Bioorganic Chemistry 59 (2015) 168–176

Contents lists available at ScienceDirect

Bioorganic Chemistry journal homepage: www.elsevier.com/locate/bioorg

Tacrine derivatives as dual topoisomerase I and II catalytic inhibitors Jana Janocˇková a, Jana Plšíková a, Ján Koval’ c, Rastislav Jendzˇelovsky´ c, Jaromír Mikeš c, Jana Kašpárková d, Viktor Brabec d, Slávka Hamulˇaková b, Peter Fedorocˇko c, Mária Kozˇurková a,e,⇑ a

Department of Biochemistry, P. J. Šafárik University in Košice, Moyzesova 11, 040 01 Košice, Slovak Republic Department of Organic Chemistry, Institute of Chemistry, P. J. Šafárik University in Košice, Moyzesova 11, 040 01 Košice, Slovak Republic c Department of Cellular Biology, Institute of Biology and Ecology, Faculty of Science, P. J. Šafárik University in Košice, Moyzesova 11, 040 01 Košice, Slovak Republic d Institute of Biophysics, Department of Molecular Biophysics and Pharmacology, Academy of Science of the Czech Republic, Brno, Czech Republic e Biomedical Research Center, University Hospital Hradec Kralove, Hradec Kralove, Czech Republic b

a r t i c l e

i n f o

Article history: Received 29 December 2014 Available online 19 March 2015 Keywords: Tacrine derivatives Topoisomerase I and II Spectroscopic techniques

a b s t r a c t This study examines the binding properties of a series of newly synthetized tacrine derivatives 1–4 and their anticancer effects. Spectroscopic techniques (UV–Vis, fluorescence spectroscopy, thermal denaturation, and linear spectropolarimetry) and viscometry were used to study DNA binding properties and to determine the types of DNA interaction with the studied derivatives. The binding constants for the complexes with DNA were obtained using UV–Vis spectroscopic titrations (K = 1.6  104– 4.0  105 M1) and electrophoretic methods were used to determine the effect of the derivatives on topoisomerase I and II activity. Monotacrine derivative 1 showed evidence of topoisomerase I relaxation activity at a concentration of 30  106 M, while bistacrine derivatives 2–4 produced a complete inhibition of topoisomerase I at a concentration of 5  106 M. The biological activities of the derivatives were studied using MTT-assay and flow cytometric methods (detection of mitochondrial membrane potential and measurement of cell viability) following incubation of 24 and 48 h with human leukemic cancer cell line HL60. The ability of the derivatives to impair cell proliferation was also tested through the analysis of cell cycle distribution. Ó 2015 Elsevier Inc. All rights reserved.

1. Introduction A large percentage of chemotherapeutic anticancer drugs are compounds which either interact directly with DNA or which prevent the proper relaxation of DNA through the inhibition of topoisomerases [1–6]. They are widely occurring enzymes that can solve the topological problems in eukaryotic and prokaryotic cells associated with DNA which emerge during key cellular processes [7,8]. The last few years have seen the development of considerable pharmacological interest in these enzymes. DNA topoisomerase I (Topo I) is an important enzyme which is found in all living organisms and which participates in many metabolic cellular processes, such as replication, transcription, recombination and repair [8–10]. Topo I acts by creating a transient break in one strand of DNA, whereas Topoisomerase II (Topo II) introduces transient doublestrand breaks [11]. Topo II has been the focus of extensive study and is the target of a wide variety of drugs which can be divided ⇑ Corresponding author at: Department of Biochemistry, P. J. Šafárik University in Košice, Moyzesova 11, 040 01 Košice, Slovak Republic and Biomedical Research Center, University Hospital Hradec Kralove, Hradec Kralove, Czech Republic. E-mail address: [email protected] (M. Kozˇurková). http://dx.doi.org/10.1016/j.bioorg.2015.03.002 0045-2068/Ó 2015 Elsevier Inc. All rights reserved.

into two main categories: DNA intercalators and non-intercalators [1,2,4,12–16]. While the majority of topoisomerase inhibitors show selectivity toward either Topo I or Topo II, a small class of compounds can act against both enzymes [17,18]. Epipodophylloids, for example, exhibit a dual inhibitory effect without stabilizing the cleavage complex induced by either enzyme (bis(phenazine-1-carboxamides)) tested against Topo I/II) [19,20]. The most potent compound is a (9-methyl) derivative which poisons Topo I at low drug concentrations and inhibits the catalytic activity of both Topo I/II activity. Topostin has also been shown to inhibit both Topo I and Topo II, although it has no toxicity against tumor cells [21]. The activity of 9-amino-1,2,3,4-tetrahydroacridine (tacrine) in neurological disorders such as Alzheimer’s disease is now well established. Numerous studies have confirmed that the drug is an effective inhibitor of acetylcholinesterase [22–24] and it has also been reported that it is not clastogenic in mammalian cells [25]. Tacrine is a relatively weak catalytic inhibitor of Topo II (in comparison with 9-aminoacridine) which has been found to inhibit topoisomerase and DNA synthesis thereby resulting in mitochondrial DNA depletion and apoptosis [26]. Tetrahydroacridines have

J. Janocˇková et al. / Bioorganic Chemistry 59 (2015) 168–176

an imperfect planarity and have been shown to bind to DNA with much lower levels of affinity than those of acridines [27]. Recent years have seen an increased academic interest in bisintercalators, as many researchers have hypothesized that the incorporation of two intercalating units into a polyfunctional ligand would result in an agent with higher DNA-binding constants, slower dissociation rates, and substantially improved sequence selectivity in comparison to those of a monointercalator. In comparison with monomers, the two planar structures connecting appropriate linker length are considered to be two factors in increasing cancer cell cytotoxicity [28]. In this paper, the biochemical and biological activities of monoand bistacrine derivatives were tested in order to examine their capacity to bind to DNA and to interfere with calf thymus Topo I and human Topo II. 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 on HL60 cell lines. 2. Material and methods 2.1. Tacrine derivatives Novel tacrine derivatives 1–4 were synthetized at Department of Organic Chemistry, P. J. Šafárik University in Košice by Dr. Hamulakova [29,30]. The studied derivatives were dissolved in DMSO at a concentration of 5  103 M. The chemical structures and characteristics of the novel derivatives are shown in Table 1. 2.2. DNA binding experiments 2.2.1. Absorption spectroscopic studies and calculation of binding constant UV–Vis spectroscopic analysis was performed in 0.01 M Tris– HCl buffer (pH = 7.4) with fixed concentrations of the tacrine derivatives (0.1  104 M in the case of ligand 1, 0.2  104 M in the case of ligands 2–4) and with gradually increasing concentrations of calf thymus DNA (ctDNA). The concentrations of ctDNA ranged from 0 to 6.42  107 M. Spectra were measured on

169

Varian Cary 100 UV–Vis spectrophotometer in a quartz cuvette (1 cm path length). Each absorption spectrum was scanned at the wavelength range of 250–500 nm. The binding constants K of derivatives 1–4 with DNA were determined from the absorption titration data using the McGhee and von Hippel equation [31].

 n1 r 1  nr ¼ Kð1  nrÞ Cf 1  ðn  1Þr where K is a binding constant, n is the binding site size in the base pairs, Cf is the molar concentration of free ligands and r is the number of ligand molecules that bind to one mol of nucleotide. The binding data were fitted using Gnu Octave 2.1.73 software [32].

2.2.2. Thermal denaturation Thermal denaturation experiments were performed in a quartz cuvette (1 cm path length) using a Varian Cary Eclipse spectrophotometer equipped with a thermostatic cell holder. The temperature was increased at a rate of 1 °C min1 over a range 40–95 °C. The thermal melting points (Tm, °C) were determined as the maximum of the first derivative plots of the melting curves. The Tm of ctDNA (6.3  107 M) in the presence and absence of the studied tacrine derivatives 1–4 (2.5  105 M) was measured in BPE buffer, pH = 7.1 (6.0  103 M Na2HPO4, 2.0  103 M NaH2PO4, 1.0  103 M EDTA).

2.2.3. Fluorescence studies The fluorescence spectra were measured on Varian Cary Eclipse spectrophotometer in a 10 mm path length quartz cuvette in a 0.01 M Tris–HCl buffer at pH = 7.4 and at laboratory temperature using excitation wavelengths of the respective absorption maxima. Emission spectra were collected from 340 to 700 nm. 10 nm slits were used for excitation and emission beams. Fluorescence intensities are expressed in arbitrary units. Fluorescence spectra were monitored at a fixed concentration of studied tacrine derivatives (0.1  104 M) with increasing concentrations of ctDNA. The concentration of ctDNA ranged from 0 to 6.42  107 M.

Table 1 Structural data of studied tacrine derivatives. Sign

Structure

Name

Molecular weight (g mol1)

Monotacrine derivative

1

N-(Piperazinoethyl)-N-(1,2,3,4-tetrahydroacridin-9-yl)amine

310.44

Bistacrine derivatives

2

1-(1,2,3,4-Tetrahydroaridin-9-yl)-3-[2-(1,2,3,4-tetrahydroacridin-9ylamino)ethyl]thiourea

481.65

3

1-(1,2,3,4-Tetrahydroaridin-9-yl)-3-[4-(1,2,3,4-tetrahydroacridin-9ylamino)butyl]thiourea

509.71

4

1-(1,2,3,4-Tetrahydroacridin-9-yl)-3-[6-(1,2,3,4-tetrahydroacridin-9ylamino) hexyl]thiourea

537.76

J. Janocˇková et al. / Bioorganic Chemistry 59 (2015) 168–176

170

2.2.4. LD spectroscopy Flow LD spectra were collected using a flow Couette cell in a Jasco J-720 spectropolarimeter adapted for LD measurements. Long molecules such as DNA (minimum length of 250 bp) can be orientated in a flow Couette cell. The flow cell consists of a fixed outer cylinder and a rotating solid quartz inner cylinder, separated by a gap of 0.5 mm, giving a total path length of 1 mm. LD spectra of ctDNA at a concentration of 3.11  104 M modified by tacrine derivatives (c = 0–1.04  104 M) were recorded at 25 °C in a 0.01 M Tris–HCl buffer in the range of 210–600 nm. 2.2.5. Viscometry The dynamic viscosity and flow times of the solution containing ctDNA (3.0  104 M) and studied derivatives (0–5.0  105 M) were measured using microviscometry (AMVn Automated MicroViscometer, Anton Paar GmbH, Austria) in a 1.6 mm capillary tube at 37 °C with 0.01 M Tris–HCl buffer. Viscosity measurements were obtained in triplicate and an average of the viscometry data was taken. The specific viscosity of the DNA solution with added ligand (g) was calculated using the flow times buffer (t0) and buffer with DNA in presence ligand (t1) using equation.

g¼

ðt1  t0 Þ t0

The root of the relative viscosity (g/g0)1/3 is plotted as a function of the moles of ligand bound per DNA-base pair (r) [33]. 2.2.6. Topoisomerase I relaxation study Negatively supercoiled plasmid pUC19 (1.4 lg) was used to assess the effects of derivatives 1–4 on topoisomerase-mediated DNA relaxation. The plasmid was incubated for 45 min at 37 °C with 2 units of calf thymus topoisomerase I (Takara Japan) in the presence and in the absence of the studied derivatives (1: 5, 30 and 60  106 M; 2–4: 1, 3, 5, 8 and 30  106 M, respectively). DNA – topoisomerase reaction buffer contains 3.5  101 M Tris– HCl (pH = 8.0), 7.2  101 M KCl, 5.0  102 M MgCl2, 2 2 5.0  10 M DTT, 5.0  10 M spermicide and 0.1% bovine serum albumin (BSA). Gel electrophoresis was performed at 7 V/cm for 2 h in a TBE buffer (8.9  102 M Tris, 8.9  102 M H3BO3, 2.0  103 M EDTA) on 0.8% agarose gel which was stained with ethidium bromide (1 mg mL1). DNA bands were visualized with UV-light. 2.2.7. Topoisomerase II decatenation assay Reactions contained 0.16 lg kinetoplast DNA (kDNA), the fresh assay buffer (mixed 10  Topo II incomplete assay Buffer A – 0.5 M Tris–HCl (pH = 8.0), 1.50 M NaCl, 0.1 M MgCl2, 5.0  103 M Dithiothreitol, 300 lg BSA/mL and 10  ATP Buffer B – 2.0  102 M ATP in water to make a 5  complete assay buffer), 2 units of human topoisomerase IIa and various concentrations of the tested compounds (5, 50 and 100  106 M) in a 20 lL final volume. These reaction mixtures were incubated for 45 min at 37 °C. The reactions were terminated with 4 lL of 5 stop loading buffer (5% Sarkosyl, 0.12% bromophenol blue, 25% glycerol) and were loaded directly onto a 1% agarose gel at 7 V/cm in TAE buffer for 2.5 h. The gel was stained with ethidium bromide and destained in water before being photographed under UV light. Known topoisomerase II poisons (100  106 M m-AMSA and 100  106 M ellipticine) were used as positive controls in the experiment. 2.2.8. Cleavage assay of topoisomerase II Reactions contained 0.2 lg pHOT1 supercoiled DNA, the assay buffer (mixed 10  Topo II incomplete assay Buffer A – 0.5 M Tris–HCl (pH = 8), 1.5 M NaCl, 0.1 M MgCl2, 5.0  103 M

dithiothreitol, 300 lg BSA/mL and 10  ATP Buffer B – 2.0  102 M ATP in water to make a 5  complete assay buffer), 5 units of human topoisomerase IIa and the test compounds (100  106 M) in a 20 lL final volume. This reaction mixtures were incubated at 37 °C for 30 min. Incubation was stopped through the addition of 10% SDS, and the reaction was then further incubated with proteinase K (50 lg mL1) for 20 min at 37 °C. The samples were then cleansed through extraction by using a mixture of chloroform and isoamyl alcohol (24:1) and run on a 1% agarose gel containing ethidium bromide (0.5 lg mL1) at 7 V/cm in TAE buffer. The poisoning activity of the tested compound was determined by the presence or absence of linear DNA forms. The known topoisomerase II poison m-AMSA (100  106 M) was used as a positive control in the experiment. 2.2.9. Cell culture and experimental design HL-60 cells (human promyelocytic leukemia, ATCC, USA) were grown in a complete RPMI medium (Gibco, USA) supplemented with 7.5% NaHCO3 (10 mL L1), penicillin (100 U mL1), streptomycin (100 lg mL1), amphotericin (25 lg mL1, Invitrogen, USA), 10% heat-inactivated fetal calf serum (FCS, PAA Laboratories GmbH, Austria) and maintained at 37 °C, 95% humidity and in a 5% CO2 atmosphere. For experiments, the cells were seeded in £60 mm Petri dishes (TPP, Switzerland) to which the studied chemical compounds were added. The results were analyzed 24 and 48 h after the addition of the compounds. 2.2.10. Detection of mitochondrial membrane potential (MMP) Cells were treated for 24 or 48 h and subsequently harvested by centrifugation, washed once with Hank’s balanced salt solution (HBSS), resuspended in HBSS supplemented with tetramethylrhodamine ethyl ester perchlorate (TMRE; 0.1  106 M/20 min/RT, Sigma–Aldrich, St. Louis, MO, USA), incubated in darkness and analyzed using a BD FACSCalibur flow cytometer (Becton Dickinson, San Jose, USA). Results are presented as a means ± standard deviation (SD) of three independent experiments. 2.2.11. Metabolic activity and viability Cells were treated for 24 or 48 h and subsequently harvested by centrifugation, washed once with HBSS, resuspended in HBSS supplemented with fluorescein diacetate (FDA) (100 ng mL1), propidium iodide (PI) (25 lg mL1) (both Sigma–Aldrich, USA), incubated in darkness for 20 (FDA) and 5 min (PI) at RT and analyzed using a flow cytometer. Results are presented as a mean ± SD of three independent experiments. 2.2.12. Analysis of cell cycle parameters Cells were treated for 24 or 48 h and subsequently harvested by centrifugation, washed in cold phosphate-buffered saline (PBS), fixed in cold 70% ethanol and kept at 4 °C overnight. Prior to analysis, cells were washed twice in PBS, resuspended in staining solution (0.1% Triton X-100, 0.137 mg mL1 of ribonuclease A and 0.02 mg mL1 of PI), incubated in darkness for 30 min at RT and analyzed using a flow cytometer. ModFit 3.0 (Verity Software House, Topsham, USA) software was used to generate DNA content frequency histograms and to quantify the number of cells in the individual cell cycle phases. Results are presented as a mean ± SD of three independent experiments. 2.2.13. Statistical analysis Results were calculated as a mean ± SD of at least three independent experiments. Statistical significance was determined with t-tests and results were deemed significant if p < 0.05.

J. Janocˇková et al. / Bioorganic Chemistry 59 (2015) 168–176

3. Results and discussion 3.1. Absorption measurement UV–Vis spectroscopy is a useful and efficient technique for identifying the binding potency of small molecules with DNA and of determining their modes of interaction [1,34]. The changes occurring in derivatives 1–4 following the addition of ctDNA were studied using this technique. The absorption spectra of mono- (1) and bistacrine (2–4) derivatives display a significant absorption band in the region of 250–400 nm. The absorption spectra of 1–4 in both the absence and presence of calf thymus DNA are given in Fig. 1. The data reveals a negligible decrease in maximal intensities (hypochromism 17–34%) and a mild bathochromic shift to higher wavelengths. It is widely accepted that hypochromism and bathochromic shifts are often associated with the intercalation mode of DNA interaction with low molecular weight compounds as a result of strong stacking interactions between DNA base pairs and the aromatic chromophore of the ligand. The scale of hypochromism is usually dependent upon the strength of the intercalative interaction of the ligand with DNA [35]. The hypochromism and bathochromic shift from our results indicate that all of the studied tacrine derivatives show evidence of interaction with ctDNA. The spectral absorption results which are characteristic of the studied tacrine derivatives is given in Table 2. The absence of a red strong shift in the UV–Vis spectra analysis suggests that the binding mode is unlikely to be intercalative in form. The binding constants K of ligand complexes 1–4 with ctDNA were calculated

3.2. Thermodynamic parameters The denaturation of the polynucleotide strand from double strand DNA results in absorption hypochromism at the wavelength of 260 nm. Tm is dependent on the strength and mode of its interaction with DNA; sharp increases in Tm of DNA are generally indicative of intercalation, while groove binding along the DNA phosphate backbone usually leads to smaller changes in Tm [42]. Table 2 and Fig. 2 show that the initial Tm of ctDNA was 80 °C but that the values change following the addition of compounds

0,6

3

0.26 0.24 0.22 0.2 0 10 20 30 40 50 60 70 Amount of added ctDNA [µl]

0,2

0

0,4

0,2

Absorbance (by λmax = 325 nm)

Absorbance

1

using McGhee and von Hippel plots in order to compare the affinity of tacrine derivatives to DNA. The range of binding constants was found to be between 1.6  104 and 2.7  105 M1, findings which prove the low affinity of the tacrine ligands to DNA and which also correspond to typical binding constants for groove binding complexes (105–109) [36,37]. The relative binding constants K calculated from our results increased as follows: 4 < 2 < 3 < 1. Comparing the values of binding constants for our studied monoand bistacrine derivatives, we found that the binding constants of monotactine (2.7  105 M1) are one order higher than those than those of the bistacrine (1.6–6.8  104 M1). The results are also in the reverse order of those of the acridine mono- and bisacridine derivatives, in which it had been discovered that the binding constants decreased with increasing substituent length [38–41]. It is possible to suggest that the different values in comparison to those of the acridine–DNA complexes are caused by the distinct geometry of the tacrine–DNA complexes, and by a different type of binding with DNA.

Absorbance

Absorbance (by λmax=337 nm)

0,4

0.65 0.55 0.45 0.35 0

350

20

400

350

0,6 0.4

4

0.35 0.3 0

20

40

60

0,2

0,4

0,2

0

0 350

Wavelength [nm]

400

Absorbance (by λmax = 325 nm)

0.25

Amount of added ctDNA [μl]

300

400

Wavelength [nm]

Absorbance

Absorbance

0,4

(by λmax = 324 nm)

Absorbance

2

60

300

Wavelength [nm] 0,6

40

Amount of added ctDNA [µl]

0 300

171

0.6 0.5 0.4 0.3

0

20

40

60

Amount of added ctDNA [µl]

300

350

400

Wavelength [nm]

Fig. 1. UV–Vis titration absorption spectra of studied monotacrines (1; concentration 0.1  104 M), and bistacrine (2–4; concentration 0.2  104 M) derivatives in 0.01 M Tris–HCl buffer (pH = 7.4; 24 °C) derivatives with increasing concentrations of ctDNA.

J. Janocˇková et al. / Bioorganic Chemistry 59 (2015) 168–176

172

Table 2 DNA binding properties of studied tacrine derivatives 1–4. Compound

1 2 3 4

kmax Free

Bound

335 324 325 324

337 325 332 325

Bathochromic shift (nm)

Hypochromicity (%)

a

2 1 7 1

17.03 20.77 30.72 34.80

85.11 79.08 79.08 78.15

Tm (°C)

DTm (°C)

K (M1)

DG (kcal mol1)

5.11 0.92 0.92 1.85

2.7  105 3.4  104 6.8  104 1.6  104

28.40 28.92 30.49 27.21

a Tm measurements were performed in BPE buffer, pH = 7.1 (6.0  103 M Na2HPO4, 2.0  103 M NaH2PO4, 1.0  103 M EDTA) using derivatives 1–4 (2.5  105 M) and ctDNA (6.3  107 M) with a heating rate of 1 °C/min. Tm of ctDNA for measurement was 80 °C.

800

∆A/∆T

1

ctDNA (6.3x10-7 M)

0,02

ctDNA + 1 (2.47x10 -5 M)

Intensity [a.u.]

ctDNA + 2 (2.47x10 -5 M) ctDNA + 3 (2.47x10 -5 M)

0,01

ctDNA + 4 (2.47x10 -5 M)

600

400

200

0 40

60

80

Temperature [°C] 0 400

Fig. 2. First derivative of denaturation curves of ctDNA (black lines, concentration for ctDNA was 6.3  107 M) with studied derivatives 1–4 (color lines, 2.47  105 M) measured at 260 nm in BPE buffer, pH = 7.1.

1–4, ranging from 78 to 85 °C. These results are indicative of low helix stability and therefore of the existence of groove binding between the studied derivatives and DNA. The stronger interaction between derivative 1, the only single structure molecule of all of the studied compounds, and DNA caused the greatest change in DNA Tm, confirming a higher level of helix stability than was found in the interactions between DNA and compounds 3–4. 3.3. Fluorescence spectroscopy Fluorescence quenching is a commonly technique which is used to determine the way in which small molecules bind to the nucleic acid structure. Fluorescence spectroscopy has a number of advantages over other techniques, such as higher selectivity, sensitivity and a larger linear concentration range. The most intense fluorescence is observed in compounds including aromatic groups with low energy p–p⁄ transition levels [43]. The fluorophore which typically binds to DNA by intercalation is ethidium bromide or acridine. In the presence of ctDNA its fluorescence decreases due to strong intercalation into adjacent base pairs [44]. Derivative 1 showed the highest intensity of all of the studied samples. Fig. 3 and SP1 show the fluorescence emission spectra of compounds 2–4 in both the absence and presence of ctDNA. The binding parameters from fluorescence measurement are shown in Table 3. The fluorescence intensity strongly decreased following the addition of ctDNA only for sample 1. 3.4. LD spectroscopy Long polymers such as DNA can be oriented by the viscous drag created by the rotation of one cylinder inside another. The extent of orientation can be assessed using linear dichroism (LD, the difference in absorption of light polarized parallel and perpendicular to an orientation axis) [45,46]. The orientation of B-DNA base pairs is approximately perpendicular to the DNA helix axis [47], which is also the orientation axis, and thus the base p–p⁄ transitions give a

450

500

550

Wavelength [nm] Fig. 3. Fluorescence emission spectra of studied tacrine derivative 1 (concentration 0.1  104 M) in 0.01 M Tris–HCl buffer (pH = 7.4; 24 °C) with increasing concentrations of ctDNA.

Table 3 Fluorescence characteristic of studied tacrine derivatives 1–4. Compound

kex (nm)

kem (nm)

Imax

a

1 2 3 4

335 324 325 324

476.85 364.00 371.07 363.07

567.38 166.77 423.29 468.80

1 0.29 0.74 0.82

F/F0

a Fluorescence quantum yields were calculated using compound 1 as standard (Uf = 1).

negative LD signal. If a molecule is bound to DNA in a specific orientation, it will also be oriented by the viscous drag and exhibit LD signals as a result. If the molecule is free/unbound or bound in a random orientation, then no LD signal is observed for its transitions. All five of the studied tacrine derivatives produced LD signals during spectroscopy with DNA (Fig. 4), a result which confirms that their DNA-binding is not random. The LD signal yielded by the sample of ctDNA in the presence of monotacrine derivative 1 showed a decrease in the region of above 300 nm (in the range of 310–370 nm); the LD spectra (Fig. 4A) show that this tacrine derivative bind to DNA in specific, non-random orientations. Moreover, the negative sign of the LD signal which arose in the region of 310–370 nm suggests that the angle of the long axis of tacrine derivative 1 to the axis of the DNA double helix is greater than 54°, the angle which is characteristic for an intercalator [48]. The DNA LD bands (220–300 nm) confirm that the DNA in the presence of tacrine derivative 1 remains in the B– DNA conformation; however, some structural changes in DNA are suggested by the increase in the amplitude of the DNA negative LD band at 260 nm upon the addition of tacrine derivative 1 (Fig. 4A). An increase in the amplitude of the negative 260 nm LD band of DNA is usually associated with DNA stiffening [47–49],

J. Janocˇková et al. / Bioorganic Chemistry 59 (2015) 168–176

173

Fig. 4. Linear dichroism spectra of ctDNA (3.11  104 M, black line) in the presence of increasing amounts of studied tacrine derivatives 1–4 (c = 0–1.04  104 M) in 0.01 M Tris–HCl solution (pH = 7.4).

which suggests that the effect of tacrine derivative 1 on this DNA LD signal is consistent with intercalation of the tacrine ligands. In addition, compound 1 also caused a significant shift (cca. 10 nm) of the main DNA band near 260 nm. The wavelength shifts in the region of DNA absorption for tacrine derivative 1 are also consistent with intercalation. Notably, in contrast to the monotacrine compound 1, the DNA LD signal arising from the bases (at 260 nm) was significantly reduced following the addition of bistacrine derivatives 2–4 (Fig. 4B–D). This could either be a result of an increase in DNA flexibility or a shortening of the DNA by bending, compaction, or aggregation [50]. The LD bands at 260 nm (Fig. 4B–D) confirm that the double-stranded B-DNA structure is also retained in the presence of compounds 2–4 and that the local structure is not significantly perturbed. Importantly, bistacrine derivatives 2–4 caused no shift of the main DNA LD band near 260 nm, which further corroborates that the preferred mode of DNA binding of the bistacrine derivatives is nonintercalative.

Fig. 5. The relative specific viscosity of ctDNA (3.0  104 M) in the presence of derivatives 1–4 in 0.01 M Tris–HCl buffer at 37 °C. Data are presented as (g/g0)1/3 versus [ligand]/[DNA], where g is the viscosity of DNA in the presence of ligand and g0 is the viscosity of DNA alone. DNA concentration is expressed as a concentration of base pairs.

3.5. Viscometry The reactions of intercalators with DNA result in the unwinding and lengthening of DNA. As the amount of intercalator bound to DNA increases, so too does the viscosity of DNA-containing solution. Therefore viscosity measurements are a useful and very sensitive method of measuring changes in DNA length. It is also known that groove binders, which do not lengthen the DNA helix, have no effect on the viscosity of DNA solution [51,52]. The effect of the studied tacrine derivatives on the viscosity of ctDNA is described in Fig. 5. Our viscosity results (Fig. 5) show that compound 1 caused a significant increase in the viscosity of the DNA solution. This result is strong evidence of DNA lengthening

Fig. 6. Electrophoresis agarose gels showing inhibition of calf thymus topoisomerase I induced DNA relaxation by compounds 1. Supercoiled DNA (lane pUC19) was incubated at 37 °C for 45 min. with topoisomerase I in the absence (lane T) or presence of ligands (lane a – 5  106 M, lane b – 30  106 M, lane c – 60  106 M). Lane CPT/EtBr  pUC19 + Topo I + camptothecin (10  106 M)/ ethidium bromide (10  106 M).

174

J. Janocˇková et al. / Bioorganic Chemistry 59 (2015) 168–176

and by extension the intercalation of 1, a finding which matches those inferred from the LD spectra results (Fig. 5). In contrast, bistacrine compounds 2–4 caused a considerably less significant increase in the viscosity of the DNA solution, indicating a less significant degree of orientation of compounds 2–4 interacting with DNA. These results indicate that the presence of two tacrine moieties in the bistacrine molecules reduces the intercalative ability into DNA, another finding which is consistent with the results of LD spectra (Fig. 4B–D). 3.6. Topoisomerase I relaxation assay DNA topoisomerases are essential enzymes that play an important role in DNA transcription and replication as well as in chromosome segregation and recombination [2,4,5,53]. Topoisomerases

cut DNA transiently to allow the unwinding of the supercoiled plasmid into both partially relaxed forms and the fully relaxed form [26]. A primary reaction of Topo I is the relaxation of supercoiled DNA which has a different electrophoretic mobility than that of a completely relaxed (not supercoiled) DNA [54]. Like other acridines, tacrine has a dose-dependent effect on topoisomerase mediated DNA cleavage and resealing [55]. The novel tacrine derivatives were studied at increasing concentrations from 1 to 60  106 M in order to determine their ability to inhibit the relaxation ability of Topo I in the presence negatively supercoiled plasmid pUC19. Monotacrine derivative 1 demonstrated topoisomerase I relaxation activity at a concentration of 30  106 M. Bistacrine derivatives 2–4 totally inhibited topoisomerase I at a concentration of 5  106 M. These results are shown in Figs. 6 and 7.

Fig. 7. Electrophoresis agarose gels showing inhibition of calf thymus topoisomerase I induced DNA relaxation by compounds 2–4. Supercoiled DNA (lane pUC19) was incubated at 37 °C for 45 min with topoisomerase I in the absence (line T) or presence of ligands (line a – 1  106 M, lane b – 3  106 M, line c – 5  106 M, line d – 8  106 M, line e – 30  106 M). Lane CPT  pUC19 + Topo I + camptothecin (10  106 M).

Fig. 8. Topoisomerase II inhibition assays – effect of studied compounds 1–4 (5–100  106 M) on the decatenation of kDNA (0.16 lg) by 2U of human topoisomerase II.

J. Janocˇková et al. / Bioorganic Chemistry 59 (2015) 168–176

3.7. Decatenation of topoisomerase II The specific reaction of Topo II with DNA is a decatenation reaction as it involves the simultaneous cleavage of two strands of the DNA-helix. The ability of derivatives 1–4 to inhibit the Topo II catalyzed decatenation of kinetoplast DNA (kDNA) was also studied. The appearance of kDNA monomers was monitored in the reaction in order to determine whether an open circular decatenated (nicked) kDNA form or a closed circular decatenated kDNA form had developed. If Topo II retains its normal function, catenated kDNA (top band with lowest mobility) would disappear and bands for open circular intermediate ability) and closed circular decatenated kDNA (migrates furthest) would appear. In contrast the absence of these bands in the results indicates inhibition of the enzyme. As is shown in Fig. 8, inhibitory effects were observed for compound 1 at concentrations of 50 and 100  106 M. Bistacrine derivatives 2–4 were found to inhibit Topo II at a concentration of 100  106 M. Topoisomerase inhibitors are classified as either poisons or suppressors based on the mode in which they interfere with the action of these enzymes. Catalytic inhibitors may prevent the binding of DNA to topoisomerase and stabilize noncovalent DNA–topoisomerase complexes. Topoisomerase poisons stabilize covalent DNA– topoisomerase complexes, thereby disabling the enzyme. After the resolution of DNA species through gel electrophoresis, the presence or absence of a linear DNA band can classify the inhibitor as a Topo II poison [1]. As is shown in Fig. 9, no linear DNA band was observed at 100  106 M of compounds, which indicates that the novel derivatives are topoisomerase catalytic inhibitors rather than poisons. A linear DNA band was clearly visible in the positive control m-AMSA (100  106 M), a compound which is a known topoisomerase II poison. 3.8. Biological studies The biological activity of the tested compounds was assessed 24 and 48 h after treatment of cancer cells, using several different concentrations (1–50  106 M) of the derivatives and applying various techniques (changes in mitochondrial membrane potential, changes in metabolic activity/viability and cell cycle distribution). Tacrine was used as a positive control in these studies, but it did not show any evidence of biological activity in our experiments. The results from the MMP analysis clearly show a negligible alteration of mitochondrial physiology and weak cytotoxic effect on all of the tested derivatives. There was no evidence of any influence being exerted on mitochondrial membrane potential (Fig. SP2). In order to address the effect of the derivatives on cell metabolic activity and viability, a double staining of the cells with FDA and PI was also carried out. The data are presented as the percentage of live and dead cells after treatment. The changes in viability show

175

Table 4 IC50 values for the MMP and viability 24 and 48 h after treatment of studied derivatives 2–4. TMRE IC50 (lM)

24 (h)

48 (h)

PI IC50 (lM)

24 (h)

48 (h)

2 3 4 T

37.51 22.86 19.46 n.d.

36.90 17.62 11.79 n.d.

2 3 4 T

33.02 26.20 21.34 n.d.

37.02 17.66 10.77 n.d.

TMRE – tetramethylrhodamine ethyl ester perchlorate, PI – propidium iodide.

Table 5 The effect of the tested compounds 1–4 (5  106 M) on cell cycle distribution. 24 h

G1

S

G2

C T 2 3 4

38.14 ± 1.82 38.28 ± 3.23 40.26 ± 0.91 43.20 ± 0.30* 45.04 ± 0.31*

43.72 ± 1.14 42.60 ± 1.53 44.49 ± 1.94 42.13 ± 0.92 41.51 ± 0.74

18.13 ± 0.91 19.11 ± 1.69 15.24 ± 1.12 14.66 ± 0.69 13.43 ± 0.86

48 h

G1

S

G2

C T 2 3 4

38.08 ± 1.39 35.45 ± 2.36 41.85 ± 1.87 43.32 ± 1.65 47.09 ± 5.03

44.85 ± 1.77 44.21 ± 2.04 44.72 ± 1.60 44.68 ± 1.79 40.87 ± 4.26

17.06 ± 1.75 18.13 ± 0.47 13.43 ± 0.28 11.99 ± 1.05 12.03 ± 0.90

The results are presented as a mean ± SD. * Statistical significance: p < 0.05 for particular experimental group compared to untreated control (C – control, T – tacrine).

a very similar trend to the changes observed in the MMP analysis (Figs. SP3 and 4), and confirm the effectiveness of the four derivatives in killing cancer cells (Table 4). As an extrapolation of the data presented in the graphs at one specific concentration (5  106 M), a comparison of the calculated IC50 values for both techniques and time points was also prepared (Table 4). Table clearly shows that both methodologies are consistent when addressing the IC50 values and the alignment of the derivatives in an order based on their cytotoxic effects on cancer cells (T < 2 < 3 < 4). The ability of derivatives to impair cell proliferation was tested through an analysis of cell cycle distribution (Table 5). The derivatives (2–4) and tacrine itself, which had been found to be ineffective in MMP testing and viability assay, did not alter cell cycle distribution significantly. The results are graphically presented as a comparison of all of the tested compounds at one chosen concentration 5  106 M (with the exception of the untreated control) due to the large amount of experimental data and in order to allow a better visualization of the differences between the derivatives. Despite the high concentrations used, tacrine itself did not show any effect on the tested cell line (HL60), although other studies have demonstrated that high concentrations of tacrine can induce apoptosis [56] and mitochondrial membrane depolarization [57] in the HepG2 cell line. 4. Conclusion

Fig. 9. Topoisomerase II inhibition mode assay – effect of tacrine derivatives 1–4 (100  106 M) on the growth of DNA – Topo II cleavage complex. pHOT1 DNA (0.2 lg) was incubated with 5U of human Topo II in the presence of the studied derivatives.

A series of tacrine derivatives, compounds 1–4, was investigated. These compounds showed evidence of DNA binding activity (K = 1.6  104–2.7  105 M1). The results of UV–vis, fluorescence spectroscopy, and linear dichroism spectropolarimetry, DNA melting techniques and viscometry indicate that the studied compounds act as effective DNA-groove binder agents. The studied compounds were also investigated for their effect on both Topo I (at 5  106 M) and II (at 100  106 M) activity. The biological activities of derivatives were studied using MTTassay and flow cytometric methods (detection of mitochondrial membrane potential, measurement of cell viability) after 24 and

J. Janocˇková et al. / Bioorganic Chemistry 59 (2015) 168–176

176

48 h incubation with human leukemic cancer cell line HL60. The ability of the derivatives to impair cell proliferation was tested through an analysis of cell cycle distribution. Tacrine and their derivatives can only induce apoptosis and mitochondrial membrane depolarization in the HL60 cell line at very high concentrations. Acknowledgments This study was supported by Slovak Research and Development Agency under contract VVCE-0001-07, VEGA Grant No. 1/0001/13, Internal Grant Program of the P. J. Šafárik University in Košice (VVGS-PF-2014-435) and CZ-DRO (UHHK, 00179906). The authors from the Institute of Biophysics, ASCR in Brno acknowledge support from the Ministry of Education, Youth and Sports of the Czech Republic (Grant LD14019) and acknowledge that their participation in the EU COST Action CM1105 enabled them to exchange regularly the most recent ideas in the field of compounds of biological significance with several European colleagues. Appendix A. Supplementary material Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.bioorg.2015.03. 002. References [1] [2] [3] [4] [5]

[6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18] [19] [20] [21]

R. Palchaudhuri, P.J. Hergenrother, Curr. Opin. Biotechnol. 18 (2007) 497–503. D.S. Thakur, Int. J. Pharm. Sci. Nanotechnol. 3 (2011) 1173–1181. C. Bailly, Chem. Rev. 112 (2012) 3611–3640. M.K. Chan, N.A. Fadzil, A.L. Chew, B.Y. Khoo, Electron. J. Biotechnol. 16 (2013) 1–10. E.A. Lafayette, S.V. De Almeida, M.G. Da Rocha Pitta, E.I.C. Beltrao, T.G. Da Silva, R.O. De Moura, I. Da Rocha Pitta, L.B. De Carvalho Júnior, Please check the author groups in Ref.[5], and correct if necessary.M.D.C. A De Lima, Molecules 18 (2013) 15035–15050. Y. Pommier, E. Leo, H.L. Zhang, C. Merchand, Chem. Biol. 17 (2010) 421–433. F. Cortes, N.P. Pastor, S. Mateos, I. Domminguez, Mutat. Res. 543 (2003) 59–66. L. Baranollo, F. Kouzino, D. Lovons, Transcription 5 (2013) 232–237. M.L. Rothenberg, Ann. Oncol. 8 (1997) 837–855. S. Bendetz-Nezev, A. Gazit, E. Priel, Mol. Pharmacol. 66 (2004) 627–634. J.M. Berger, S.J. Gamblin, S.C. Harrison, J.C. Wang, Nature 379 (1996) 25–232. S. Zhang, X. Li, F. Zhang, P. Yang, X. Gao, Q. Song, Bioorg. Med. Chem. 14 (2006) 3888–3895. A.C. Ketron, W.A. Denny, D.E. Graves, N. Osteroff, Biochemistry 51 (2012) 1730–1739. G. Cholewinski, K. Dzierbicka, A.M. Kolodziejcik, Pharmacol. Rep. 63 (2011) 305–336. S.M. Vos, E.M. Tretter, B.H. Schmidt, J.M. Berger, Nat. Rev. 12 (2011) 827–841. J. Plsikova, J. Stepankova, J. Kasparkova, V. Brabec, M. Backor, M. Kozurkova, Toxicol. In Vitro 28 (2014) 182–186. W.A. Denny, B.C. Baguley, Curr. Top. Med. Chem. 3 (2003) 339–353. J.A. Spicer, S.A. Gamage, G.J. Atwell, G.J. Finlay, B.C. Baguley, W.A. Denny, J. Med. Chem. 40 (1997) 1919–1929. D. Perrin, B. Van Hille, J.M. Barret, A. Kruczynski, C. Etiévant, T. Imbert, B.T. Hill, Biochem. Pharmacol. 59 (2000) 807–819. J.A. Spicer, S.A. Gamage, G.W. Rewcastle, G.J. Finlay, D.J. Bridewell, B.C. Baguley, W.A. Denny, J. Med. Chem. 43 (2000) 1350–1358. C. Bailly, Curr. Med. Chem. 7 (2000) 39–58.

[22] M. Kozurkova, P. Kristian, in: P. Kristian (Ed.), Acridine Isothiocyanates: Chemistry and Biology, Lambert Academic Press, 2014, pp. 206–233 (Chapter 13). [23] M. Kozurkova, S. Hamulakova, Z. Gazova, H. Paulikova, P. Kristian, Pharmaceuticals 4 (2011) 382–418. [24] V. Tumiatti, A. Minarini, M.L. Bolognesi, A. Milelli, M. Rosini, C. Melchiorre, Curr. Med. Chem. 17 (2010) 1825–1838. [25] R.D. Snyder, M.R. Arone, Mutat. Res. 503 (2002) 21–35. [26] A. Mansouri, D. Haouzi, V. Descatoire, C.H. Demeilliers, A. Sutton, N. Vadrot, B. Fromenty, G. Feldmann, D. Pessayre, A.M.D. Berson, Hepatology 38 (2003) 715–725. [27] A.D. Papaphilis, Y.H. Shaw, Biochim. Biophys. Acta 476 (1977) 122–130. [28] S.S. Wang, Y.-J. Lee, S.-C. Hsu, H.-O. Chang, W.-K. Yin, L.-S. Chang, S.-Y. Chou, Bioorg. Med. Chem. 15 (2007) 735–748. [29] S. Hamulakova, J. Imrich, L. Janovec, P. Kristian, I. Danihel, O. Holas, M. Pohanka, S. Böhm, M. Kozurkova, K. Kuca, Int. J. Biol. Macromol. 70 (2014) 435–439. [30] S. Hamulakova, P. Kristian, D. Jun, K. Kuca, J. Imrich, I. Danihel, S. Bohm, K.D. Klika, Heterocycles 76 (2008) 1219–1235. [31] J.D. McGhee, P.H. von Hippel, J. Mol. Biol. 86 (1974) 469–489. [32] J. Busa, Octave, Technical University in Košice, Košice, 2006. [33] P.C. Dedon, Current Protocols in Nucleic Acid Chemistry, John Wiley & Sons Inc., Hoboken, NJ, 2001. [34] C. Liu, S. Liu, Y. Wang, S. Wang, J. Zhang, S. Li, X. Qin, X. Li, K. Wang, Q. Zhou, Med. Chem. Res. 23 (2014) 1899–1907. [35] H. Wu, J. Yuan, G. Pan, Y. Zhang, X. Wang, F. Shi, X. Fan, J. Photochem. Photobiol. B 122 (2013) 37–44. [36] P. Pandya, M. Islam, G.S. Kumar, B. Jayaram, S. Kumar, J. Chem. Sci. 122 (2010) 247–257. [37] H. Ihmels, D. Otto, Top. Curr. Chem. 258 (2005) 161–204. [38] L. Janovec, D. Sabolova, M. Kozurkova, H. Paulikova, P. Kristian, J. Ungvarsky, E. Moravcikova, M. Bajdichova, D. Podhradsky, J. Imrich, Bioconjug. Chem. 18 (2007) 93–100. [39] M. Kozurkova, D. Sabolova, L. Janovec, J. Mikes, J. Koval Ungvarsky, M. Stefanisinova, P. Fedorocko, P. Kristian, J. Imrich, Bioorg. Med. Chem. 16 (2008) 3976–3984. [40] L. Janovec, M. Kozurkova, D. Sabolova, J. Ungvarsky, H. Paulikova, J. Plsikova, Z. Vantova, J. Imrich, Bioorg. Med. Chem. 19 (2011) 1790–1801. [41] J. Plsikova, L. Janovec, J. Koval, J. Ungvarsky, J. Mikes, R. Jendzelovsky, P. Fedorocko, J. Imrich, P. Kristian, J. Kasparkova, V. Brabec, M. Kozurkova, Eur. J. Med. Chem. 57 (2012) 283–295. [42] M. Endang, T. Wahyuni, H. Tjahjono, N. Yoshioka, H. Inoue, Spectrochim. Acta Part A Mol. Biomol. Spectrosc. 77 (2010) 528–534. [43] J.R. Lakowicz, Principles of Fluorescence Spectroscopy, third ed., Springer, 2006. [44] N. Li, Y. Ma, L. Guo, X. Yang, Biophys. Chem. 116 (2005) 199–205. [45] B. Norden, M. Kubista, T. Kurucsev, Quart. Rev. Biophys. 25 (1992) 51–170. [46] A. Rodger, B. Norden, Circular Dichroism and Linear Dichroism, Oxford University Press, Oxford, New York, Tokyo, 1997. [47] P.J. Chou, W.C. Johnson, J. Am. Chem. Soc. 115 (1993) 1205–1214. [48] A. Rodger, R. Marington, M.A. Geeves, M. Hicks, L. de Alwis, D.J. Halsall, T.R. Dafforn, Phys. Chem. Chem. Phys. 8 (2006) 3161–3171. [49] T. Ihara, T. Ikegami, T. Fujii, Y. Kitamura, S. Sueda, M. Takagi, A. Jyo, J. Inorg. Biochem. 100 (2006) 1744–1754. [50] M.J. Hannon, V. Moreno, M.J. Prieto, E. Moldrheim, E. Sletten, I. Meistermann, C.J. Isaac, K.J. Sanders, A. Rodger, Angew. Chem. Int. Ed. 40 (2001) 879–884. [51] S. Satyanarayana, J.C. Dabrowiak, J.B. Chaires, Biochemistry 31 (1992) 9319– 9324. [52] S. Satyanarayana, J.C. Dabrowiak, J.B. Chaires, Biochemistry 32 (1993) 2573– 2584. [53] J. Leppard, J. Champoux, Chromosoma 114 (2005) 75–85. [54] J.L. Nitiss, E. Soans, A. Rogojina, A. Seth, M. Mishina, Curr. Protoc. Pharmacol. (2012) (Chapter 3). [55] A. Skaladanowski, S.Y. Plisov, J. Konopa, A.K. Larsen, Mol. Pharmacol. 49 (1996) 772–780. [56] C. Gao, Y. Ding, L. Zhong, L. Jiang, C. Geng, X. Yao, J. Cao, Toxicol. In Vitro 28 (2014) 667–674. [57] M.J. Ezoulin, C.Z. Dong, Z. Liu, J. Li, H.Z. Chen, F. Heymans, L. Lelièvre, J.E. Ombetta, F. Massicot, Toxicol. In Vitro 20 (2006) 824–831.