Synthesis, spectral characterization, DNA binding ability and anti-cancer screening of new acridine-based derivatives

Article abstract of DOI:10.1007/s00044-017-1931-9

In this study, a series of newly synthesized acridine derivatives, compounds 4, 6a, and 6b, are described and their biological activity on HL-60 cell lines is assessed using a number of different techniques. Binding studies were also performed between the

Full text of DOI:10.1007/s00044-017-1931-9

MEDICINAL
CHEMISTRY
RESEARCH
Med Chem Res
DOI 10.1007/s00044-017-1931-9
ORIGINAL RESEARCH
Synthesis, spectral characterization, DNA binding ability and
anti-cancer screening of new acridine-based derivatives
Othman M. Salem1 Mária Vilková2 Jana Janočková1,3 Rastislav Jendželovský4
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Peter Fedoročko4 Ján Imrich2 Mária Kožurková1,3
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Received: 4 March 2016 / Accepted: 22 May 2017
© Springer Science+Business Media New York 2017
Abstract In this study, a series of newly synthesized acri-
dine derivatives, compounds 4, 6a, and 6b, are described
and their biological activity on HL-60 cell lines is assessed
using a number of different techniques. Binding studies
were also performed between the derivatives and DNA in
order to characterize the mechanism of the agents’ effect in
more detail. The results of ultraviolet–visible absorption
spectroscopy prove that the binding of derivatives 4, 6a,
and 6b had occurred with a binding constant value of
K = 3.5 × 104–4.0 × 104 M−1. These findings are indicative
of a strong interaction between the derivatives and DNA,
and this hypothesis is supported by the results of the
fluorescence emission, linear dichroism, and viscometric
assays.
Introduction
As the costs of testing and licensing new drugs continue to
rise, there is a growing trend among researchers of rein-
vestigating and improving drugs that are already in the
market. One example of a widely available drug group that
could be reassessed in this manner would be acridine
derivatives. These compounds form one of the best-known
classes of multi-target anti-cancer agents, and the drugs
operate by interfering with DNA synthesis and inhibiting
topoisomerases. A fuller understanding of acridine deriva-
tives would be of considerable interest in our work in the
development DNA topoisomerase inhibitors as chemother-
apeutic agents (Vantova et al. 2009; Janočková et al. 2015a,
b; Salem et al. 2015, 2016). The dynamic nature of the
DNA molecule permits essential biological functions such
as replication, transcription, recombination, repair and DNA
segregation, and topoisomerase enzymes regulate these
topological changes by introducing or removing super-
coiling, knots, or catenations in DNA molecules (Cham-
poux 2001; Vos et al. 2011; Yao et al. 2015). DNA is
generally the primary intracellular target of anti-cancer
drugs, so the interaction between small molecules and DNA
can cause DNA damage in cancer cells and block the
division of cancer cells, thereby resulting in cell death. As
cancer cells exhibit high levels of topoisomerase activity
and demonstrate remarkable sensitivity to DNA-targeted
drugs, a large number of anti-cancer drugs have been
designed and synthesized that specifically target the inter-
action between DNA and topoisomerases (Lafayette et al.
2013; Lang et al. 2013; Li et al. 2014). Despite the side
effects associated with DNA-targeted compounds, topoi-
somerase inhibitors are still recognized as being the main
drug of choice in prolonging the lives of patients, but these
agents are unfortunately accompanied by a number of side
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Keywords Acridine derivatives Topoisomerases I and II
HL-60 cells DNA binding
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* Mária Kožurková
maria.kozurkova@upjs.sk
1
Department of Biochemistry, P. J. Šafárik University, Košice,
Slovakia
2
Department of Organic Chemistry, Institute of Chemistry, P. J.
Šafárik University, Košice, Slovakia
3
Biomedical Research Center, University Hospital Hradec Kralove,
Hradec Kralove, Czech Republic
4
Department of Cellular Biology, Faculty of Science, Institute of
Biology and Ecology, P. J. Šafárik University, Košice, Slovakia

Med Chem Res
effects. Therefore, research into novel DNA-binding agents
that act as topoisomerase inhibitors remains a high priority
in the fight against cancer (Rajendran and Nair 2006; Di
Giorgio et al. 2008; Goodell et al. 2008; Moukharskaya and
Verschraegen 2012; Barros et al. 2012, 2013).
analysis was performed on a Perkin Elmer CHN 2400 ele-
mental analyzer.
All purchased chemical were of reagent grade and used
without further purification. Ethidium bromide (EB),
dimethyl sulfoxide (DMSO), ctDNA, propidium iodide
(PI), EB and tetramethylrhodamine ethyl ester (TMRE),
plasmid pUC 19 (2761 bp, DH 5α), agarose (type II No-A-
6877) were obtained from Sigma-Aldrich Chemie (Ger-
many); EDTA, RNase A, and proteinase K were purchased
from Serva (Germany); fluorescein diacetate (FDA) from
BD Pharmingen (USA); RPMI-1640 medium, penicillin,
streptomycin, amphotericin, fetal bovine serum from Gibco
(Invitrogen, USA); and all other chemicals were purchased
from Lachema (the Czech Republic). HL-60 cell lines were
purchased from American Type Culture Collection (ATCC,
USA).
Our work focuses on the development of novel acridine
derivatives that possess bioactive heterocyclic substituents,
which are able to interact or interfere with cell processes or
metabolites. We also aim to produce new approaches and
techniques for the synthesis and preparation of such agents
that would be attractive and viable from a chemical point of
view. As a continuation of our research interest (Ungvarská
Maľučká et al. 2016) in the chemical transformation of
acridine moiety, we decided to investigate other
carbon–carbon double-bond forming reactions. In this
study, we synthesize and describe three new acridine deri-
vatives that possess a biologically active CH=CH moiety, a
feature that would be expected to demonstrate some degree
of anti-cancer activity. The biochemical and biological
activities of acridine derivatives 4, 6a, and 6b were tested in
order to examine their capacity to bind to DNA and to
interfere with calf thymus topo I. The binding constants of
these dyes with calf thymus DNA (ctDNA) were deter-
mined using ultraviolet–visible (UV–Vis) measurements
and these results will be presented as an approach to the
investigation of in vivo interactions. Linear dichroism (LD)
spectroscopy, viscosity measurement, and DNA melting
studies were also performed in order to obtain a deeper
insight into the DNA-binding affinity of these novel deri-
vatives. The biological activity of the novel compounds
were assessed using a number of techniques, which include
examinations of cell cycle distribution and changes in the
mitochondrial membrane potential (MMP) in HL-60 cell
lines.
Synthesis of (2E)-3-(acridin-9-yl)prop-2-enoic acid (2) A
solution of sodium hydroxide (2 equiv) was added to a
solution of methyl (2E)-3-(acridin-9-yl)prop-2-enoate (1,
500 mg, 1.90 mmol) in methanol (10 mL). After stirring for
8 h, the solvent was evaporated in vacuo. The reaction was
quenched by the addition of water (25 mL). The aqueous
portion was acidified to pH 2 with 6 M hydrochloric acid.
The carboxylic acid was precipitated on standing, filtered
off, and dried (Vilková et al. 2016).
Yield 420 mg, 89%. Mp 225–226 °C. Bright yellow solid.
[Found: C 77.21; H 4.22; N 5.49; C16H11NO2 (249.269)
1
requires C 77.10; H 4.45; N 5.62%]. H NMR (600 MHz,
DMSO-d6): 8.14 (1H, d, J 16.8, H-3), 8.19 (2H, dd, J 7.2,
H-1′,8′), 8.18 (2H, dd, J 7.2, H-4′,5′), 7.88 (2H, ddd, J 6.6,
1.2, H-3′,6′), 7.66 (2H, ddd, J 8.4, H-2′,7′), 6.43 (1H, d, J
16.2, H-2); 13C NMR (150 MHz, DMSO-d6): 166.8 (C-1),
148.5 (C-4′a,10′a), 140.7 (C-9′), 138.5 (C-3), 131.0 (C-2),
130.9 (C-3′,6′), 130.0 (C-4′,5′), 127.1 (C-2′,7′), 125.9 (C-
1′,8′), 123.6 (C-8′a,9′a) ppm.
Materials and methods
Chemistry
Synthesis of (2E)-3-(acridin-9-yl)prop-2-enoyl chloride
(3) The acid chloride was prepared through the reaction
of carboxylic acid 2 (500 mg, 2.00 mmol) and thionyl
chloride (480 mg, 4.03 mmol) in freshly distilled dichlor-
omethane (10 mL) by heating under reflux for 8 h. The
excess of thionyl chloride was removed under reduced
pressure and the resulting product was used in the next step
without further purification (Ilis et al. 2011).
General information and materials
Nuclear magnetic resonance (NMR) spectra were recorded
at room temperature on a Varian VNMRS spectrometer
operating at 599.87 MHz for H, 150.84 MHz for 13C, and
1
60.79 MHz for 15N. Spectra were recorded in CDCl3 or
DMSO-d6 and the chemical shifts were referenced to the
Yield 480 mg, 88%. 1H NMR (600 MHz, DMSO-d6):
8.59 (2H, d, J 9.0, H-4′,5′), 8.56 (1H, d, J 16.2, H-3), 8.48
(2H, d, J 7.2, H-1′,8′), 8.30 (2H, t, J 7.2, H-3′,6′), 7.94 (2H,
t, J 7.2, H-2′,7′), 6.58 (1H, d, J 16.2, H-2); 13C NMR (150
MHz, DMSO-d6): 165.6 (C-1), 151.9 (C-9′), 140.1 (C-4′
a,10′a), 136.2 (C-3), 136.1 (C-3′,6′), 133.1 (C-2), 128.0 (C-
2′,7′), 127.0 (C-1′,8′), 123.6 (C-8′a,9′a), 121.1 (C-4′,5′)
ppm.
residual solvent signal (CDCl3: H NMR 7.26 ppm, 13C
1
NMR 77.0 ppm; DMSO-d6: H NMR 2.50 ppm, 13C NMR
1
39.5 ppm). 2D gCOSY, gHSQC, gHMBC (optimized for a
long-range coupling of 8 Hz) methods were employed
(Vilková et al. 2016). Melting points were determined on a
Boetius hot-stage apparatus and are uncorrected. Elemental

Med Chem Res
1
Synthesis of propyl (2E)-3-(acridin-9-yl)prop-2-enoate
(4) (E)-3-(acridin-9-yl)propenoyl chloride (3, 1 equiv)
was added to a stirred cooled dichloromethane solution (0 °
C, 1 mL) of triethylamine (2 equiv) and ethanol (5 equiv).
After stirring for 24 h at room temperature, the reaction
mixture was then concentrated in vacuo, and the residue
was purified by column chromatography on silica gel (1:1
hexane-ethyl acetate) to produce ester 4.
77.29; H 4.32; N 8.58%]; H (600 MHz, DMSO-d6): 8.72
(1H, t, J 2.0, H-2″), 8.45 (1H, d, J 17.0, H-2), 8.43 (2H, d, J
8.5, H-1′,8′), 8.37 (1H, d, J 7.8, H-6″), 8.25 (1H, dd, J 8.2,
2.4, H-4″), 8.19 (2H, d, J 8.7, H-4′,5′), 7.88 (2H, ddd, J 8.8,
6.5, 1.3, H-3′,6′), 7.78 (1H, t, J 8.0, H-5″), 7.65 (2H, ddd, J
8.8, 6.5, 1.3, H-2′,7′), 7.31 (1H, d, J 16.7, H-1); 13C (150
MHz, DMSO-d6): 148.4 (C-3″), 148.2 (C-4′a,10′a), 142.2
(C-9′), 138.2 (C-1″), 137.1 (C-1), 133.3 (C-6″), 130.3 (C-
6′), 130.2 (C-5″), 129.5 (C-4′,5′), 126.1 (C-1′,8′; C-2′,7′),
125.4 (C-2), 123.7 (C-8′a,9′a), 123.0 (C-4″), 121.8 (C-2″);
15N (61 MHz, DMSO-d6): −75.1 (N-10′), −9.3 (NO2) ppm.
The novel acridine derivatives 4, 6a, and 6b were
dissolved in absolute DMSO at a concentration of 5 ×
10−3 M. The solutions were stored in darkness at 4 °C. A
stock solution of ctDNA was prepared in a TE buffer (pH =
7.4). The concentrations of the DNA samples per mole
phosphate were determined spectrophotometrically at 260
Yield 70 mg, 65%. Bright yellow solid. Mp 179−180 °C.
[Found: C 78.21; H 5.66; N 4.65; C19H17NO2 (291.349)
1
requires C 78.33; H 5.88; N 4.81%]. H NMR (600 MHz,
CDCl3): 8.54 (1H, d, J 16.2, H-3), 8.29 (2H, d, J 9.0, H-
4′,5′), 8.19 (2H, d, J 8.4, H-1′,8′), 7.81 (2H, t, J 9.0, 7.8, H-
3′,6′), 7.58 (2H, t, J 8.4, 7.8, H-2′,7′), 6.48 (1H, d, J 16.2,
H-2), 4.30 (2H, t, J 14.1, H-1″), 1.82 (2H, k, J 14.1, 7.2, H-
2″), 1.05 (3H, t, J 7.2, H-3″); 13C NMR (150 MHz, CDCl3):
165.7 (C-1), 148.0 (C-4′a,10′a), 140.4 (C-9′), 139.0 (C-3),
130.5 (C-3′,6′), 129.6 (C-2, C-4′,5′), 126.5 (C-2′,7′), 125.2
(C-1′,8′), 123.8 (C-8′a,9′a), 66.8 (C-1″), 22.1 (C-2″), 10.5
(C-3″) ppm.
nm using a molar extinction coefficient of 6600 cm−1 M−1
.
UV–Vis spectroscopic studies
General procedure for the synthesis of 9-[(E)-2-
UV–Vis spectra were obtained using a Varian Cary 100
UV–Vis spectrophotometer in pH 7.4 buffer at
(nitrophenyl)ethenyl]acridine 6a, b
a
room temperature using 1 × 1 cm quartz cuvettes.
The UV–Vis spectra of compounds 4, 6a, and 6b and
compounds–ctDNA complexes were recorded at a wave-
length range of 200−600 nm, in a 10 mM Tris-HCl buffer,
pH 7.4. Experiments were carried out in with fixed con-
centrations of derivatives 4, 6a, and 6b (6.25 × 10−5 M)
and varying concentrations of DNA (0–31 × 10−4 M).
A mixture of 9-methylacridine (5) (1.0 g, 5.17 mmol),
appropriate aldehyde (0.782 g, 5.17 mmol), and zinc
chloride (0.776 g, 5.70 mmol) was heated at 130 °C for 3 h.
The reaction mixture was cooled to room temperature and
was partitioned between aqueous sodium hydroxide and
chloroform. The organic phase was washed with water and
brine, dried over magnesium sulfate, filtered, and con-
centrated in vacuo. The residue was purified by silica gel
column chromatography (4:1 hexane-acetone) to produce
compound 6 (Tsukamoto et al. 2009).
Thermal denaturation
Thermal studies were carried out using a Varian Cary
Eclipse spectrophotometer, equipped with a temperature
controlling programmer ( 0.1 °C). The absorbance at 260
nm was continuously monitored (10 mM Tris-HCl buffer,
pH 7.4) for solutions of ctDNA (3.2 × 10−3 M) in the
absence and presence of the complexes (7.5 × 10−6 M). The
9-[(E)-2-(4-nitrophenyl)ethenyl]acridine (6a) Yield 1.12
mg, 66%. Bright yellow solid. Mp 286−287 °C. [Found: C
77.17; H 4.25; N 8.34; C21H14N2O2 (326.354) requires C
1
77.29; H 4.32; N 8.58%]; H (600 MHz, DMSO-d6): 8.49
(1H, d, J 16.2, H-2), 8.41 (2H, d, J 9.0, H-1′,8′), 8.34 (2H,
d, J 8.7, H-3″,5″), 8.20 (2H, d, J 8.5, H-4′,5′), 8.16 (2H, d, J
8.7, H-2″,6″), 7.89 (2H, ddd, J 8.8, 6.5, 1.4, H-3′,6′), 7.66
(2H, ddd, J 8.9, 6.5, 1.3, H-2′,7′), 7.32 (1H, d, J 16.6, H-1);
13C (150 MHz, DMSO-d6): 148.2 (C-4′a,10′a), 147.0 (C-
4″), 142.9 (C-1″), 141.9 (C-9′), 137.3 (C-1), 130.3 (C-3′,6′),
129.6 (C-4′,5′), 128.3 (C-2″,6″), 127.2 (C-2), 126.2 (C-
2′,7′), 125.9 (C-1′,8′), 124.0 (C-3″,5″), 123.6 (C-8′a,9′a);
15N (61 MHz, DMSO-d6): −74.4 (N-10′), −9.8 (NO2) ppm.
temperature of the solution was increased by 0.4 °C min−1
.
Equilibrium binding titration
The binding affinities were calculated from absorbance
spectra according to the McGhee and von Hippel method
using data from a Scatchard plot (McGhee and von Hippel
1974; Jenkins 1997). The binding data were fitted using
GNU Octave 2.1.73 software (Busa 2006).
9-[(E)-2-(3-nitrophenyl)ethenyl]acridine (6b) Yield 1.38
g, 82%. Bright yellow solid. Mp 208−209 °C. [Found: C
77.15; H 4.14; N 8.29; C21H14N2O2 (326.354) requires C

Med Chem Res
Fluorescence measurements
the acridine derivatives (5, 30, 60, and 80 μM, respectively)
and these observations were used to estimate the effect.
Fluorescence spectra were recorded at room temperature at
a range of 520–800 nm with an excitation wavelength at
500 nm. The widths of both the excitation and emission slits
were set at 10 nm. The intercalating effect of derivatives 4,
6a, and 6b (1.2 × 10−4 M) with the ctDNA–EB complex
was studied by adding specific quantities of the compound
into the quartz cuvette containing a fixed concentration of
the ctDNA–EB complex solution. The concentration of EB
was 6.0 × 10−5 M, while that of the ctDNA was 1.0 × 10−6
M. The influence of the addition of each compound to the
DNA–EB complex solution was observed by recording
variations in the fluorescence emission spectra.
Cell culture conditions
HL-60 cell lines were purchased from ATCC. Cells were
grown in a RPMI-1640 medium supplemented with fetal
bovine serum and antibiotics (100 U mL−1 penicillin; 100
µg mL−1 streptomycin; 25 µg mL−1 amphotericin) at 37 °C,
95% humidity and in an atmosphere of 5% CO2 (Janočková
et al. 2015b). For the experiments, cells were seeded in 96-
well plates (MTT assay) or in 6-well plates (MMP dis-
sipation, viability, cell cycle) (Trasadingen, Switzerland) to
which the acridine derivatives were added. The results were
analyzed 48 and 72 h after the addition of the compounds.
Acridine was used as a positive control.
LD spectroscopy
Flow LD spectra were collected by using a flow Couette cell
in a Jasco J-720 spectropolarimeter adapted for LD mea-
surements. It is possible to orient long molecules such as
DNA (minimum length of ~250 bp) 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 pathlength of 1 mm. LD spectra of
ctDNA at a concentration of 2.0 × 10−4 M modified by
acridine derivatives 4, 6a, and 6b (0.79–3.96 × 10−5 M)
were recorded at 25 °C in a 10 mM Tris-HCl buffer (pH 7.4)
in the range of 220–600 nm.
MTT assay
The metabolic activity of cells was evaluated using a MTT
(3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bro-
mide) reduction assay. At scheduled time points (48 and
72 h) MTT (5 mg mL−1) was added and the cells (1 × 104)
were incubated for another 4 h at 37 °C. The reaction was
stopped and formazan crystals were dissolved by the addi-
tion of 100 µL sodium dodecyl sulfate (10%). The absor-
bance was measured at 584 nm using a BMG FLUOstar
Optima (BMG Labtechnologies GmbH, Germany). Results
were evaluated as percentages of absorbance of the
untreated control.
Viscometric measurements
Viscosity experiments were conducted on a AMVn Auto-
mated Micro Viscometer (Anton Paar GmbH, Austria)
using a 1.6-mm capillary tube at constant temperature of 25
0.1 °C. Titrations were performed by adding various
concentrations of derivatives 4, 6a, and 6b (2.5–10 × 10−5
M) to a ctDNA solution (1.6 × 10−3 M) which had been
placed in the viscometer. Data were presented as (η/η0)1/3
vs. the ratio of the concentration of the compound to
ctDNA, where η is the viscosity of ctDNA in the presence
of the compound and η0 is the viscosity of ctDNA in the
absence of the compound. Viscosity values were calculated
from the observed flow time of the DNA containing solu-
tions (t) and corrected for buffer solution (t0), η = (t–t0)/t0.
Detection of MMP dissipation, metabolic activity and
viability
HL-60 cells (1 × 106) were treated with varying con-
centrations of the studied acridine derivatives (10–50 ×
10−6 M) and with acridine (2–10 × 10−6 M) as a positive
control. Detection of MMP dissipation, metabolic activity,
and viability was performed as described by Janočková
et al. (2015a). After 48 and 72 h, incubation cells were
harvested using centrifugation (5 min, 24 °C, 800 rpm),
washed once with Hank’s balanced salt solution and stained
with TMRE (0.1 µmol dm−3) in the case of MMP dissipa-
DNA topoisomerase I relaxation assay
tion detection or double staining with FDA (100 ng mL−1
)
The effects of the acridine derivatives on the relaxation of
plasmid DNA with topoisomerase I were performed using
the methods described previously by Janočková et al.
(2015a, b). A plasmid pBR322 (1.4 μg) was incubated for
45 min at 37 °C with two units of calf thymus topoisome-
rase I (Takara, Japan) in both the absence and presence of
and PI (25 µg mL−1) in the case of metabolic activity and
viability detection. These experiments were examined using
a BD FACS Calibur flow cytometer (Becton Dickinson,
USA), and the results were analyzed using FlowJo software
(TreeStar Inc., USA).

Med Chem Res
Cell cycle analysis
HL-60 cells (1 × 106) were treated with varying con-
centrations of the studied acridine derivatives (10–50 ×
10−6 M) and with acridine (2–10 × 10−6 M) as a positive
control for 48 and 72 h using the method described by
Salem et al. (2016). HL-60 cells were analyzed using a BD
FACSCalibur flow cytometer. ModFit 3.0 software (Verity
Software House, USA) was used to generate DNA content
frequency histograms and to quantify the number of cells in
the individual cell cycle phases.
Statistical analysis
Each experiment was performed at least three times and the
results were expressed as mean values SD. The t-test was
used to determine the significance levels that are indicated
in the legend to each particular figure as p < 0.05 (*),
p < 0.01 (**), and p < 0.001 (***).
Scheme 1 Synthesis of propyl(2E)-3-(acridin-9-yl)prop-2-enoates 4
R
1''
1
H
CH3
H
1'
2
8'
5'
R-CHO
ZnCl2
8'a
9'a
4'a
Results and discussion
Chemistry
130 °C
3 h
10'a
N
N
4'
5
6a,b
a: R = 4-nitrophenyl
b: R = 3-nitrophenyl
The esterification of carboxylic acids and transesterification
of esters have a wide range of academic and industrial
applications (Ramalinga et al. 2002; Khan et al. 2003).
While a variety of procedures exist for the transesterification
of esters, these protocols are often hindered by the need for
excess reagents and/or expensive catalysts, and the pro-
duction process can also produce undesirable side products.
Therefore, we decided to synthesize a new compound,
derivative 4, using conditions that were simple, fast, and
that did not require the use of catalysts. The synthetic
strategies adopted in order to synthesize compound 4 under
the above-mentioned conditions are depicted in Scheme 1.
Ester 4 was synthesized by adapting methodologies that
have been described in previous literature with some very
minor modifications (Scheme 1) (Khurana et al. 2004; Ilis
et al. 2011). The key intermediate, compound 3, was pre-
pared in an excellent yield in two consecutive steps. Methyl
(2E)-3-(acridin-9-yl)prop-2-enoate (1) (Vilková et al. 2016)
was first transformed to (2E)-3-(acridin-9-yl)prop-2-enoic
acid (2) using basic hydrolysis with NaOH in methanol at
room temperature to achieve a complete conversion over the
course of 24 h.
Scheme 2 Synthesis of 9-[(E)-2-(nitrophenyl)ethenyl]acridine 6a and
6b
data determined that the coupling constants between alkenic
protons for derivative 4 had the value of 16.2 Hz, a value
that confirms a trans (E) configuration of alkenes. These
spectral data are consistent with those which have been
reported previously (Vilková et al. 2016).
In a previously published study (Vilková et al. 2016), we
described the synthesis of 9-(2-styryl)acridine from acridin-
9-carbaldehyde using the Wittig reaction. However, on this
occasion, we decided to synthesize 9-(2-styryl)acridine
using the method described by Tsukamoto et al. (2009),
which was simpler, faster, and which did not require the use
of expensive catalysts. Following Tsukamoto’s procedure,
we carried out a condensation of 9-methylacridine (5) with
3- and 4-nitrobenzaldehyde under zinc chloride catalysis
without a solvent to obtain 9-[(E)-2-(nitrophenyl)ethenyl]
acridines (compounds 6a, 6b) (Scheme 2). Compounds 6a
and 6b had an E configuration of the exocyclic double
bond, as was determined from the presence of doublet
Chloride 3 was prepared from carboxylic acid 2 and
thionyl chloride, and was subsequently transformed to ester
4 with appropriate alcohol and trimethylamine at a good
yield as a bright yellow solid product. The new compound
was characterized using elemental analysis and 1D and 2D
1
signals in their H NMR spectra due to olefinic protons at
7.31 (H-1) and 8.45 (H-2) ppm for derivative 6a and 7.32
(H-1) and 8.49 (H-2) ppm for derivative 6b with a coupling
constant of approximately 16 Hz.
1
NMR spectroscopy. The results of the H NMR spectral

Med Chem Res
Absorption titration
at the absorption band into McGhee and von Hippel plots.
The calculated values of the binding constants K ranged
from 3.6 × 104 to 4.0 × 104 M−1, and these results prove
the strong affinity of the acridine ligands toward DNA and
the calculated values also correspond to the range of bind-
ing constants which are characteristic for intercalators (104
to 106).
DNA melting is another important technique that is used
to study the interaction of small molecules with nucleic
acids. The thermal denaturation behavior of DNA in the
presence of different complexes can give an insight into
their conformational changes at increasing temperatures and
offer information about the mode and strength of interaction
of the complexes with DNA (Neyhart et al. 1993). Inter-
calative binding between DNA and small molecules can
stabilize the DNA double-helical structure and this process
increases the Tm by about 5–8 °C. However non-
intercalative binding such as groove binding or electro-
static binding causes little or no perceptible increase in Tm.
Denaturation of DNA is usually measured photometrically
at the temperature-dependent extinction point at 260 nm.
Table 1 shows that Tm of ctDNA was 66.02 °C and that ΔTm
The interaction of the newly synthesized 4, 6a, and 6b
derivatives (Schemes 1 and 2) with ctDNA was monitored
using UV–Vis absorption spectroscopy. This procedure
uses changes in the spectral observations to determine the
existence of interaction between complexes and DNA and
can also to define the mode of interaction that has occurred.
In general, hypochromism may be detected in the case of
intercalative binding of π–π* stacking interactions (Li et al.
1997), while red shifts (bathochromism) are observed when
the DNA duplex has been stabilized (Lang et al. 2013). The
binding of derivatives 4, 6a, and 6b complexes to DNA
helices was determined by following the changes in
absorbance and shifts in wavelength following each addi-
tion of DNA solution to the complex. The acridine deri-
vative complexes in the DMSO-buffer mixture exhibited an
intense transition in the region of 300–450 nm, a result
which is suggestive of π–π* intraligand transitions. A slight
decrease in absorption and a small red shift were observed
following the titration of ctDNA with complexes 4, 6a, and
6b. The absorption spectra for compounds 4, 6a, and 6b
clearly show that the addition of DNA to the complexes
produced hypochromism and a small red shift. These
spectral characteristics strongly suggest that the interaction
of the complexes with DNA involves stacking between the
aromatic chromophore and the base pairs of DNA. The
changes observed in the absorption spectra of ctDNA in the
presence of complex 4 are given in Fig. 1. The results
obtained from the UV–Vis titration experiments (Table 1)
suggest that binding had occurred between the compounds
and ctDNA, although it was not possible to determine the
exact binding mode using UV–Vis spectroscopy alone. In
most cases, the existence of hypochromism would be con-
sidered as convincing evidence of interaction between the
base pairs of ctDNA, and this possibility cannot be ruled
out. It is important to note that the planarity of ligands and
their geometry play a significant role in the ability of the
compounds to interact with DNA.
0,4
Sample 4 : 6.25 × 10-5 M
ctDNA : 0-31× 10-4
M
0,3
0,2
0,1
0
300
350
400
450
500
Wavelenght [nm]
Fig. 1 UV–Vis absorption spectra of studied acridine derivatives
(upper line, derivative 4, 6.25 × 10−5 M) in the presence of increasing
concentrations of ctDNA (color lines, 0–31 × 10−4 M, increment 2 ×
10−4 M) in a 0.01 M Tris-HCl buffer (pH 7.4; 24 °C). Arrow indicates
the absorption change upon increasing amounts of ctDNA
The binding constants, K, of the ligand–DNA complexes
were calculated by integrating the variances in absorbance
Table 1 UV–Vis absorption
data of acridine derivatives 4,
6a, and 6b
a
a
Compounds λmax
λmax
bound
(nm)
Δλ
(nm) (%)
Hypochromicity K × 104
Tm
ΔTm
ΔG
free
(nm)
(M−1
)
(°C) (°C)
(kJ mol−1
)
4
360
373
386
360
374
387
0
1
1
16.21
3.6
3.9
4.0
67.18 1.16
67.18 1.16
67.22 1.20
−19.59
−19.39
−19.32
6a
6b
a
23.88
21.35
Tm measurements were performed in a Tris-HCl buffer (0.01 M), pH 7.4 using derivatives 4, 6a, and 6b
(7.5 × 10−6 M) and ctDNA (3.2 × 10−3 M) at a heating rate of 0.4 °C min−1. Tm of ctDNA for measurement
was 66.02 °C

Med Chem Res
700
600
500
400
300
200
100
0
increased from 0.84 to 1.16 °C in the presence of com-
pounds 4, 6a, and 6b. The high ΔTm value for the com-
plexes is suggestive of an intercalative mode of binding of
complexes to DNA. In our case, only a slight shift increase
in Tm was observed in the presence of the studied com-
plexes in comparison with that recorded for free DNA. The
small increase in DNA melting temperature in the presence
of the complexes indicates that groove binding is the pre-
dominant mode of binding between the complexes and
DNA (Kamatchi et al. 2013; Chitrapriya et al. 2011).
4
550
600
650
700
750
800
Fluorescence studies
Wavelenght [nm]
700
600
500
400
300
200
100
0
In order to investigate the binding ability of studied deri-
vatives and DNA in more detail, fluorescence absorption
spectroscopy was employed. In the case of EB displacement
assay, a solution containing a fixed concentration of EB and
ctDNA was titrated with increasing concentrations of deri-
vatives 4, 6a, and 6b. EB–ctDNA complexes were excited
at 500 nm and emission spectra were recorded from 520 to
800 nm. EB is a well-known intercalator that is often used
as a spectral probe to estimate the binding mode of small
molecules to double-helical DNA. The representative
emission spectra of EB bound to DNA in both the absence
and the presence of compounds 4, 6a, and 6b are given in
Fig. 2. If the titration of ctDNA-EB with compounds 4, 6a,
and 6b results in a decrease in fluorescence intensity, this
would indicate that the studied derivatives compete with EB
for interaction sites in DNA and dissociate EB from the
system. As a consequence, derivatives 6a and 6b demon-
strated a clear reduction in emission intensity, thereby
suggesting that competitive binding interaction between
derivatives 6a and 6b and EB with ctDNA had occurred,
and also that derivatives 6a and 6b may interact with
ctDNA mainly through an intercalative binding mode.
Unlike derivatives 6a and 6b, derivative 4 showed a very
slight reduction in emission spectra, indicating that this
derivative had not displaced EB from the DNA–EB com-
plex. These findings would suggest that derivative 4 is
unlikely to interact with ctDNA through intercalation, and
that any interaction is likely to be through an alternative
mode of binding. However, not only the DNA intercalators
but also groove DNA binders could cause the reduction in
the emission intensities of DNA bound (Cain et al. 1978)
but moderately for the latter case. For example, the non-
intercalating DNA groove-binding agents of berenil, bisa-
midines, spermine, and spermidine were reported to be
capable of displacing EB from DNA (Patel 1979).
6a
550
600
650
700
750
800
Wavelenght [nm]
700
600
500
400
300
200
100
0
6b
550
600
650
700
750
800
Wavelenght [nm]
Fig. 2 Fluorescent displacement of EB (6.0 × 10−5 M, black line)
bound to DNA (1.0 × 10−6 M) with increasing concentrations of
derivatives 4, 6a, and 6b (0–1.2 × 10−4 M), λex = 500 nm, in a 0.01 M
Tris-HCl (pH 7.4, 24 °C), corresponding to the curves from top to
bottom, respectively
embedded into the DNA helical axis. Small molecules that
are unbound or that are bound randomly to ctDNA are not
oriented and show no signal. However, small molecules that
are bound in a specific orientation with respect to ctDNA
become oriented by the flow and their orientation shows a
LD signal (Terenzi et al. 2013). The LD of these samples is
characterized as the difference between the absorption of
light polarized parallel (A║) and perpendicular (A┴) to the
axis. LD measurements were also applied in order to
investigate the DNA-binding mode of derivatives 4, 6a, and
Linear dichroism
LD measurements are often used to determine the orienta-
tion of ligands and their chromophores that are potentially

Med Chem Res
0.01
0.00
can cause a decrease or increase of this band. Decreases in
intensity could be connected to the bending or coiling
potency of the ligands, while increases in intensity could be
a result of intercalation, or the stiffening of the DNA. The
LD spectra of ctDNA (2.0 × 10−4 M) in both the absence
and presence of increasing concentrations of compounds 4,
6a, and 6b (0.79–3.96 × 10−5 M) are presented in Fig. 3.
The results show that compounds 4 and 6b produced an
increase in the intensity of the DNA negative band and that
compound 6a caused a decrease in the signal in this region.
Moreover, the positive LD signals observed in the region of
300–600 nm suggest that compounds 6a and 6b bind to
DNA in an oriented non-random fashion (Phelps et al.
2013).
4
-0.01
-0.02
-0.03
-0.04
300
400
500
600
Wavelength [nm]
Viscometry
In order to clarify the mode of binding of derivatives to
ctDNA, viscosity measurements were also carried out using
varying concentrations of derivatives 4, 6a, and 6b in order
to gather more information on the possible binding modes
of the studied compounds and ctDNA. Intercalation binding
typically causes an increase in the viscosity of the
ligand–DNA complex because this form of binding requires
a large space between adjacent base pairs in order to
accommodate the ligand and lengthen the double helix
(Hannon et al. 2001). In contrast, small molecules that bind
exclusively in DNA grooves by partial or non-typical
intercalation usually cause little or no change in the visc-
osity of the DNA solution (Ling et al. 2008; Rehman et al.
2014). The results obtained from the viscosity measure-
ments of the ligand–ctDNA complex at increasing amounts
of derivatives 4, 6a, and 6b are presented in Fig. 4. No
significant increase in the relative viscosity of ctDNA was
observed at increasing concentrations of derivatives 4, 6a,
and 6b, and this suggests that derivatives 4, 6a, and 6b
Fig. 3 LD spectra of ctDNA (2.0 × 10−4 M, black line) in the pre-
sence of increasing amounts of derivatives 4, 6a, and 6b (0.79–3.96 ×
10−5 M, color line) in a 0.01 M Tris-HCl solution (pH 7.4, 24 °C)
6b in more detail. The DNA base pairs are perpendicular to
the helical axis, and ctDNA in its B-form demonstrates a
representative negative band in the range of 220–300 nm, a
finding which is likely the result of π–π* transitions within
the bases. Ligands that are able to alter the structure of DNA
Fig. 4 Effect of increasing concentration of derivatives 4, 6a, and 6b
(2.5–10 × 10−5 M) on the viscosity of ctDNA. Concentration of
ctDNA was constant (1.6 × 10−3 M) while the concentration of deri-
vatives 4, 6a, and 6b varied (0.01 M Tris-HCl buffer, pH 7.4; 24 °C)

Med Chem Res
prefer to interact with DNA through non-intercalative
binding modes.
in Fig. 5, the supercoiled pBR322 DNA was fully relaxed
by the enzyme in the absence of the studied derivatives
(lane pBR322 + topo I). Derivatives 4, 6a, and 6b
demonstrated topo I inhibitory activity at 60 µM, with the
exception of derivative 6a which partially inhibited the
activity of topo I at a concentration of 30 µM. All of these
results indicate that these derivatives show some evidence
of interaction with DNA and the inhibition of topo I, which
in turn suggests that the compounds could be potential
DNA topoisomerase I inhibition assay
In order to establish whether our newly synthesized deri-
vatives possess the ability to inhibit topo I, a relaxation
assay was performed using derivatives 4, 6a, and 6b and
plasmid pBR322 DNA mediated by topo I. As can be seen
Fig. 5 Effect of the compounds on the
relaxation of plasmid DNA by human
topoisomerase I. pBR322 (Lane
pBR322) was incubated with topo I
(lane pBR322 + topo I) and studied
compounds 4, 6a and 6b (5, 30, 60, and
80 µM) 45 min at 37 °C. The DNA
samples were run on agarose gel
followed by EB staining. Sc supercoiled
plasmid, oc open circular plasmid
Fig. 6 MTT assay: effect of derivatives 4, 6a, and 6b and acridine (ACR) (0–100 × 10−6 M) on the metabolic activity of HL-60 cells at 24, 48,
and 72 h after treatment

Med Chem Res
Fig. 7 The effect of acridine
(ACR, positive control, 2–10 ×
10−6 M) and derivatives 4, 6a,
and 6b (10–50 × 10−6 M) on
changes in MMP 48 and 72 h
after treatment. The results are
presented as a mean SD,
statistical significance: p < 0.05
(*), p < 0.01 (**), p < 0.005
(***) for particular experimental
group compared to untreated
control (C)
Fig. 8 The effect of acridine
(ACR, positive control, 2–10 ×
10−6 M) and derivatives 4, 6a,
and 6b (10–50 × 10−6 M) on
changes in viability 48 and 72 h
after treatment. The results are
presented as a mean SD,
statistical significance: p < 0.05
(*), p < 0.01 (**), p < 0.005
(***) for each experimental
group compared to untreated
control (C)
targets candidates for development as new topo I inhibitors
(Basili et al. 2008; Bera et al. 2008).
leukemia cells (CCRF-CEM), or 3,4-dihydro-1H-[1,3]oxa-
zino[4,5-c] acridines (Ouberai et al. 2006) were character-
ized with cytotoxic activity against HT-29 colon
adenocarcinoma cell line. In this study, we also investigated
the effectiveness of new acridine derivatives 4, 6a, and 6b
on the HL-60 human promyelocytic leukemia cell line using
various flow cytometer techniques.
The effect of derivatives 4, 6a, and 6b on the metabolic
activity of HL-60 cells was determined by examining the
number of MTT metabolizing cells and their metabolic
potency after treatment with varying concentrations of the
Cytotoxic studies
While considerable research has focused on the develop-
ment of acridine derivatives as potential anti-cancer agents,
the highly cytotoxic nature of the compounds has limited
their clinical application to very specific uses. For example,
9-anilino acridine derivatives (Su et al. 2006) exhibited
potent in vitro cytotoxicity against human lymphoblastic

Med Chem Res
studied derivatives (0–100 × 10−6 M) over 24, 48, or 72 h.
The results show that the studied derivatives had a dose-
dependent and time-dependent inhibitory effect on the
metabolic activity of HL-60 cells; the effect was relatively
weak at lower concentrations and over shorter time periods,
but at concentrations greater than 20 × 10−6 M after 72 h
incubation the effect was markedly stronger (Fig. 6).
Further tests were then performed in order to determine
whether the cytotoxic effects of the studied derivatives can
be related to the changes in the MMP, the mechanism
implicated in cell death and in the interference of the cell
cycle progression.
Cell death can be activated by a mitochondria-dependent
apoptotic process, which includes the opening of mito-
chondrial permeability transition complex pore, mitochon-
drial transmembrane potential dissipation, and other factors.
Table 2 IC50 values for the MMP and viability evaluated 48 and 72 h
after treatment with acridine (ACR) and derivatives 4, 6a, and 6b
TMRE IC50 (μM) 48 h
72 h
PI IC50 (μM) 48 h
72 h
4
31.58 30.58
4
36.15 27.85
6a
nd
nd
6a
nd
nd
6b
ACR
nd
nd
6b
ACR
nd
nd
6.37
5.23
6.40
5.04
TMRE tetramethylrhodamine ethyl ester, PI propidium iodide
Fig. 9 The effect of acridine
(ACR, positive control, 2–10 ×
10−6 M) and derivatives 4, 6a,
and 6b (10–50 × 10−6 M) on
cell cycle distribution 48 and
72 h after treatment. The results
are presented as a mean SD,
statistical significance: p < 0.05
(*), p < 0.01 (**), p < 0.005
(***) for each experimental
group compared to untreated
control (C)

Med Chem Res
MMP is lost during the process of apoptosis through the
disruption of energy metabolism (Salem et al. 2016),
therefore we decided to investigate the rate of MMP dis-
sipation induced by derivatives 4, 6a, and 6b (Fig. 7).
Derivative 4 caused a significant reduction in the MMP of
HL-60 cells after 72-h treatment at a concentration of at 50
μM. Treatment with derivatives 6a and 6b had a weaker
effect on the MMP in comparison to that recorded for the
positive control, acridine. The analysis of cell viability
revealed similar results as that of the MMP analysis. Deri-
vative 4 and the acridine control were more effective than
derivatives 6a and 6b (Fig. 8). IC50 values were also cal-
culated for both techniques (MMP and viability) and for
different time points (Table 2).
derivative 4 (27.85 µM, 72 h) is several times lower than
that of model amsacrine (IC50 = 0.34 µM).
Our results obtained from this work identify various
areas of valuable information relating to ligand–DNA
interactions, which could be useful for rational drug design
and helpful in the development of their potential pharma-
ceutical and biological effect.
Acknowledgements This study was supported by Slovak Research
and Development Agency under contract VVCE-0001-07, VEGA
grants No. 1/0001/13, and 1/0672/11 and MH CZ-DRO (UHHK,
00179906). The authors would like to thank Jana Kašpárkova and
Viktor Brabec for their advice on LD measurement and viscometric
techniques.
As can be seen in Fig. 9, a significant increase in the
number of cells in the G1 phase of cell cycle was observed
following 48-h exposure to 10, 20, and 50 µM of derivatives
4 and 6a and the acridine control. Moreover, derivatives 4
and 6b caused a reduction of cells in the S phase at 50 μM.
A corresponding increase in the cell population in the G2
phase of cell cycle was also observed after 48-h exposure to
50 μM of derivatives 4 and the acridine control. Similar
results were obtained following the incubation of HL-60
cells with 10, 20, and 50 µM of the studied derivatives for
72 h. Derivative 4 had a clear effect on cell cycle distribu-
tion, visible as an increased proportion of cells in the G2
phase with a simultaneous reduction in the number of cells
in the G1 phase.
Compliance with ethical standards
Conflict of interest The authors declare that they have no competing
interests.
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