International Journal of Biological Macromolecules 64 (2014) 53–62

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Interaction of cholinesterase modulators with DNA and their cytotoxic activity Jana Janockova a , Zuzana Gulasova a , Jana Plsikova a , Kamil Musilek b , Kamil Kuca c , Jaromir Mikes d , Lubomir Culka d , Peter Fedorocko d , Maria Kozurkova a,∗ Institute of Chemistry, Department of Biochemistry, P. J. Sˇ afárik University, Faculty of Science, Moyzesova 11, 04001 Kosice, Slovak Republic University of Hradec Kralove, Faculty of Science, Department of Chemistry, Rokitanskeho 62, 50003 Hradec Kralove, Czech Republic c University Hospital, Sokolska 581, 500 05 Hradec Kralove, Czech Republic d Institute of Biology and Ecology, Department of Cellular Biology, P. J. Sˇ afárik University, Faculty of Science, Moyzesova 11, 04001 Kosice, Slovak Republic a

b

a r t i c l e

i n f o

Article history: Received 14 October 2013 Received in revised form 22 November 2013 Accepted 25 November 2013 Available online 1 December 2013 Keywords: Cholinesterase modulators Oximes DNA Topoisomerase I HL-60 Cytotoxicity

a b s t r a c t This research was focused on a study of the binding properties of a series of cholinesterase reactivators compounds K075 (1), K027 (2) and inhibitors compounds K524, K009 and 7-MEOTA (3–5) with calf thymus DNA. The nature of the interactions between compounds 1–5 and DNA were studied using spectroscopic techniques (UV–vis, fluorescence spectroscopy and circular dichroism). The binding constants for complexes of cholinesterase modulators with DNA were determined from UV–vis spectroscopic titrations (K = 0.5 × 104 –8.9 × 105 M−1 ). The ability of the prepared analogues to relax topoisomerase I was studied with electrophoretic techniques and it was proved that ligands 4 and 5 inhibited this enzyme at a concentration of 30 ␮M. The biological activity of the novel compounds was assessed through an examination of changes in cell cycle distribution, mitochondrial membrane potential and cellular viability. Inhibitors 3–5 exhibited a cytotoxic effect on HL-60 (human acute promyelocytic leukaemia) cell culture, demonstrated a tendency to affect mitochondrial physiology and viability, and also forced cells to accumulate in the G1 /G0 -phase of the cell cycle. The cholinesterase reactivators 1 and 2 were found relatively save from the point of view of DNA binding, whereas cholinesterase inhibitors 3–5 resulted as strong DNA binding agents that limit their plausible use. © 2013 Elsevier B.V. All rights reserved.

1. Introduction The natural world is a rich source of chemically novel products with a broad spectrum of bioactivity and many compounds which are derived from organisms have generated scientific interest due to their naturally occurring cytotoxicity [1]. Schwartsmann et al. [2] have reported the existence of a number of interesting pyridinium derivatives which have been isolated from marine sponges. These biologically active compounds exert different biological activities, including cytotoxicity, hemolysis, antibacterial, antifungal or insecticidal effects, and also the inhibition of acetylcholinesterase (AChE) enzyme activity [1,3]. It has been suggested that AChE may be involved in development, differentiation, and pathogenic processes such as Alzheimer’s disease and tumorigenesis [4]. Efforts to inhibit acetylcholinesterase activity and the potential regulatory effects on cancer progression which could result from these efforts have

∗ Corresponding author. Tel.: +421 556223582; fax: +421 556222124. E-mail addresses: [email protected], [email protected] (M. Kozurkova). 0141-8130/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.ijbiomac.2013.11.022

been the focus of studies by several authors. Studies published by Hyatt et al. [5] 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 which is observed in cancer patients. Zovko et al. [1] found that some tumour types exhibit general changes during acetylcholinesterase activity. The presence of molecules related to the cholinergic signal system (in both healthy and carcinogenic lung tissue) has led to the hypothesis that substances which inhibit or affect the cholinergic signalling system could also possess anticancer effects. Cytotoxicity tests carried out on cell lines derived from lung tumours showed an AChE inhibition-dependent selective reduction of cell viability [1]. The same cells exhibited a loss in mitochondrial potential when exposed to non-toxic AChE inhibitors, which is a reaction characteristic of early apoptotic events. The same cells also exhibited a positive response to the annexin V affinity assay which is a specific feature of apoptosis. Paleari et al. [6] reports that polymeric alkylpyridinium salts may be a potential source of very promising anti-cancer agents due to their biological activity against tumours. The fact that they leave non-cancer cells, such as lymphocytes and organs, such as liver, heart and kidney, almost completely unaffected by their action.

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They suggest that the main anti-cancer effect may be exerted through AChE activity inhibition, because such an inhibition would enhance the permanence of acetylcholine (ACh) at the reciprocal sites. Recent years have also seen an increased use of AChE inhibitors in the treatment of a number of inflammatory and carcinogenic diseases in which ACh receptors may play a role [7]. Pyridinium oximes are used mainly as antidotes in cases of poisoning by organophosphorus compounds [8–10]. Other oximes are used in treatments of the cardiovascular system because of their anti-inflammatory, anti-viral, antibacterial and antifungal activities [11–16]. Oximes are also applied in agriculture, where are widely used as herbicides, pesticides and fungicides [17,18]. The cholinesterase modulators 1 [(E)-1,4-bis(4-hydroxyiminomethyl pyridinium)-but-2-ene dibromide, K075] and 2 [1-(4-carbonylpyridinium)-3-(4-hydroxyiminomethyl pyridinium)-propane dibromide, K027] have been tested for cholinesterase activity in the past and they were initially designed as acetylcholinesterase reactivators which counteract poisoning by organophosphorus compounds [19,20]. Both molecules have at least one oxime (hydroxyiminomethyl) functional group which is essential for their reactivation ability. Compound 2 was found to be a very effective reactivator of organophosphorus pesticides and also has the benefit of being a low toxic drug candidate [21–23]. Compound 1 was evaluated as a very potent reactivator of tabun-inhibited acetylcholinesterase, but its level of toxicity is above commercially acceptable standards [24,25]. Compounds 3 (1,10-bis(quinolinium)-dec-1,10-diyl dibromide, K524), 4 (10-methylacridinium iodide, K009) and 5 (9-amino7-methoxy-1,2,3,4-tetrahydroacridine hydrochloride, 7-MEOTA) were designed as cholinesterase inhibitors. Compound 5 was originally prepared and developed as a potential drug for the treatment of Alzheimer’s disease on account of its inhibitory ability towards acetylcholinesterase [25]. The resulting drug is a potent centrally active acetylcholinesterase inhibitor with minimal hepatotoxicity [26] which was also prepared with quaternary moiety [27]. Similarly, compound 3 was introduced as a peripherally acting cholinesterase inhibitor with implications in the treatment of Myasthenia gravis [28]. Compound 4 was designed as AChE inhibitor with potential to modulate activity of acetylcholine (muscarinic or nicotinic) receptors [29]. All compounds resulted in potent human AChE inhibitors on the ␮M or nM scale. The various groups of cholinesterase modulators vary in their toxicity profile. The oxime reactivators (e.g. 1 and 2) are usually found to be more toxic compounds, because they are just used once or continuously within 1–2 weeks after organophosphorus intoxication [30]. It means that their toxicity issues are minor compared to organophosphorus agents and they are overlapped by their lifesaving potency. Additionally, the used oximes have high hydrophilicity that enables their elimination from the body mostly within few hours [31]. The rapid oxime elimination also decreases their toxicity, but it is worthwhile to study it from the point of view of human safe use. On the other hand, the AChE inhibitors are administered in long-term treatment plan (Alzheimer disease, Myasthenia gravis) and they have to be low toxic compounds [26]. In case of peripheral AChE inhibitors (e.g. 3 and 4), they are also strongly hydrophilic compounds that have rapid clearance, which should support their low toxicity, which should be studied in longterm view. In contrast, neutral AChE inhibitors (e.g. 5) should have enhanced lipophilicity to be able BBB penetration and long lasting effect in CNS that presumes to study their toxicity profile. The rational for the investigation of the DNA interaction of these compounds lies in the toxicity to mice which they have demonstrated in vivo [32]. Most of the anticancer chemotherapeutic drugs in wide use today are DNA-damaging agents, and the targeting of DNA has been proven to result in the relatively potent and selective destruction of tumour cells. Thus, it had been supposed that

the interaction of the compounds examined in this study with DNA was due to their structural factors (e.g. condensed or closely connected aromatic rings). In this paper, the biochemical and biological activities of selected cholinesterase modulators (reactivators and inhibitors) are described and tested from the perspective of their activity as DNA-binding compounds. Their capacity to bind to DNA and to interfere with human topoisomerase I is studied, and the biological activity of the novel compounds are assessed using different techniques, such as examinations of cell cycle distribution and changes in mitochondrial membrane potential. 2. Material and methods 2.1. Materials The studied analogues (Table 1) were prepared prior to the experiment and were dissolved in DMSO to a final concentration of 2 × 104 ␮M. All chemicals and reagents purchased were of reagent grade and were used without further purification. Propidium iodide (PI), Hoechst 33342, ethidium bromide, Triton X-100, a reduced form of glutathione (GSH), dimethyl sulfoxide (DMSO, Serva) and calf thymus DNA were obtained from Sigma–Aldrich Chemie (Germany). EDTA, RNase A and proteinase K were purchased from Serva (Germany). Plazmid pUC 19 (2761 bp, DH 5˛), agarose (type II NoA-6877) (Sigma), 5,5-dithio-bis(2-nitrobenzoic acid) (DTNB) were purchased from Merck (Germany) and all other chemicals were purchased from Lachema (Czech Republic). 2.2. Methods 2.2.1. UV–vis absorption measurements UV–vis spectra were measured on a Varian Cary 100 UV-vis spectrophotometer in a 0.01 M Tris-HCl buffer (pH 7.4). A solution of calf thymus DNA (ctDNA) in a TE (Tris–EDTA) buffer was sonicated for 5 min and the DNA concentration was determined from its absorbance at 260 nm. The purity of DNA was determined by monitoring the value of A260 /A280 . DNA concentration was measured at 260 nm and expressed as micromolar equivalents of the base pairs ranging from 0 to 40 ␮M bp. Compounds 1–5 were dissolved in DMSO stock solution, from which working solutions were prepared through its dilution to a concentration of 25 ␮M using a 0.01 M Tris-HCl buffer. All measurements were performed at 24 ◦ C. 2.2.2. Fluorescence measurements Fluorescence measurements were carried out on a Varian Cary Eclipse spectrofluorimeter. All measurements were made using a 10 mm lightpath cuvette in a 0.01 M Tris-HCl buffer at pH 7.4. Emission spectra of derivatives 3–5 were recorded in the region of 350–600 nm using an excitation wavelength of 310–360 nm. Fluorescence intensities are expressed in arbitrary units. Fluorescence titrations were conducted by adding increasing amounts of ctDNA directly into the cell containing solutions of ligands 3–5 (c3 = 4.0 ␮M, c4 = 0.8 ␮M, c5 = 1.2 ␮M) in 0.01 M Tris-HCl buffer, pH 7.4. All measurements were performed at 24 ◦ C. The binding of ligands 1 and 2 to ctDNA was investigated using FID assay methods with ethidium bromide (EtBr) and a premixed solution of ctDNA (25.3 × 10−3 ␮M) and increasing amounts of 1 (c1 = 0–2.24 × 102 ␮M) and 2 (c2 = 0–1.68 × 102 ␮M) in a 0.01 M Tris-HCl buffer, pH 7.4. Emission spectra were collected from 550 to 800 nm with a 510 nm excitation of EtBr at 24 ◦ C. 2.2.3. Tm measurements Thermal denaturation studies were conducted using a Varian Cary Eclipse spectrophotometer equipped with a thermostatic

J. Janockova et al. / International Journal of Biological Macromolecules 64 (2014) 53–62

55

Table 1 The characteristics of studied compounds 1–5. Sign

Name

Mw [g mol−1 ]

Structure

HON=HC 1

2 Br N

(E)-1,4-bis(4hydroxyiminomethylpyridinium)but-2-ene dibromide (K075)

1,10-Bis(quinolinium)-dec1,10-diyl dibromide (K524)

4

10-Methylacridinium iodide (K009)

5

C H =N O H

2 Br

1-(4-Carbonylpyridinium)3-(4hydroxyiminomethylpyridinium)propane dibromide

3

458.15

CH=NOH

H 2N O C 2

N

N

446.14

N

558.39

N CH3

9-Amino-7-methoxy1,2,3,4-tetrahydroacridine hydrochloride (7-MEOTA)

cell holder. The temperature was controlled with a thermostatic bath (±0.1 ◦ C). The absorbance at 260 nm was monitored for either ctDNA (31.6 × 10−2 M) or a mixture of ctDNA with 1–5 (50 × 10−6 M) in BPE buffer, pH 7.1 (6 mM Na2 HPO4 , 2 mM NaH2 PO4 , 1 mM EDTA), with a heating rate of 1 ◦ C min−1 . The melting temperatures were determined as the maximum of the first derivative plots of the melting curves.

2.2.4. Equilibrium binding titration The binding affinities were calculated from the absorbance spectra according to the method developed by McGhee and von Hippel [33–35] using data points from a Scatchard plot. The binding data were fitted using GNU Octave 2.1.73 software [36].

321.16

I

250.72

2.2.5. Circular dichroism CD spectra of complexes ctDNA (31.6 × 10−1 ␮M bp) – ligands 1–5 (0.4 × 103 ␮M) were recorded 10 min after mixing on a J-810 Jasco spectropolarimeter at 24 ◦ C. All measurements were taken in a 0.01 M Tris-HCl buffer (pH 7.4). A rectangular quartz cell of 1 mm path length was used to obtain the spectra from 230 to 600 nm. Results are presented as a mean of three scans, from which the buffer background was electronically subtracted. 2.2.6. Topoisomerase I relaxation assay In order to determine topoisomerase I relaxation activity, calf thymus topoisomerase I (Takara, Japan) and pUC19 DNA (1.4 ␮g) were used 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 ␮L) containing 0.1% bovine serum albumin (BSA). The ligand was added and the reaction was initiated by the addition of 3

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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 a 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.

2.2.7. Cell culture and experimental design Cell culture HL-60 cells (human acute promyelocytic leukaemia) (ATCC, Rockville, MD, USA) were grown in suspension in a RPMI-1640 medium (Gibco Invitrogen Corp., Carlsbad, CA, USA) supplemented with a 10% foetal calf serum (FCS, Gibco Invitrogen Corp.) and antibiotics (penicillin 100 U mL−1 , streptomycin 100 ␮g mL−1 , and amphotericin 25 ␮g mL−1 ) (Gibco). The cultures were maintained at 37 ◦ C in a humidified atmosphere of 5% CO2 /95% air. Briefly description, 5 × 105 exponentially growing cells were seeded in Ø 60 mm Petri dishes (TPP, Switzerland) in a volume of 2 mL. Prior to the administering of the drug, the cells were allowed to settle for 1 h.

Fig. 1. Absorption spectra of derivative 3 (25 ␮M) in the absence and presence varying concentration of ctDNA (from top to bottom) at 24 ◦ C in a 0.01 M Tris-HCl medium buffered at pH 7.4.

2.2.8. Analysis of cell cycle parameters Cells were treated for 24, 48, or 72 h and subsequently harvested by centrifugation, washed in cold phosphate-buffered saline (PBS), fixed in cold 70% ethanol and stored overnight at 4 ◦ C. Prior to analysis, cells were washed twice in PBS, re-suspended in a staining solution (0.1% Triton X-100, 0.137 mg mL−1 of ribonuclease A and 0.02 mg mL−1 of propidium iodide or PI), incubated in darkness at rt for 30 min and analyzed using a FACSCalibur flow cytometer (Becton Dickinson, San Jose, CA, USA). ModFit 3.0 (Verity Software House, Topsham, ME, USA) software was used to generate DNA content frequency histograms and to quantify the number of cells in the individual cell cycle phases.

2.2.11. Statistical analysis The results were calculated as mean ± standard deviation (SD) of at least three independent experiments. The statistical significance was determined by one-way ANOVA followed by Tukey’s post hoc tests and the results were deemed significant if p < 0.01. 3. Results and discussion Compounds 1–5 were prepared according to the method which has been published previously [22–28] and are shown in Table 1. 3.1. Spectral and DNA binding studies

2.2.9. Detection of mitochondrial membrane potential (MMP) Cells (5 × 105 ) were treated for 24, 48, or 72 h and subsequently harvested by centrifugation, washed once with Hank’s balanced salt solution (HBSS), re-suspended in HBSS supplemented with 0.1 ␮M tetramethylrhodamine ethyl ester perchlorate (TMRE; Sigma–Aldrich, St. Louis, MO, USA), incubated for 20 min at 37 ◦ C in darkness and analyzed using a BD FACSCalibur flow cytometer. The cells (1 × 104 per sample) were gated according to FSC × SSC dot blot and analyzed in FL2 channel.

Conformational and structural changes in DNA following binding with the drug cause changes in the spectral features of the corresponding DNA, and hyperchromic and hypochromic effects in the double helix structure of DNA can be observed. Hypochromism is a result of contraction and changes in conformation of DNA in the helix axis, while hyperchromism results from damage of the DNA double helix structure [16]. The absorption spectra of compounds 1–5 (Table 1, 25 ␮M) were recorded at regular increased concentrations of ctDNA with corrections being made for the small volume changes which occurred during titration. They all exhibited broad absorption bands in the region of 300–450 nm which are typical for transitions between the ␲-electronic energy levels of the skeleton. The absorption spectra of compound 3 in the absence and presence of calf-thymus DNA (0–60.0 × 10−6 ␮M) are shown in Fig. 1. As the concentrations of DNA increased, the curve showed significant hypochromicity (13–42%) and a slight bathochromic shift indicating that the compounds can insert themselves into the DNA base. UV–vis data for all compounds 1–5 are displayed in Table 2. The intercalative mode of binding usually results in hypochromism and red shift due to strong stacking interactions

2.2.10. Viability/cellular metabolism Viability/cellular metabolism was analyzed using fluorescein diacetate (FDA; BD Pharmingen, Franklin Lanes, NJ, USA) and propidium iodide (PI; Sigma–Aldrich) double-staining. Cells (5 × 105 ) were treated for 24, 48, or 72 h and subsequently harvested using centrifugation, washed once with Hank’s balanced salt solution (HBSS) and stained with FDA in a HBSS buffer for 20 min. Prior to measurements, they were stained with PI (1 ␮g mL−1 ) and analyzed using the BD FACSCalibur flow cytometer. Results were analyzed using CellQuest Pro software.

Table 2 UV–vis absorption data of compounds 1–5. Tm (◦ C)

Compound

max free (nm)

max bound (nm)

 (nm)

Hypochromicity (%)

a

1 2 3 4 5

342 343 316 357 334

338 342 316 357 336

−4 −1 0 0 2

41.4 15.5 28.6 33.2 13.1

74.9 77.9 77.0 74.9 74.0

K (M−1 ) 6.0 × 104 0.5 × 104 8.9 × 105 7.0 × 105 7.9 × 104

a Tm measurements were performed in BPE buffer, pH 7.1 (6 mM Na2 HPO4 , 2 mM NaH2 PO4 , 1 mM EDTA) using 50.10−6 M drug and 32.6 × 10−2 M bp ctDNA with a heating rate of 1 ◦ C min−1 . Tm of ctDNA is 72.02 ◦ C.

J. Janockova et al. / International Journal of Biological Macromolecules 64 (2014) 53–62

3a

ctDNA

400

Fluorescence

ctDNA + 3

ΔA / ΔT

deltaA/deltaT

0,016

57

0,008

200

0

0 40

60

80

560

Temperature [°C]

720

800

400

Fig. 2. First derivative of the helix denaturation curves of ctDNA (solid line) with compound 3 (dashed line) measured at 260 nm in BPE buffer, pH 7.1.

Fluorescence [a.u.]

3b 300

200

100

0 360

400

440

480

520

Wavelength [nm] Fig. 3. (a) Fluorescence emission spectra of EtBr bound to DNA (25.3 × 10−3 ␮M) with increasing concentrations of ligand 1 (0–34 × 102 ␮M) at 2 ␮M steps, ex = 510 nm, in 10 mM Tris-HCl (pH = 7.4). (b) Spectrofluorimetric titration of derivative 3 (4.0 ␮M) in 0.01 M Tris-HCl buffer (pH 7.4, 24 ◦ C) by increasing the concentration of ctDNA (from top to bottom, 0–35 × 10−2 ␮M bp, at 2 ␮M intervals), ex = 316 nm.

4

ctDNA

2

ctDNA + 3

0

CD [mdeg]

between aromatic chromophores and the base pair of DNA. The extent of red shift and hypochromism are typically found to correlate to the intercalative binding strength, and complexes which bind non-intercalatively or electrostatistically with DNA may result in either hyperchromism or hypochromism. The low hypochromism effects observed in the presence of the studied compounds imply that they do not intercalate very strongly or deeply between the DNA base pairs [33]; it is evident that three of the complexes exhibit almost identical DNA binding affinities. This data was used to determine the binding constants of ligand complexes 1–5 with ctDNA using McGhee and von Hippel plots [34,35]. The constants were derived from nonlinear curve fitting, a process which was described previously [37]. The binding parameters from UV–vis spectroscopic analysis are summarized in Table 2. Large values of binding constants K in the range of from 0.5 × 104 to 8.9 × 105 M−1 prove a high affinity of ligands to DNA-base pairs. They are comparable to typical binding constants for intercalation of organic dyes into DNA (104 –106 M−1 ) and in good agreement with the binding affinity of standard proflavine hemisulphate (K = 7.95 × 105 M−1 ) and proflavine derivatives (K = 2.9 − 7.6 × 105 M−1 ) [33]. Thermal denaturation studies were used in order to establish further evidence of intercalation into DNA. Duplex DNA was thermally denatured into single-strand components in both the presence and absence of compounds. Compounds that physically associate with DNA typically stabilize the duplex when their presence increases the midpoint denaturation temperature (Tm ), and therefore the helix denaturation of DNA can be determined by monitoring the absorbance of DNA bases at 260 nm as a function of temperature (Tm ). However, Tm will increase slightly (Tm < 0.6 ◦ C), when interactions of small molecules with DNA through nonspecific electrostatic interactions with the phosphate backbone of DNA occur [38]. The first derivative of the DNA melting curve in the presence and absence of compound 3 is presented in Fig. 2. Tm of the ctDNA was 72.02 ◦ C in the absence of compounds 1–5 and increased from 74.9 to 77.9 ◦ C in the presence of 1–5 (Table 2). Therefore, it is possible to suggest that analogues 1–5 interact with DNA through the intercalative mode. However, changes in Tm alone are not sufficient proof of DNA intercalation, since these fluctuations can also be influenced by other kinds of molecules which recognize DNA, for example groove binders. A more accurate indicator of DNA intercalation is ethidium bromide displacement. Ligands 1 and 2 showed very low fluorescence intensity, therefore their binding was investigated using fluorescence titration of ethidium bromide (EtBr) with a premixed solution of ctDNA (25.3 × 10−3 ␮M). The maximal excitation of EtBr was recorded at 510 nm. Extrusion of EtBr from ctDNA was observed for both ligands 1 and 2 (Fig. 3a).

640

Wavelength [nm]

-2 -4 -6 -8

-10 250

300

350

400

450

500

Wavelength [nm] Fig. 4. CD spectrum of ctDNA (9.48 ␮M) in the absence and presence of compounds 3 and 5 (0.6 × 103 ␮M) in 0.01 M Tris-HCl buffer (pH 7.4).

Fig. 5. Electrophoresis agarose gel showing inhibition of calf thymus topoisomerase I induced DNA relaxation by compounds 1–5. Native supercoiled pUC19 (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 ␮M, lane b – 30 ␮M, lane c – 60 ␮M).

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Fig. 6. Cell cycle analysis. Cells were treated with derivatives 3–5 (individual concentrations are indicated) for up to 72 h. Cell cycle histograms are shown to demonstrate significant differences in cell cycle regulation and cell death induction (sub G1 /G0 phase). Representative set of data from three independent experiments is presented.

J. Janockova et al. / International Journal of Biological Macromolecules 64 (2014) 53–62 Table 3 Fluorescence characteristics of compounds 3–5. Compound

ex (nm)

em (nm)

F/F0 a

3 4 5

316 357 334

407 488 391

0.6 0.6 1.0

a

Fluorescence quantum yields were calculated using 5 as standard (˚f = 1).

The competitive binding of small molecules and EtBr to DNA can provide indirect information regarding the nature and relative strength of DNA binding. It is well known that while the intrinsic fluorescent intensity of EtBr is largely insignificant due to the quenching by the solvent molecules, it emits strong fluorescence in the presence of DNA [39]. If the second molecule also binds to DNA through the intercalative mode, the binding sites of DNA available for EtBr will be reduced and the fluorescent intensity of EtBr will consequently be quenched [40]. As was expected, the addition of DNA to the EtBr solution resulted in a significant enhancement of the fluorescent intensity of EtBr. When increased amounts of oxime compounds 1 and 2 were added to the EtBr–DNA mixture solution, the emission intensities of EtBr–DNA were seen to attenuate, indicating that more and more EtBr molecules had been released from the EtBr–DNA complex. This also suggests that oxime compounds 1 and 2 bind to DNA through an intercalation mode similar to that observed for EtBr. Free ligands 3–5 exhibited a broad emission band in the range of 350–600 nm. In Fig. 3b, the spectrofluorimetric titration of ligand 3 is depicted. The excitation wavelength was 316 nm and spectra were monitored at a fixed concentration of 4.0 ␮M. Titration with increasing concentrations of ctDNA (0–35 × 10−2 ␮M) continued until no further changes in the spectra of the drug–DNA complexes were recorded. Binding of the probes to DNA helix was found to reduce the fluorescence of the probes and this therefore represents further proof of intercalation. The binding parameters obtained from the spectrofluorimetric analysis of ligands 3–5 are summarized in Table 3. Circular dichroism (CD) is a powerful and reliable tool which is used to determine the conformation of biomacromolecules. Many DNA-binding ligands are achiral and are therefore optically inactive. Upon interaction with DNA, a molecule can acquire an induced CD signal through the coupling of electric transition moments of the ligand and the DNA bases. The observation of a CD signal with the absorption bands of the chiral ligand can provide confirmation of ligand–DNA interactions [41,42]. The CD spectrum of free ctDNA has a negative band at 245 nm due to helicity, and a positive band at 279 due to base stacking, which is characteristic of DNA in the

59

right-handed B form. When compounds 1–5 were incubated with ctDNA, the CD spectra displayed changes in both the positive and negative CD bands. The representative spectrum is displayed in Fig. 4. During the course of the binding titration, changes in the intensity alone would indicate a preference for the single binding mode; the shape of the curve of the observed CD would not be sufficient. Changes in the shape of CD signals may suggest either multiple ligand-binding modes, changes in DNA conformation or ligand–ligand interactions. 3.2. Topoisomerase I relaxation activity DNA topoisomerases are important enzymes in all living organisms; they are involved in the supercoiling of DNA and thus play important roles in many cellular metabolic processes including replication, transcription, recombination, repair and chromosome condensation. There has been considerable scientific interest in these enzymes, because a wide range of anticancer drugs are focused on the inhibition of DNA topoisomerases [6,43]. DNA topoisomerase I (Topo I) is an essential enzyme in mammalian cells [7,8] which creates a single strand break in DNA allowing relaxation of the DNA for replication [8,43]. The single strand break is then religated and the DNA double strand is restored. It has been shown that DNA intercalators can significantly interfere with this physiological process. The ligand that occupies the topoisomerase binding site may suppress the association of topoisomerase with DNA and influence the activity of topoisomerase. DNA intercalators, which inhibit topoisomerase activity or form stabilized ternary complexes with DNA and topoisomerase, have a high potential for the development of DNA-targeted anticancer drugs [44]. The cholinesterase modulators studied in this paper were evaluated for their Topo I relaxation activity. Fig. 5 illustrates the calf thymus DNA Topo I relaxation activity of compounds 1–5, all of which were incubated with topoisomerase I and pUC19. As shown, the supercoiled DNA was fully relaxed by the enzyme in the absence of the studied samples. The compounds 1–3 showed Topo I relaxation activity in the presence of ligands at a concentration of 60 ␮M. A larger fraction of the relaxed form observed for compounds 4 and 5 at a concentration of 30 ␮M. Comparable results were obtained for acridine [45] and similar oxime structures [46]. These findings may be a result of topoisomerase I relaxation together with the concurrent intercalation of ligands 1–5 into the topoisomerase binding site. We assume that the ternary complex consisting of topoisomerase, DNA and ligand is significantly stabilized under these conditions.

Table 4 Cell cycle distribution of compound 3. G1 24 h

48 h

72 h

Control 15 ␮M 25 ␮M 50 ␮M 100 ␮M Control 15 ␮M 25 ␮M 50 ␮M 100 ␮M Control 15 ␮M 25 ␮M 50 ␮M 100 ␮M

33.02 33.48 38.82 50.54 62.26 32.88 40.50 48.70 66.17 80.11 41.58 54.50 61.94 77.00 78.80

S ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.91 1.67 1.76* 3.34* 2.78* 0.45 4.89 3.68* 5.73* 3.94* 1.72 3.35 4.44* 4.22* 2.45*

49.59 50.04 46.42 38.84 28.40 48.77 49.35 40.87 24.68 13.36 43.13 35.95 28.44 16.23 19.48

G2 /M ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.56 2.26 1.47 2.72 2.78* 1.70 3.54 2.26 4.80 3.83* 2.31 2.66 3.89 3.60* 1.40*

17.39 16.47 14.75 10.62 9.34 18.35 10.15 10.43 9.15 6.52 15.29 9.54 9.62 6.77 2.58

± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

1.44 0.59 0.51 0.99* 0.03* 2.06* 1.38* 1.44* 0.95* 0.97* 2.46 0.75 0.81 0.70* 1.49*

The results are presented as a mean ± SD, statistical significance: p < 0.01 (*) for particular experimental group compared to untreated control.

Fig. 7. The effect of the compounds 3–5 on changes in mitochondrial membrane potential (MMP). The results are presented as a mean ± SD, statistical significance: p < 0.001 (*) for particular experimental group compared to untreated control.

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Fig. 8. The effect of the compound 3–5 on changes cell death induction. The results are presented as a mean ± SD, statistical significance: p < 0.001 (*) for particular experimental group compared to untreated control: (a) 24 h, (b) 48 h, (c) 72 h.

J. Janockova et al. / International Journal of Biological Macromolecules 64 (2014) 53–62 Table 5 Cell cycle distribution of compound 4. G1 24 h

48 h

72 h

Control 15 ␮M 25 ␮M 35 ␮M 50 ␮M Control 15 ␮M 25 ␮M 35 ␮M 50 ␮M Control 15 ␮M 25 ␮M 35 ␮M 50 ␮M

they also demonstrated high cytotoxic potential at concentrations of up to 100 ␮M. S

33.02 33.56 73.67 49.05 40.20 32.88 26.80 26.77 49.94 43.31 41.58 32.68 33.15 44.26 41.57

± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.91 7.39 24.14 21.64 0.58 0.45 6.35 5.44 11.09 3.25 1.72 3.60 4.53 4.75 13.58

61

49.59 37.91 7.79 51.72 49.57 48.77 49.13 84.16 39.95 46.83 43.13 50.44 44.63 39.58 39.27

G2 /M ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.56 9.38 2.74* 8.05 1.07 1.70 15.80 21.78 19.79 5.22 2.31 7.59 8.94 7.09 23.58

17.39 28.53 18.54 11.02 10.23 18.35 24.07 26.05 10.11 9.86 15.29 16.89 22.22 16.17 19.16

± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

1.44 2.38* 21.49 2.14 0.49 2.06 9.62 44.19 9.05 2.51* 2.46 4.18 13.19 9.77 14.30

For explanation see Table 4.

3.5. Viability/cellular metabolism Simultaneous analysis of viability (propidium iodide) and activity of cellular metabolism (fluorescein diacetate) showed similar results to those obtained in the analysis of  m dissipation. The results proved that molecules 1 and 2 are ineffective at concentrations of less than 100 ␮M. On the other hand, derivative 4 proved to be highly effective at the same concentration; for this reason, it was only tested at concentrations of up to 50 ␮M as the higher concentrations were highly cytotoxic when they were tested after 24 h of incubation (Fig. 8A). Compared to compound 4, compounds 3 and 5 demonstrated a delayed, although still distinct effect leading to cell destruction. The action of these three compounds was found to be dependent on both time and dose (Fig. 8).

3.3. Cell cycle

4. Conclusion

The tested compounds showed a cytotoxic effect, and molecules 3–5 in particular demonstrated a tendency to accumulate in the G1 /G0 -phase of the cell cycle. In case of the molecule 3, this trend was dependent on both dose and time over a 72 h period. Compounds 4–5 were also able to induce cell cycle arrest in G1 /G0 phase, however only at lower doses and within the initial 24 h (5), although higher concentrations of these compounds resulted in some changes which were deemed insignificant due to their high variability (Tables 4–6). Considering the other results which prove the high toxicity levels of these compounds ( m , viability/cellular metabolism), we presume that the cytotoxic action was fast and caused rapid cell destruction, given that a high proportion of the cell population was exterminated within the first 24 h (Fig. 6).

A series of cholinesterase modulators, compounds 1–5, was investigated. These compounds showed DNA binding activity (K = 0.5 × 104 to 8.9 × 105 M−1 ). UV–vis, fluorescence, circular dichroism spectroscopy, DNA melting techniques indicated that the studied compounds act as effective DNA-interacting agents. We have also confirmed their biological activity through experiments focused on analyzing their potential anticancer effects. Only derivatives 3–5 (cholinesterase inhibitors) proved to be highly effective against human acute promyelocytic leukaemia cells HL-60 (with derivatives 4 and 5 proving to be especially effective), evoking rapid mitochondrial membrane potential dissipation combined with a drop in cell metabolism and loss of viability at concentrations of up to 100 ␮M. Derivative 4 proved to be the most effective as it caused an almost total eradication of cells at a concentration of 50 ␮M. Based on obtained findings, cholinesterase reactivators 1 and 2 were found relatively save from the point of view of DNA binding, whereas cholinesterase inhibitors 3–5 (especially compound 4) resulted as strong DNA binding agents that limit their plausible use. We suggest that some of the derivatives presented in this study (compound 4) could have potential applications as anti-cancer drugs.

3.4. Dissipation of the mitochondrial membrane potential The molecules designated as 3–5 demonstrated a strong ability to affect mitochondrial physiology (Fig. 7). The dissipation of mitochondrial membrane potential  m is the hallmark of programmed cell death as it is accompanied by the release of proapoptotic molecules into the cytoplasm. Fundamentally, it is also a concomitant result of cell death in general. In concentrations ranging from 35 to 50 ␮M, compound 4 was able to evoke  m dissipation in approximately 80% of cells within 24 h and caused cell death with an efficiency approaching 100% after 48 h and 72 h incubation. The other two derivatives, 3 and 5, were less effective, but

Table 6 Cell cycle distribution of compound 5. G1 24 h

48 h

72 h

Control 25 ␮M 50 ␮M 75 ␮M 100 ␮M Control 25 ␮M 50 ␮M 75 ␮M 100 ␮M Control 25 ␮M 50 ␮M 75 ␮M 100 ␮M

33.02 38.81 46.82 53.14 46.43 32.88 39.79 57.69 45.55 29.39 41.58 41.84 54.68 39.71 50.87

For explanation see Table 4.

S ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.91 3.32 2.68* 3.46* 1.91* 0.45 3.44 5.70* 13.48 14.75 1.72 1.70 7.82 20.29 8.62

49.59 48.75 41.43 30.07 32.61 48.77 47.54 31.06 40.05 59.84 43.13 44.72 34.37 50.22 34.86

Acknowledgements This study was supported by Slovak Research and Development Agency (VVCE-0001-07, VEGA grant No. 1/0001/13, APVV-0280ˇ 11), Internal Grant Programme of the P. J. Safárik University in Koˇsice (VVGS-PF-2013-78), University Hospital in Hradec Kralove (long term development plan) and University Hradec Kralove (long term development plan).

G2 /M ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.56 2.23 2.73 1.28* 1.98* 1.70 1.66 4.61 15.98 28.72 2.31 0.73 6.24 29.81 4.34

17.39 12.43 11.75 16.79 20.96 18.35 12.66 11.25 14.40 10.77 15.29 13.44 10.94 15.10 14.27

± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

1.44 1.23* 0.61 2.29 3.40 2.06 1.79* 1.20* 2.50* 13.97 2.46 1.05 1.60 9.58 12.02

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