7.4, 24 °C) with increasing concentration of ctDNA (0–45.8  102 lM).

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ctDNA (31.6  102 lM) or a mixture of ctDNA with 1–4 (50 lM) in BPE buffer, pH 7.1 (6 mM Na2HPO4, 2 mM NaH2PO4, 1 mM EDTA), with a heating rate of 1 °C min1. The melting temperatures were determined as the maximum of the first derivative plots of the melting curves [40].

Table 2 UV–Vis absorption data and DNA binding properties of compounds 1–4. Compound

kmax (nm) Free

Bond

1 2 3 4

343 342 343 335

342 340 340 335

Dk (nm)

Hypochromicity (%)

a

1 2 3 0

11.7 28.8 41.6 4.07

74.0 74.9 74.0 77.9

Tm

K (M1)

1.4  105 3.5  104 9.0  104 6.8  104

Fluorescence measurements The fluorescent measurements of the DNA interaction with selected compounds 1–4 were scanned on a Varian Cary Eclipse spectrofluorimeter. All measurements were made using a 10 mm lightpath cuvette in a 0.01 M Tris buffer at pH 7.4. For excitation and emission beams were used 10 nm slits. Fluorescence intensities are expressed in arbitrary units. The binding of ligands 1–3 to ctDNA was investigated by fluorescent intercalator displacement assay (FID) with ethidium bromide (EtBr) and premixed solution of ctDNA (25.3  103 lM) and increasing amounts of ligand 1 (c1 = 0–1.36  102 lM), 2 (c2 = 0–2.0  102 lM) and 3 (c3 = 0– 2.0  102 lM) in 0.01 M Tris buffer, pH 7.4. Emission spectra were collected from 550 to 800 nm with 510 nm excitation of EtBr at 24 °C [41]. Emission spectrum of compound 4 was recorded in the region 350–480 nm using excitation wavelength 335 nm. Fluorescence

a Tm measurements were performed in BPE buffer, pH 7.1 (6 mM Na2HPO4, 2 mM NaH2PO4, 1 mM EDTA) using compound 1–4 (50 lM) and 31.6  102 lM ctDNA with a heating rate of 2 °C min1. Tm of ctDNA is 72.0 °C.

from its absorbance at 260 nm. The purity of DNA was determined by monitoring a value A260/A280. All measurements were performed at 24 °C [40]. Tm measurements Thermal denaturation studies were conducted using a Varian Cary Eclipse spectrophotometer equipped with a thermostatic cell holder. The temperature was controlled by a thermostatic bath (±0.1 °C). The absorbance at 260 nm was monitored for either

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Wavelength (nm) Fig. 2. (A) Fluorescent intercalator displacement of EtBr bound to DNA (25.3  103 lM) with increasing concentrations of derivative 2 (0–2.0  102 lM), kex = 510 nm, in 10 mM Tris (pH = 7.4, 24 °C), and (B) spectrofluorimetric titration of derivative 4 (4.0 lM) in 0.01 M Tris buffer (pH 7.4, 24 °C) by increasing the concentration of ctDNA (from top to bottom, 0–0.43 lM bp, at 2 lM intervals), kex = 335 nm.

J. Janockova et al. / Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy 115 (2013) 364–369

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10

poly(A-T) (25 µM)

CD [mdeg]

5

poly(A-T) + 1 (4 x 10 2 µM) poly(A-T) + 2 (4 x 10 2 µM)

0

poly(A-T) + 3 (4 x 10 2 µM)

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poly(A-T) + 4 (4 x 10 2 µM)

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Wavelength [nm] Fig. 3. Circular dichroism spectra of poly(A–T) (c = 25 lM) in the absence and presence of compounds 1–4 (c = 4  102 lM) in 0.01 M Tris buffer (pH 7.4, 24 °C).

Fig. 4. Electrophoresis agarose gel showing inhibition of calf thymus topoisomerase I induced DNA relaxation by compounds 1–4. Native supercoiled pBR322 (lane 0) was incubated for 45 min at 37 °C with topoisomerase I in the absence (lane T) or presence of ligands (lane a – 5, lane b – 30, lane c – 60 lM).

Fig. 5. Effect of samples 1–4 on the relaxation of relaxed pBR322 plasmid DNA by topoisomerase I. Relaxed pBR322 (1.4 mg, lane 0) was incubated for 45 min at 37 °C with 3 units of topoisomerase I in the absence (lane T) or presence of ligands (lane a – 5, lane b – 30, lane c – 60 lM). Line CPT – relax pBR322 + Topo I + campothecine (10 lM), line H – relax pBR322 + Topo I + Hoechst (10 lM), line EtBr – relax pBR322 + Topo I + ethidium bromide (5 lM). The DNA samples were run on agarose gel followed by ethidium bromide staining.

titrations were carried out by adding increasing amounts of DNA (0–0.43 lM) directly into the cuvette containing solution of analogue 4 (c = 4 lM) in 0.01 M Tris buffer, pH 7.4 at 24 °C [42].

quires a determination of Cf and, hence, the amount r of drug bound per unit of DNA as a function of added titrant. For determination Cf and r we used equitation:

Circular dichroism

a ¼ C b =C ¼ ð1  C f =CÞ ¼ ðA0f  AÞ=ðA0f  A0b Þ

Circular dichroism spectra were recorded on a J-810 Jasco spectropolarimeter in a rectangular quartz cell of 1 mm path length. All measurements were performed in 0.01 M Tris buffer (pH = 7.4) at 24 °C, the concentration of DNA was 3.16 lM and the concentration of studied derivatives was 4  102 lM. CD spectra were measured from 230 to 400 nm. Results are presented as a mean of three scans [42]. Equlibrium binding titration

where a is the drug binding fraction, A0f and A0b are the measured absorption for the free and fully bound drug at the monitoring wavelength. Then, r = aC/CDNA and Cf = (1  a)C, where CDNA is the total concentration of DNA [43]. Data from UV–Vis measurements were used to determine the binding constants K of 1–4 with DNA. The binding data were applied to McGhee and von Hippel equation [44] to identify the value of K:

This technique has found universal application in DNA–drug binding studies. The absorbance A measured at any wavelength reflects both the free and DNA-bound species:

 n1 r 1  nr ¼ Kð1  nrÞ Cf 1  ðn  1Þr

A ¼ Af þ Ab ¼ ef  C f þ eb  C b where C is the fixed drug concentration (i.e., Cf + Cb) and ef and eb represent the respective extinction coefficient. Binding analysis re-

where K is a binding constant, n is the binding site size in the base pairs. The binding data were fitted using GNU Octave 2.1.73 software [45].

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Topoisomerase relaxation assay Evaluation of topoisomerase activity was performed via gel electrophoresis assay. In these experiments, calf thymus topoisomerase I (Takara, Japan) and negatively supercoiled or relaxed plasmid pBR322 DNA (1.4 lg) as the substrate in the reaction buffer (350 mM Tris–HCl, pH = 8, 720 mM KCl, 50 mM MgCl2, 50 mM DTT, 50 mM spermidine; 20 lL) containing 0.1 % bovine serum albumin (BSA) were used. Appropriate inhibitor with increasing concentration (5 lM, 30 lM, 60 lM) was added and the reaction was initiated by the addition of 3 units of topoisomerase I. The reactions were carried out at 37 °C for 45 min. A gel electrophoresis was performed at 7 V/cm for 2 h in TBE (Tris + boric acid + EDTA) buffer on a 1% agarose gel. The gel was stained with ethidium bromide (1 mg/ml) and photographed under UV light [42].

Results and discussion Absorption, circular dichroism and fluorescence give different type of information and thus they are used together to provide a powerful toolbox for investigation of molecular structures including the different modes of DNA binding. UV–Vis absorption spectra showed significant absorption of samples 1–4 in the range 300–400 nm. Fig. 1 provides an example that illustrates characteristic changes in the absorption spectra during titration of compound 2 with increasing amounts of ctDNA. The binding of drugs 1–4 to DNA resulted in different spectral changes characterized by shift and hypochromism (Table 2). UV– Vis titration data have been used to determine binding constant of ligands 1–4 with ctDNA using McGhee and von Hippel plots. Binding parameters from spectrophotometric analysis are summarized in Table 2. The values of binding constants K, (in the range 3.5  104 to 1.4  105 M1) were indicative of high binding affinity (compared to amsacrine is 4.0  104 M1) [46]. Additional evidence for interaction of sample 1–4 with DNA was obtained from thermal denaturation studies. Binding of small molecules to DNA is assumed to stabilize the helix against its thermal denaturation and therefore it is accompaniment by the rise of translation temperature (Tm) for the double- to single-stranded DNA chain. As it is shown in Table 2, the DNA melting experiment revealed that Tm of calf thymus DNA was 72.0 °C and it increased from 74.0 to 77.9 °C in the presence of compounds 1–4. Thus, confirming heightened helix stability as the results of intercalation studied derivatives into DNA [40,47]. The binding of compounds 1–4 to DNA has been characterized through fluorescence spectroscopy. Fig. 2 shows the fluorescence emission spectra of compounds 2 in the absence and presence of ctDNA. Because of low fluorescence, the binding of ligands 1–3 to ctDNA was investigated by FID assay methods with ethidium bromide (EtBr) and premixed solution of ctDNA. The compounds 1–3 have an emission maximum at around 600 nm by 510 nm excitation of EtBr. This also indicated that compounds 1–3 also bound to DNA through an intercalation mode similar as the EtBr did. Ligand 4 had an emission maximum at 375 nm, when it was excited at 335 nm. The fluorescence intensity increased with the addition of ctDNA, where the wavelength shift was not observed. The stronger enhancement of fluorescence emission spectra may be caused by the planar groups of molecules on the booth sides of molecules that can stack between adjacent base pairs of ctDNA. The fluorescence quenching curve is given in the inset of Fig. 2. Among the methods available for analysis of structures of drug-DNA complexes in solution, the circular dichroism (CD) is particularly simple, and provides information, which cannot be obtained as conveniently by the other methods. One of the advantages of the method is the fact that some important information

may already be derived without using any complex theory [48,49]. CD spectra monitor the asymmetric environment of the compounds when they are bound to DNA and therefore they can be used to obtain information on their binding mode. The free compounds do not have CD spectra, but they might induce CD, when they bound to DNA. The CD spectrum of the DNA is sensitive to changes in conformation resulting from ligand binding, but an interpretation of the CD signal is complex. It has been shown (Fig. 3) that the CD spectrum of poly(A–T) is significantly altered in the presence of ligands 1–4. Changes of negative bands at 245 nm with a non-significant red shift were observed for all tested compounds. The positive band at 279 nm showed a decrease of molar elipticity, and mild red shift of the band maxima along with an increase of intensity upon addition of the compounds 1–4 to poly(A–T). The CD spectrum of free ctDNA has a negative band at 245 nm due to helicity, and a positive band at 279 nm due to base stacking which is characteristic of DNA in the right-handed B form. Kozˇurková et al. [47] referred that proflavine compounds incubated with ctDNA, displayed changes in both positive and negative CD bands. When the positive band at 279 nm showed an increase of molar elipticity following the addition of derivatives to the DNA, this phenomenon could be due to the stabilization of a right-handed B form of DNA by intercalation. In our case changes in elipticity prove that these compounds not intercalate into DNA, but interact with it in a different mode. DNA-topoisomerases (Topo I), a family of DNA-processing enzymes, represent the pharmacological target of major clinically useful chemotherapeutic agents. Topoisomerase-targeted drugs can be divided into two broad classes that vary widely in their mechanisms of action [50–51]. The class I include the drugs that act by stabilizing covalent topoisomerase-DNA complexes. They are also referred to as ‘‘topoisomerase poisons’’, because they transform the enzyme into a potent cellular toxin. In contrast, class II drugs interfere with the catalytic function of the enzyme without trapping the covalent complex. The drugs in this class are referred to as ‘‘topoisomerase inhibitors’’ [52,53]. The ligand that occupies the topoisomerase binding site may suppress the association of topoisomerase with DNA, thus it influences the topoisomerase activity. DNA intercalators have the high developmental potential as DNA targeted anticancer drugs [54]. Initially, the topoisomerase I inhibitory properties of the selected compounds were examined using a DNA relaxation assay. Negatively supercoiled plasmid pBR322 was incubated with topoisomerase I at three compound concentrations (5, 30 and 60 lM). The photopicture of 1–4 agarose gel electrophoresis experiment is presented in Fig. 4. As it is shown, supercoiled pBR322 was fully relaxed by the enzyme in the absence of the studied samples (line 0 and T). Strong scDNA bands and very light ocDNA bands can be seen for compounds 1–4. Thus, 1–4 all significantly inhibit Topo I catalysis, when tested at 30 or 60 lM concentration. The gel shows that compound 2 shows Topo I relaxation activity at a very low ligand concentration (5 lM). The better differentiation between the topoisomerase inhibition or DNA intercalation by tested compounds was performed as a DNA unwinding assay using relaxed plasmid DNA (pBR322) and calf thymus topoisomerase I in the presence of three compound concentrations (5, 30 and 60 lM). In the presence of the drugs, the intensity of the band corresponding to the open circular form of DNA had no change. Interestingly at concentration >5 lM, compounds inhibited the relaxation of scDNA by Topo I. The relaxed plasmid did not revert back to its supercoiled form (typical for intercalators) indicating that 1–4 constituents did not inhibit the topoisomerase I activity through intercalation into the DNA (Fig. 5). In order to verify the activities of tested compounds, they were compared with the known Topo I inhibitor Campothecine

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(CPT, 10 lM), poison Hoechst 33258 (H, 10 lM) and intercalator ethidium bromide (EtBr, 5 lM) as controls. Conclusions The interactions between selected cholinesterase modulators and DNA were investigated at physiological pH = 7.4 using UV– Vis fluorescence and circular dichroism spectroscopic techniques. It was shown that modulators 1–4 could bind to DNA with a high affinity. The binding constants were calculated (K = 3.5  104 to 1.4  105 M1). All observations were consistent with a non-intercalation mode for selected compounds binding to DNA. The relaxed plasmid did not revert back to its supercoiled form indicating that samples constituents inhibit the topoisomerase I activity through non-intercalative mode. This work could help to go further in understanding of the binding mechanism of cholinesterase modulators with DNA and possibly designing some similar new and efficient molecules targeted to DNA.

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Acknowledgments This study was supported by Slovak Research and Development Agency (VEGA Grant No. 1/0001/13, APVV-0280-11), Internal Grant Program of the P.J. Šafárik University in Kosice (VVGS 40/ 12-13, VVGS-PF-2012-16), University Hospital in Hradec Kralove (long term development plan) and University Hradec Kralove (long term development plan). References [1] R. Martinez, L. Chacón-Garcia, Curr. Med. Chem. 12 (2005) 1345–1359. [2] H. Ihmels, D. Otto, Top. Curr. Chem. 258 (2005) 161–204. [3] G. Cholewinski, K. Dzierbicka, A.M. Kolodziejcyk, Pharm. Rep. 63 (2011) 305– 336. [4] L.R. Ferguson, W.A. Denny, Mutat. Res. 258 (1991) 123–160. [5] P. Belmont, J. Bosson, T. Godet, M. Tiano, Anticancer Agents Med. Chem. 7 (2007) 139–169. [6] M. Demeunynck, F. Charmantray, A. Martelli, Curr. Pharm. Des. 7 (2001) 1170– 1724. [7] T.C. Rowe, V. Weissig, J.W. Laerence, Adv. Drug Deliv. Rev. 49 (2001) 175–187. [8] H.A. Azab, B.H. Hussein, A.L.I. Al Falouji, J. Fluoresc. 22 (2012) 639–649. [9] D. Qin, Z. Yang, B. Wang, Spectrochim. Acta Part A 68 (2007) 912–917. [10] M. Sarma, T. Chatterjee, S. Ghanta, S.K. Das, J. Org. Chem. 77 (2012) 432–444. [11] A.G. Majouga, A.V. Udina, E.K. Beloglazkina, D.A. Skvortsov, M.I. Zvereva, O.A. Dontsova, N.V. Zyk, N.S. Zefirov, Tetrahedron Lett. 53 (2012) 51–53. [12] Z. Frohlichova, J. Imrich, I. Danihel, P. Kristian, S. Bohm, D. Sabolova, M. Kozurkova, O. Hritzova, B. Horvath, T. Busova, K.D. Klika, Spectrochim. Acta Part A 73 (2009) 238–248. [13] M. Stefanisinova, V. Tomeckova, M. Kozurkova, A. Ostro, M. Marekova, Spectrochim. Acta Part A 81 (2011) 666–671. [14] B.C.C. Souza, T.B. De Oliveira, T.M. Aquino, M.C.A. De Lima, I.R. Pitta, S.L. Galdino, E.O. Lima, T. Gonçalves-Silva, G.C.G. Militão, L. Scotti, M.T. Scotti, F.J.B. Mendonça, Acta Pharm. 62 (2012) 221–236. [15] F.W.A. Barros, T.G. Silva, M.G. da Rocha Pitta, D.P. Bezerra, L.V. Costa-Lotufo, M.O. de Moraes, C. Pessoa, M.A.F.B. de Moura, F.C. de Abreu, M.C. Alves de

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