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

Contents lists available at ScienceDirect

International Journal of Biological Macromolecules

jo ur nal homep age: www.elsev ier .com/ locate / i jb iomac

nteraction of cholinesterase modulators with DNA and theirytotoxic activity

ana Janockovaa, Zuzana Gulasovaa, Jana Plsikovaa, Kamil Musilekb, Kamil Kucac,aromir Mikesd, Lubomir Culkad, Peter Fedorockod, Maria Kozurkovaa,∗

Institute of Chemistry, Department of Biochemistry, P. J. Safárik University, Faculty of Science, Moyzesova 11, 04001 Kosice, Slovak RepublicUniversity of Hradec Kralove, Faculty of Science, Department of Chemistry, Rokitanskeho 62, 50003 Hradec Kralove, Czech RepublicUniversity Hospital, Sokolska 581, 500 05 Hradec Kralove, Czech RepublicInstitute of Biology and Ecology, Department of Cellular Biology, P. J. Safárik University, Faculty of Science, Moyzesova 11, 04001 Kosice, Slovak Republic

r t i c l e i n f o

rticle history:eceived 14 October 2013eceived in revised form2 November 2013ccepted 25 November 2013vailable online 1 December 2013

eywords:holinesterase modulators

a b s t r a c t

This research was focused on a study of the binding properties of a series of cholinesterase reactiva-tors compounds K075 (1), K027 (2) and inhibitors compounds K524, K009 and 7-MEOTA (3–5) with calfthymus DNA. The nature of the interactions between compounds 1–5 and DNA were studied using spec-troscopic techniques (UV–vis, fluorescence spectroscopy and circular dichroism). The binding constantsfor complexes of cholinesterase modulators with DNA were determined from UV–vis spectroscopic titra-tions (K = 0.5 × 104–8.9 × 105 M−1). The ability of the prepared analogues to relax topoisomerase I wasstudied with electrophoretic techniques and it was proved that ligands 4 and 5 inhibited this enzymeat a concentration of 30 �M. The biological activity of the novel compounds was assessed through an

ximesNAopoisomerase IL-60ytotoxicity

examination of changes in cell cycle distribution, mitochondrial membrane potential and cellular via-bility. Inhibitors 3–5 exhibited a cytotoxic effect on HL-60 (human acute promyelocytic leukaemia) cellculture, demonstrated a tendency to affect mitochondrial physiology and viability, and also forced cellsto accumulate in the G1/G0-phase of the cell cycle. The cholinesterase reactivators 1 and 2 were foundrelatively save from the point of view of DNA binding, whereas cholinesterase inhibitors 3–5 resulted asstrong DNA binding agents that limit their plausible use.

. Introduction

The natural world is a rich source of chemically novel productsith a broad spectrum of bioactivity and many compounds which

re derived from organisms have generated scientific interest dueo their naturally occurring cytotoxicity [1]. Schwartsmann et al. [2]ave reported the existence of a number of interesting pyridiniumerivatives which have been isolated from marine sponges. Theseiologically active compounds exert different biological activities,

ncluding cytotoxicity, hemolysis, antibacterial, antifungal or insec-icidal effects, and also the inhibition of acetylcholinesterase (AChE)nzyme activity [1,3].

It has been suggested that AChE may be involved inevelopment, differentiation, and pathogenic processes such as

lzheimer’s disease and tumorigenesis [4]. Efforts to inhibit acetyl-holinesterase activity and the potential regulatory effects onancer progression which could result from these efforts have

∗ Corresponding author. Tel.: +421 556223582; fax: +421 556222124.E-mail addresses: maria.kozurkova@upjs.sk, mariakozurkova@yahoo.com

M. Kozurkova).

141-8130/$ – see front matter © 2013 Elsevier B.V. All rights reserved.ttp://dx.doi.org/10.1016/j.ijbiomac.2013.11.022

© 2013 Elsevier B.V. All rights reserved.

been the focus of studies by several authors. Studies publishedby Hyatt et al. [5] indicate that CPT-11 (irinotecan, 7-ethyl-10-[4-(1-piperidino)-1-piperidino]carbonyloxycamptothecin) is a potentinhibitor of AChE, and this probably accounts for the choliner-gic syndrome which is observed in cancer patients. Zovko et al.[1] found that some tumour types exhibit general changes dur-ing acetylcholinesterase activity. The presence of molecules relatedto the cholinergic signal system (in both healthy and carcinogeniclung tissue) has led to the hypothesis that substances which inhibitor affect the cholinergic signalling system could also possess anti-cancer effects. Cytotoxicity tests carried out on cell lines derivedfrom lung tumours showed an AChE inhibition-dependent selectivereduction of cell viability [1]. The same cells exhibited a loss in mito-chondrial potential when exposed to non-toxic AChE inhibitors,which is a reaction characteristic of early apoptotic events. Thesame cells also exhibited a positive response to the annexin V affin-ity 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 dueto their biological activity against tumours. The fact that they leavenon-cancer cells, such as lymphocytes and organs, such as liver,heart and kidney, almost completely unaffected by their action.

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4 J. Janockova et al. / International Journa

hey suggest that the main anti-cancer effect may be exertedhrough AChE activity inhibition, because such an inhibition wouldnhance the permanence of acetylcholine (ACh) at the recipro-al sites. Recent years have also seen an increased use of AChEnhibitors in the treatment of a number of inflammatory and car-inogenic diseases in which ACh receptors may play a role [7].

Pyridinium oximes are used mainly as antidotes in cases of poi-oning by organophosphorus compounds [8–10]. Other oximes aresed in treatments of the cardiovascular system because of theirnti-inflammatory, anti-viral, antibacterial and antifungal activi-ies [11–16]. Oximes are also applied in agriculture, where areidely used as herbicides, pesticides and fungicides [17,18].

The cholinesterase modulators 1 [(E)-1,4-bis(4-hydroxy-minomethyl pyridinium)-but-2-ene dibromide, K075] and 21-(4-carbonylpyridinium)-3-(4-hydroxyiminomethyl pyrid-nium)-propane dibromide, K027] have been tested forholinesterase activity in the past and they were initially designeds acetylcholinesterase reactivators which counteract poison-ng by organophosphorus compounds [19,20]. Both moleculesave at least one oxime (hydroxyiminomethyl) functional grouphich is essential for their reactivation ability. Compound 2 was

ound to be a very effective reactivator of organophosphorusesticides and also has the benefit of being a low toxic drugandidate [21–23]. Compound 1 was evaluated as a very potenteactivator of tabun-inhibited acetylcholinesterase, but its levelf toxicity is above commercially acceptable standards [24,25].ompounds 3 (1,10-bis(quinolinium)-dec-1,10-diyl dibromide,524), 4 (10-methylacridinium iodide, K009) and 5 (9-amino--methoxy-1,2,3,4-tetrahydroacridine hydrochloride, 7-MEOTA)ere designed as cholinesterase inhibitors. Compound 5 was

riginally prepared and developed as a potential drug for thereatment of Alzheimer’s disease on account of its inhibitorybility towards acetylcholinesterase [25]. The resulting drug is aotent centrally active acetylcholinesterase inhibitor with minimalepatotoxicity [26] which was also prepared with quaternary moi-ty [27]. Similarly, compound 3 was introduced as a peripherallycting cholinesterase inhibitor with implications in the treatmentf Myasthenia gravis [28]. Compound 4 was designed as AChEnhibitor with potential to modulate activity of acetylcholinemuscarinic or nicotinic) receptors [29]. All compounds resultedn potent human AChE inhibitors on the �M or nM scale.

The various groups of cholinesterase modulators vary in theiroxicity profile. The oxime reactivators (e.g. 1 and 2) are usuallyound to be more toxic compounds, because they are just usednce or continuously within 1–2 weeks after organophosphorusntoxication [30]. It means that their toxicity issues are minor com-ared to organophosphorus agents and they are overlapped byheir lifesaving potency. Additionally, the used oximes have highydrophilicity that enables their elimination from the body mostlyithin few hours [31]. The rapid oxime elimination also decreases

heir toxicity, but it is worthwhile to study it from the point ofiew of human safe use. On the other hand, the AChE inhibitorsre administered in long-term treatment plan (Alzheimer disease,yasthenia gravis) and they have to be low toxic compounds [26].

n case of peripheral AChE inhibitors (e.g. 3 and 4), they are alsotrongly hydrophilic compounds that have rapid clearance, whichhould support their low toxicity, which should be studied in long-erm view. In contrast, neutral AChE inhibitors (e.g. 5) should havenhanced lipophilicity to be able BBB penetration and long lastingffect in CNS that presumes to study their toxicity profile.

The rational for the investigation of the DNA interaction of theseompounds lies in the toxicity to mice which they have demon-

trated in vivo [32]. Most of the anticancer chemotherapeutic drugsn wide use today are DNA-damaging agents, and the targeting ofNA has been proven to result in the relatively potent and selec-

ive destruction of tumour cells. Thus, it had been supposed that

logical Macromolecules 64 (2014) 53– 62

the interaction of the compounds examined in this study with DNAwas due to their structural factors (e.g. condensed or closely con-nected aromatic rings). In this paper, the biochemical and biologicalactivities of selected cholinesterase modulators (reactivators andinhibitors) are described and tested from the perspective of theiractivity as DNA-binding compounds. Their capacity to bind to DNAand to interfere with human topoisomerase I is studied, and thebiological activity of the novel compounds are assessed using dif-ferent techniques, such as examinations of cell cycle distributionand changes in mitochondrial membrane potential.

2. Material and methods

2.1. Materials

The studied analogues (Table 1) were prepared prior to theexperiment and were dissolved in DMSO to a final concentration of2 × 104 �M.

All chemicals and reagents purchased were of reagent gradeand were used without further purification. Propidium iodide (PI),Hoechst 33342, ethidium bromide, Triton X-100, a reduced form ofglutathione (GSH), dimethyl sulfoxide (DMSO, Serva) and calf thy-mus 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 No-A-6877) (Sigma), 5,5-dithio-bis(2-nitrobenzoic acid) (DTNB) werepurchased from Merck (Germany) and all other chemicals werepurchased from Lachema (Czech Republic).

2.2. Methods

2.2.1. UV–vis absorption measurementsUV–vis spectra were measured on a Varian Cary 100 UV-vis

spectrophotometer in a 0.01 M Tris-HCl buffer (pH 7.4). A solu-tion of calf thymus DNA (ctDNA) in a TE (Tris–EDTA) buffer wassonicated for 5 min and the DNA concentration was determinedfrom its absorbance at 260 nm. The purity of DNA was determinedby monitoring the value of A260/A280. DNA concentration was mea-sured at 260 nm and expressed as micromolar equivalents of thebase pairs ranging from 0 to 40 �M bp. Compounds 1–5 were dis-solved in DMSO stock solution, from which working solutions wereprepared through its dilution to a concentration of 25 �M using a0.01 M Tris-HCl buffer. All measurements were performed at 24 ◦C.

2.2.2. Fluorescence measurementsFluorescence measurements were carried out on a Varian Cary

Eclipse spectrofluorimeter. All measurements were made usinga 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 regionof 350–600 nm using an excitation wavelength of 310–360 nm.Fluorescence intensities are expressed in arbitrary units. Fluores-cence titrations were conducted by adding increasing amounts ofctDNA 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, pH7.4. All measurements were performed at 24 ◦C.

The binding of ligands 1 and 2 to ctDNA was investigated usingFID assay methods with ethidium bromide (EtBr) and a premixedsolution 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 MTris-HCl buffer, pH 7.4. Emission spectra were collected from 550to 800 nm with a 510 nm excitation of EtBr at 24 ◦C.

2.2.3. Tm measurementsThermal denaturation studies were conducted using a Var-

ian Cary Eclipse spectrophotometer equipped with a thermostatic

J. Janockova et al. / International Journal of Biological Macromolecules 64 (2014) 53– 62 55

Table 1The characteristics of studied compounds 1–5.

Sign Name Structure Mw [g mol−1]

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

2 B rN

N

H O N = H C

C H = N O H

458.15

2 1-(4-Carbonylpyridinium)-3-(4-hydroxyiminomethylpyridinium)-propanedibromide

N N2 Br

H2N O C C H =N O H446.14

3 1,10-Bis(quinolinium)-dec-1,10-diyl dibromide(K524)

558.39

4 10-Methylacridiniumiodide (K009) N

CH3 I321.16

5 9-Amino-7-methoxy-1,2,3,4-tetrahydroacridinehydrochloride (7-MEOTA)

250.72

csf1Nid

2

svT[

ell holder. The temperature was controlled with a thermo-tatic bath (±0.1 ◦C). The absorbance at 260 nm was monitoredor either ctDNA (31.6 × 10−2 M) or a mixture of ctDNA with–5 (50 × 10−6 M) in BPE buffer, pH 7.1 (6 mM Na2HPO4, 2 mMaH2PO4, 1 mM EDTA), with a heating rate of 1 ◦C min−1. The melt-

ng temperatures were determined as the maximum of the firsterivative plots of the melting curves.

.2.4. Equilibrium binding titrationThe binding affinities were calculated from the absorbance

pectra according to the method developed by McGhee andon Hippel [33–35] using data points from a Scatchard plot.he binding data were fitted using GNU Octave 2.1.73 software36].

2.2.5. Circular dichroismCD 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-810Jasco spectropolarimeter at 24 ◦C. All measurements were taken ina 0.01 M Tris-HCl buffer (pH 7.4). A rectangular quartz cell of 1 mmpath length was used to obtain the spectra from 230 to 600 nm.Results are presented as a mean of three scans, from which thebuffer background was electronically subtracted.

2.2.6. Topoisomerase I relaxation assayIn 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 ligandwas added and the reaction was initiated by the addition of 3

5 l of Biological Macromolecules 64 (2014) 53– 62

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6 J. Janockova et al. / International Journa

nits of topoisomerase I. The reactions were carried out at 37 ◦Cor 45 min. A gel electrophoresis was performed at 7 V/cm for 2 hn a TBE (Tris, boric acid, EDTA) buffer on a 1% agarose gel. The gel

as stained with ethidium bromide (1 mg/ml) and photographednder UV light.

.2.7. Cell culture and experimental designCell culture HL-60 cells (human acute promyelocytic leukaemia)

ATCC, Rockville, MD, USA) were grown in suspension in aPMI-1640 medium (Gibco Invitrogen Corp., Carlsbad, CA, USA)upplemented with a 10% foetal calf serum (FCS, Gibco Invitro-en Corp.) and antibiotics (penicillin 100 U mL−1, streptomycin00 �g mL−1, and amphotericin 25 �g mL−1) (Gibco). The culturesere maintained at 37 ◦C in a humidified atmosphere of 5% CO2/95%

ir. Briefly description, 5 × 105 exponentially growing cells wereeeded in Ø 60 mm Petri dishes (TPP, Switzerland) in a volume of

mL. Prior to the administering of the drug, the cells were allowedo settle for 1 h.

.2.8. Analysis of cell cycle parametersCells were treated for 24, 48, or 72 h and subsequently harvested

y centrifugation, washed in cold phosphate-buffered saline (PBS),xed in cold 70% ethanol and stored overnight at 4 ◦C. Prior to anal-sis, cells were washed twice in PBS, re-suspended in a stainingolution (0.1% Triton X-100, 0.137 mg mL−1 of ribonuclease A and.02 mg mL−1 of propidium iodide or PI), incubated in darknesst rt for 30 min and analyzed using a FACSCalibur flow cytometerBecton Dickinson, San Jose, CA, USA). ModFit 3.0 (Verity Softwareouse, Topsham, ME, USA) software was used to generate DNA con-

ent frequency histograms and to quantify the number of cells inhe individual cell cycle phases.

.2.9. Detection of mitochondrial membrane potential (MMP)Cells (5 × 105) were treated for 24, 48, or 72 h and subse-

uently harvested by centrifugation, washed once with Hank’salanced salt solution (HBSS), re-suspended in HBSS supplementedith 0.1 �M tetramethylrhodamine ethyl ester perchlorate (TMRE;

igma–Aldrich, St. Louis, MO, USA), incubated for 20 min at 37 ◦Cn darkness and analyzed using a BD FACSCalibur flow cytometer.he cells (1 × 104 per sample) were gated according to FSC × SSCot blot and analyzed in FL2 channel.

.2.10. Viability/cellular metabolismViability/cellular metabolism was analyzed using fluorescein

iacetate (FDA; BD Pharmingen, Franklin Lanes, NJ, USA) and pro-idium iodide (PI; Sigma–Aldrich) double-staining. Cells (5 × 105)ere treated for 24, 48, or 72 h and subsequently harvested using

entrifugation, washed once with Hank’s balanced salt solution

HBSS) and stained with FDA in a HBSS buffer for 20 min. Prior to

easurements, they were stained with PI (1 �g mL−1) and analyzedsing the BD FACSCalibur flow cytometer. Results were analyzedsing CellQuest Pro software.

able 2V–vis absorption data of compounds 1–5.

Compound �max free (nm) �max bound (nm) ��

1 342 338 −4

2 343 342 −1

3 316 316 0

4 357 357 0

5 334 336 2

a Tm measurements were performed in BPE buffer, pH 7.1 (6 mM Na2HPO4, 2 mM NaH2

ate of 1 ◦C min−1. Tm of ctDNA is 72.02 ◦C.

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

2.2.11. Statistical analysisThe results were calculated as mean ± standard deviation (SD) of

at least three independent experiments. The statistical significancewas determined by one-way ANOVA followed by Tukey’s post hoctests and the results were deemed significant if p < 0.01.

3. Results and discussion

Compounds 1–5 were prepared according to the method whichhas been published previously [22–28] and are shown in Table 1.

3.1. Spectral and DNA binding studies

Conformational and structural changes in DNA following bind-ing with the drug cause changes in the spectral features of thecorresponding DNA, and hyperchromic and hypochromic effects inthe double helix structure of DNA can be observed. Hypochromismis a result of contraction and changes in conformation of DNA in thehelix axis, while hyperchromism results from damage of the DNAdouble helix structure [16]. The absorption spectra of compounds1–5 (Table 1, 25 �M) were recorded at regular increased concentra-tions of ctDNA with corrections being made for the small volumechanges which occurred during titration. They all exhibited broadabsorption bands in the region of 300–450 nm which are typical fortransitions between the �-electronic energy levels of the skeleton.The absorption spectra of compound 3 in the absence and pres-ence of calf-thymus DNA (0–60.0 × 10−6 �M) are shown in Fig. 1. Asthe concentrations of DNA increased, the curve showed significanthypochromicity (13–42%) and a slight bathochromic shift indicat-

ing 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 inhypochromism and red shift due to strong stacking interactions

(nm) Hypochromicity (%) aTm (◦C) K (M−1)

41.4 74.9 6.0 × 104

15.5 77.9 0.5 × 104

28.6 77.0 8.9 × 105

33.2 74.9 7.0 × 105

13.1 74.0 7.9 × 104

PO4, 1 mM EDTA) using 50.10−6 M drug and 32.6 × 10−2 M bp ctDNA with a heating

J. Janockova et al. / International Journal of Biological Macromolecules 64 (2014) 53– 62 57

806040

0,016

0,008

0

Temperature [°C]

deltaA/deltaT

ctDNA

ctDNA + 3ΔA

/ ΔT

Fc

becwmhcdt

lpcbmrob(ip

ltpaetitwcoecfpt

iobb

fo(51

800720640560

400

200

0

Wavelength [nm]

Fluorescence

520480440400360

400

300

200

100

0

Wavelength [nm]

Fluorescence[a.u.]

3b

3a

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 ofderivative 3 (4.0 �M) in 0.01 M Tris-HCl buffer (pH 7.4, 24 ◦C) by increasing theconcentration of ctDNA (from top to bottom, 0–35 × 10−2 �M bp, at 2 �M intervals),�ex = 316 nm.

500450400350300250

4

2

0

-2

-4

-6

-8

-10

Wavelength [nm]

CD

[mde

g]

ctDNA

ctDNA + 3

Fig. 4. CD spectrum of ctDNA (9.48 �M) in the absence and presence of compounds3 and 5 (0.6 × 103 �M) in 0.01 M Tris-HCl buffer (pH 7.4).

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

etween aromatic chromophores and the base pair of DNA. Thextent of red shift and hypochromism are typically found toorrelate to the intercalative binding strength, and complexeshich bind non-intercalatively or electrostatistically with DNAay result in either hyperchromism or hypochromism. The low

ypochromism effects observed in the presence of the studiedompounds imply that they do not intercalate very strongly oreeply between the DNA base pairs [33]; it is evident that three ofhe complexes exhibit almost identical DNA binding affinities.

This data was used to determine the binding constants ofigand complexes 1–5 with ctDNA using McGhee and von Hip-el plots [34,35]. The constants were derived from nonlinearurve fitting, a process which was described previously [37]. Theinding parameters from UV–vis spectroscopic analysis are sum-arized in Table 2. Large values of binding constants K in the

ange of from 0.5 × 104 to 8.9 × 105 M−1 prove a high affinityf ligands to DNA-base pairs. They are comparable to typicalinding constants for intercalation of organic dyes into DNA104–106 M−1) and in good agreement with the binding affin-ty of standard proflavine hemisulphate (K = 7.95 × 105 M−1) androflavine derivatives (K = 2.9 − 7.6 × 105 M−1) [33].

Thermal denaturation studies were used in order to estab-ish further evidence of intercalation into DNA. Duplex DNA washermally denatured into single-strand components in both theresence and absence of compounds. Compounds that physicallyssociate with DNA typically stabilize the duplex when their pres-nce increases the midpoint denaturation temperature (Tm), andherefore the helix denaturation of DNA can be determined by mon-toring the absorbance of DNA bases at 260 nm as a function ofemperature (Tm). However, Tm will increase slightly (Tm < 0.6 ◦C),hen interactions of small molecules with DNA through nonspe-

ific electrostatic interactions with the phosphate backbone of DNAccur [38]. The first derivative of the DNA melting curve in the pres-nce and absence of compound 3 is presented in Fig. 2. Tm of thetDNA was 72.02 ◦C in the absence of compounds 1–5 and increasedrom 74.9 to 77.9 ◦C in the presence of 1–5 (Table 2). Therefore, it isossible to suggest that analogues 1–5 interact with DNA throughhe intercalative mode.

However, changes in Tm alone are not sufficient proof of DNAntercalation, since these fluctuations can also be influenced byther kinds of molecules which recognize DNA, for example grooveinders. A more accurate indicator of DNA intercalation is ethidiumromide displacement.

Ligands 1 and 2 showed very low fluorescence intensity, there-ore their binding was investigated using fluorescence titration

f ethidium bromide (EtBr) with a premixed solution of ctDNA25.3 × 10−3 �M). The maximal excitation of EtBr was recorded at10 nm. Extrusion of EtBr from ctDNA was observed for both ligands

and 2 (Fig. 3a).

Fig. 5. Electrophoresis agarose gel showing inhibition of calf thymus topoisomeraseI 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) orpresence of ligands (lane a – 5 �M, lane b – 30 �M, lane c – 60 �M).

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

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 demonstratesignificant 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 Bio

Table 3Fluorescence characteristics of compounds 3–5.

Compound �ex (nm) �em (nm) F/F0a

3 316 407 0.64 357 488 0.6

cssqiDawotctct1o

33twtcfros

uDtClalha

TC

Tp

the concurrent intercalation of ligands 1–5 into the topoisomerase

5 334 391 1.0

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

The competitive binding of small molecules and EtBr to DNAan provide indirect information regarding the nature and relativetrength of DNA binding. It is well known that while the intrin-ic fluorescent intensity of EtBr is largely insignificant due to theuenching by the solvent molecules, it emits strong fluorescence

n the presence of DNA [39]. If the second molecule also binds toNA through the intercalative mode, the binding sites of DNA avail-ble for EtBr will be reduced and the fluorescent intensity of EtBrill consequently be quenched [40]. As was expected, the addition

f DNA to the EtBr solution resulted in a significant enhancement ofhe fluorescent intensity of EtBr. When increased amounts of oximeompounds 1 and 2 were added to the EtBr–DNA mixture solution,he emission intensities of EtBr–DNA were seen to attenuate, indi-ating that more and more EtBr molecules had been released fromhe EtBr–DNA complex. This also suggests that oxime compounds

and 2 bind to DNA through an intercalation mode similar to thatbserved for EtBr.

Free ligands 3–5 exhibited a broad emission band in the range of50–600 nm. In Fig. 3b, the spectrofluorimetric titration of ligand

is depicted. The excitation wavelength was 316 nm and spec-ra were monitored at a fixed concentration of 4.0 �M. Titrationith increasing concentrations of ctDNA (0–35 × 10−2 �M) con-

inued until no further changes in the spectra of the drug–DNAomplexes were recorded. Binding of the probes to DNA helix wasound to reduce the fluorescence of the probes and this thereforeepresents further proof of intercalation. The binding parametersbtained from the spectrofluorimetric analysis of ligands 3–5 areummarized in Table 3.

Circular dichroism (CD) is a powerful and reliable tool which issed to determine the conformation of biomacromolecules. ManyNA-binding ligands are achiral and are therefore optically inac-

ive. Upon interaction with DNA, a molecule can acquire an inducedD signal through the coupling of electric transition moments of the

igand and the DNA bases. The observation of a CD signal with the

bsorption bands of the chiral ligand can provide confirmation ofigand–DNA interactions [41,42]. The CD spectrum of free ctDNAas a negative band at 245 nm due to helicity, and a positive bandt 279 due to base stacking, which is characteristic of DNA in the

able 4ell cycle distribution of compound 3.

G1 S G2/M

24 h Control 33.02 ± 0.91 49.59 ± 0.56 17.39 ± 1.4415 �M 33.48 ± 1.67 50.04 ± 2.26 16.47 ± 0.5925 �M 38.82 ± 1.76* 46.42 ± 1.47 14.75 ± 0.5150 �M 50.54 ± 3.34* 38.84 ± 2.72 10.62 ± 0.99*100 �M 62.26 ± 2.78* 28.40 ± 2.78* 9.34 ± 0.03*

48 h Control 32.88 ± 0.45 48.77 ± 1.70 18.35 ± 2.06*15 �M 40.50 ± 4.89 49.35 ± 3.54 10.15 ± 1.38*25 �M 48.70 ± 3.68* 40.87 ± 2.26 10.43 ± 1.44*50 �M 66.17 ± 5.73* 24.68 ± 4.80 9.15 ± 0.95*100 �M 80.11 ± 3.94* 13.36 ± 3.83* 6.52 ± 0.97*

72 h Control 41.58 ± 1.72 43.13 ± 2.31 15.29 ± 2.4615 �M 54.50 ± 3.35 35.95 ± 2.66 9.54 ± 0.7525 �M 61.94 ± 4.44* 28.44 ± 3.89 9.62 ± 0.8150 �M 77.00 ± 4.22* 16.23 ± 3.60* 6.77 ± 0.70*100 �M 78.80 ± 2.45* 19.48 ± 1.40* 2.58 ± 1.49*

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

logical Macromolecules 64 (2014) 53– 62 59

right-handed B form. When compounds 1–5 were incubated withctDNA, the CD spectra displayed changes in both the positive andnegative CD bands. The representative spectrum is displayed inFig. 4. During the course of the binding titration, changes in theintensity alone would indicate a preference for the single bindingmode; the shape of the curve of the observed CD would not besufficient. Changes in the shape of CD signals may suggest eithermultiple ligand-binding modes, changes in DNA conformation orligand–ligand interactions.

3.2. Topoisomerase I relaxation activity

DNA topoisomerases are important enzymes in all living orga-nisms; they are involved in the supercoiling of DNA and thus playimportant roles in many cellular metabolic processes includingreplication, transcription, recombination, repair and chromosomecondensation. There has been considerable scientific interest inthese enzymes, because a wide range of anticancer drugs arefocused on the inhibition of DNA topoisomerases [6,43].

DNA topoisomerase I (Topo I) is an essential enzyme in mam-malian cells [7,8] which creates a single strand break in DNAallowing relaxation of the DNA for replication [8,43]. The sin-gle strand break is then religated and the DNA double strand isrestored. It has been shown that DNA intercalators can significantlyinterfere with this physiological process. The ligand that occupiesthe topoisomerase binding site may suppress the association oftopoisomerase with DNA and influence the activity of topoiso-merase. DNA intercalators, which inhibit topoisomerase activity orform stabilized ternary complexes with DNA and topoisomerase,have a high potential for the development of DNA-targeted anti-cancer drugs [44].

The cholinesterase modulators studied in this paper were eval-uated for their Topo I relaxation activity. Fig. 5 illustrates the calfthymus DNA Topo I relaxation activity of compounds 1–5, all ofwhich were incubated with topoisomerase I and pUC19. As shown,the supercoiled DNA was fully relaxed by the enzyme in the absenceof the studied samples. The compounds 1–3 showed Topo I relax-ation activity in the presence of ligands at a concentration of 60 �M.A larger fraction of the relaxed form observed for compounds 4 and5 at a concentration of 30 �M. Comparable results were obtainedfor acridine [45] and similar oxime structures [46]. These find-ings may be a result of topoisomerase I relaxation together with

binding site. We assume that the ternary complex consisting oftopoisomerase, DNA and ligand is significantly stabilized underthese conditions.

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

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

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 experimentalgroup compared to untreated control: (a) 24 h, (b) 48 h, (c) 72 h.

J. Janockova et al. / International Journal of Bio

Table 5Cell cycle distribution of compound 4.

G1 S G2/M

24 h Control 33.02 ± 0.91 49.59 ± 0.56 17.39 ± 1.4415 �M 33.56 ± 7.39 37.91 ± 9.38 28.53 ± 2.38*25 �M 73.67 ± 24.14 7.79 ± 2.74* 18.54 ± 21.4935 �M 49.05 ± 21.64 51.72 ± 8.05 11.02 ± 2.1450 �M 40.20 ± 0.58 49.57 ± 1.07 10.23 ± 0.49

48 h Control 32.88 ± 0.45 48.77 ± 1.70 18.35 ± 2.0615 �M 26.80 ± 6.35 49.13 ± 15.80 24.07 ± 9.6225 �M 26.77 ± 5.44 84.16 ± 21.78 26.05 ± 44.1935 �M 49.94 ± 11.09 39.95 ± 19.79 10.11 ± 9.0550 �M 43.31 ± 3.25 46.83 ± 5.22 9.86 ± 2.51*

72 h Control 41.58 ± 1.72 43.13 ± 2.31 15.29 ± 2.4615 �M 32.68 ± 3.60 50.44 ± 7.59 16.89 ± 4.1825 �M 33.15 ± 4.53 44.63 ± 8.94 22.22 ± 13.1935 �M 44.26 ± 4.75 39.58 ± 7.09 16.17 ± 9.77

F

3

3GtCpasvtmcc

3

iopparsdb

TC

F

50 �M 41.57 ± 13.58 39.27 ± 23.58 19.16 ± 14.30

or explanation see Table 4.

.3. Cell cycle

The tested compounds showed a cytotoxic effect, and molecules–5 in particular demonstrated a tendency to accumulate in the1/G0-phase of the cell cycle. In case of the molecule 3, this

rend was dependent on both dose and time over a 72 h period.ompounds 4–5 were also able to induce cell cycle arrest in G1/G0-hase, however only at lower doses and within the initial 24 h (5),lthough higher concentrations of these compounds resulted inome changes which were deemed insignificant due to their highariability (Tables 4–6). Considering the other results which provehe high toxicity levels of these compounds (�� m, viability/cellular

etabolism), we presume that the cytotoxic action was fast andaused rapid cell destruction, given that a high proportion of theell population was exterminated within the first 24 h (Fig. 6).

.4. Dissipation of the mitochondrial membrane potential

The molecules designated as 3–5 demonstrated a strong abil-ty to affect mitochondrial physiology (Fig. 7). The dissipationf mitochondrial membrane potential �� m is the hallmark ofrogrammed cell death as it is accompanied by the release ofroapoptotic molecules into the cytoplasm. Fundamentally, it islso a concomitant result of cell death in general. In concentrationsanging from 35 to 50 �M, compound 4 was able to evoke �� m dis-

ipation in approximately 80% of cells within 24 h and caused celleath with an efficiency approaching 100% after 48 h and 72 h incu-ation. The other two derivatives, 3 and 5, were less effective, but

able 6ell cycle distribution of compound 5.

G1 S G2/M

24 h Control 33.02 ± 0.91 49.59 ± 0.56 17.39 ± 1.4425 �M 38.81 ± 3.32 48.75 ± 2.23 12.43 ± 1.23*50 �M 46.82 ± 2.68* 41.43 ± 2.73 11.75 ± 0.6175 �M 53.14 ± 3.46* 30.07 ± 1.28* 16.79 ± 2.29100 �M 46.43 ± 1.91* 32.61 ± 1.98* 20.96 ± 3.40

48 h Control 32.88 ± 0.45 48.77 ± 1.70 18.35 ± 2.0625 �M 39.79 ± 3.44 47.54 ± 1.66 12.66 ± 1.79*50 �M 57.69 ± 5.70* 31.06 ± 4.61 11.25 ± 1.20*75 �M 45.55 ± 13.48 40.05 ± 15.98 14.40 ± 2.50*100 �M 29.39 ± 14.75 59.84 ± 28.72 10.77 ± 13.97

72 h Control 41.58 ± 1.72 43.13 ± 2.31 15.29 ± 2.4625 �M 41.84 ± 1.70 44.72 ± 0.73 13.44 ± 1.0550 �M 54.68 ± 7.82 34.37 ± 6.24 10.94 ± 1.6075 �M 39.71 ± 20.29 50.22 ± 29.81 15.10 ± 9.58100 �M 50.87 ± 8.62 34.86 ± 4.34 14.27 ± 12.02

or explanation see Table 4. [

logical Macromolecules 64 (2014) 53– 62 61

they also demonstrated high cytotoxic potential at concentrationsof up to 100 �M.

3.5. Viability/cellular metabolism

Simultaneous analysis of viability (propidium iodide) and activ-ity of cellular metabolism (fluorescein diacetate) showed similarresults to those obtained in the analysis of �� m dissipation. Theresults proved that molecules 1 and 2 are ineffective at concentra-tions of less than 100 �M. On the other hand, derivative 4 provedto be highly effective at the same concentration; for this reason,it was only tested at concentrations of up to 50 �M as the higherconcentrations were highly cytotoxic when they were tested after24 h of incubation (Fig. 8A). Compared to compound 4, compounds3 and 5 demonstrated a delayed, although still distinct effect lead-ing to cell destruction. The action of these three compounds wasfound to be dependent on both time and dose (Fig. 8).

4. Conclusion

A series of cholinesterase modulators, compounds 1–5, wasinvestigated. These compounds showed DNA binding activity(K = 0.5 × 104 to 8.9 × 105 M−1). UV–vis, fluorescence, circulardichroism spectroscopy, DNA melting techniques indicated thatthe studied compounds act as effective DNA-interacting agents. Wehave also confirmed their biological activity through experimentsfocused on analyzing their potential anticancer effects. Only deriva-tives 3–5 (cholinesterase inhibitors) proved to be highly effectiveagainst human acute promyelocytic leukaemia cells HL-60 (withderivatives 4 and 5 proving to be especially effective), evoking rapidmitochondrial membrane potential dissipation combined with adrop in cell metabolism and loss of viability at concentrations ofup to 100 �M. Derivative 4 proved to be the most effective as itcaused an almost total eradication of cells at a concentration of50 �M. Based on obtained findings, cholinesterase reactivators 1and 2 were found relatively save from the point of view of DNAbinding, whereas cholinesterase inhibitors 3–5 (especially com-pound 4) resulted as strong DNA binding agents that limit theirplausible use. We suggest that some of the derivatives presentedin this study (compound 4) could have potential applications asanti-cancer drugs.

Acknowledgements

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

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