Synthesis and preliminary biological evaluation for the anticancer activity of organochalcogen (S/se) tethered chrysin-based organometallic RuII(η6-p-cymene) complexes

Article abstract of DOI:10.1080/07391102.2018.1513867

Organochalcogen (S/Se) functionalized chrysin derivatives were synthesized and coordinated with Ru^II(η⁶-p-cymene) to efficiently form ruthenium-based chemotherapeutic drug entities [C₃₁H₃₅O₄SRuCl]; [C

Full text of DOI:10.1080/07391102.2018.1513867

Journal of Biomolecular Structure and Dynamics
ISSN: 0739-1102 (Print) 1538-0254 (Online) Journal homepage: http://www.tandfonline.com/loi/tbsd20
Synthesis and preliminary biological evaluation
for the anticancer activity of organochalcogen
(S/Se) tethered chrysin based organometallic
RuII(η6–p–cymene) complexes
Shipra Yadav & Jai Deo Singh
To cite this article: Shipra Yadav & Jai Deo Singh (2018): Synthesis and preliminary biological
evaluation for the anticancer activity of organochalcogen (S/Se) tethered chrysin based
organometallic RuII(η6–p–cymene) complexes, Journal of Biomolecular Structure and Dynamics,
DOI: 10.1080/07391102.2018.1513867
To link to this article: https://doi.org/10.1080/07391102.2018.1513867
Accepted author version posted online: 20
Aug 2018.
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Synthesis and preliminary biological evaluation for the anticancer activity of
organochalcogen (S/Se) tethered chrysin based organometallic RuII(η6–p–cymene)
complexes
Shipra Yadav and Jai Deo Singh*
Department of Chemistry, Indian Institute of Technology Delhi (IITD), Hauz Khas, New Delhi
110016, India
*Corresponding author
E–mail: jaideo@chemistry.iitd.ernet.in
Abstract
Organochalcogen (S/Se) functionalized chrysin derivatives were synthesized and coordinated
with RuII(η6–p–cymene) to efficiently form ruthenium based chemotherapeutic drug entities
[C31H35O4SRuCl]; [C31H35O4SeRuCl]; [C33H31O4SRuCl] and [C33H31O4SeRuCl]. The complexes
were thoroughly characterized by analytical and various spectroscopic techniques which include
elemental analysis, UV–vis, IR, NMR (1H, 13C and 77Se NMR) and HR–MS. The interaction
studies of these Ru(II) complexes were carried out with CT DNA/HSA by employing UV–vis,
fluorescence and circular dichroic techniques in view to examine their chemotherapeutic
potential. The complexes demonstrated predominant binding towards CTDNA via electrostatic
interaction while, the extent of binding was quantified by calculating intrinsic binding constant
(Kb) and binding constant (K) values which revealed higher binding affinity of selenium based
chrysin complexes as compared to their thio–analogs, following the order [C31H35O4SeRuCl] >
[C33H31O4SeRuCl] > [C31H35O4SRuCl] > [C33H31O4SRuCl]. Moreover, interaction of these
complexes with human serum albumin (HSA) was also investigated which suggested
spontaneous interactions of complexes with the protein by hydrogen bonding and van der Waals
forces. To visualize the preferential binding sites and affinity of complexes with DNA and HSA
molecular docking studies were performed. Additionally, in vitro anticancer activity of the
complexes were evaluated by SRB assay on selected cancer cell lines viz., HeLa (cervical),
MIA–PA–CA–2 (pancreatic), MCF–7 (breast), Hep–G2 (Hepatoma) and SK–OV–3 (ovarian)
which exhibited the superior cytotoxicity of complex [C31H35O4SeRuCl] as compared to other
analogs on selective cancer phenotypes.
1

Keywords: Organochalcogen; RuII(η6–p–cymene)complexes; DNA/HSA interaction; In vitro
Cytotoxicity; Molecular docking.
Abbreviations: Chry, Chrysin; Cym, Cymene; HSA, Human serum albumin; Kb, Binding
constant; Kq Quenching constant; Ksv, Stern-Volmer constant.
1. Introduction
Platinum based drugs, cisplatin and the follow–on (carboplatin and oxaliplatin) are dominated
metallo–chemotherapeutic agents used in a wide number of anticancer treatment regimens
(Kelland, 2007). However, their incidences of indiscriminant “off‒target” side effects including
toxicity and tumor metastasis have limited their effective use as chemotherapeutic agents
(Wheate, Walker, Craig and Oun, 2010; Alessio, 2011; Jaouen and Metzler–Nolte, 2010).
Consequently, the current cancer treatment scenario demands an urgent and alternative need to
develop potential metal complexes that could surmount Pt–based resistances, improve clinical
efficacy with reduced toxicity. In this pursuit, Ru(II)–complexes have emerged as a notable class
of anticancer drugs because of their favorable kinetic aspects, rich redox chemistry, inherently
less toxicity and high selectivity for cancer cells (Allardyce and Dyson, 2001; Clarke, 2003;
Hall, Beer, Buchner, Cardin and Cardin, 2015). Significantly, ruthenium complexes might
interfere with multiple targets including DNA, proteins and enzymes (Nazarov, Hartinger and
Dyson, 2014; Adhireksan et al., 2014; Noffke, Habtemariam, Pizarro and Sadler, 2012,),
resulting in comparable or superior cytotoxicity profile against a wide spectrum of cancer
phenotypes including the cisplatin–resistant strains (Dyson, 2007) in contrast to Pt–drugs that
exhibit cytotoxicity mainly due to covalent interactions with DNA (Reedijk, 1996).
Notably, the success of Ru(III)–complexes NAMI‒A, [trans‒RuCl4(1H‒imidazole) (DMSO–S)],
KP1019, [trans‒RuCl4(1H‒indazole)2] and its Na+ analogue NKP1339, [trans‒RuCl4(1H‒
indazole)2] in clinical and preclinical trials has led interest in the development of ruthenium
based complexes as a new anticancer agents with improved cytotoxicity (Hartinger et al., 2008;
Hartinger et al., 2006; Rademaker–Lakhai, Van Den Bongard, Pluim, Beijnen and Schellens,
2004; Weiss et al., 2014). However, in recent years, half‒sandwich organoruthenium(II)–arene
complexes have exhibited promising in vivo antimetastatic, antiangiogenic and anticancer
properties (Clavel et al., 2015; Weiss et al., 2015; Berndsen et al., 2017). At the forefront,
RAPTA‒C
([RuII(η6–p–cymene)(PTA)Cl2],
PTA
=
1,3,5‒triaza‒7‒
phosphatricyclo[3.3.1.1]decane), RAED‒C ([RuII(η6–p–cymene)(en)Cl]+) and RM175
2

([RuII(η6‒biphenyl)(en)Cl], en = ethylenediamine) are the representative examples among this
class that are under preclinical trials (Nowak–Sliwinska et al., 2011; Guichard et al., 2006)
(Chart I).
Chart I. Structures of some Ru(II)/(III) promising anticancer agents.
In general, organometallic Ru(II)‒arene complexes structurally offer hydrophobic arene group
which influences the cellular uptake and kinetic reactivity of Ru(II) complexes (Wang et al.,
2005), while the coordinated ligand afford intrinsic control over reactivity and selectivity
towards biomolecular targets (Paunescu, McArthur, Soudani, Scopelliti and Dyson, 2016; Riedl
et al., 2017) (Fig. 1). A wide variety of modifications in Ru(II)‒arene complexes have been
employed at both the arene moiety and the ancillary mono‒ or multidentate ligand systems in an
effort to precisely tune its anticancer activity. This can be arbitrarily done by inclusion of
bioactive organic ligand of known biological functions (Kurzwernhart et al., 2012; Kubanik et
al., 2015; Maria et al., 2009).
3

Fig. 1. Graphical scheme showing the general function of different structural elements.
Among them, flavonoids (plant polyphenols) are the most robust and versatile ligands with wide
range of pharmaceutical properties viz., anti–inflammatory, antimicrobial, antioxidant and
anticancer activity (Cushnie, and Lamb, 2005; Middleton, Kandaswami and Theoharides, 2000;
Pawara, Tandela, Kunabevub and Jaldappagari, 2018). In particular, chrysin is a natural flavone
that has shown to induce growth inhibition and apoptosis in multiple cancer cell lines (Yang et
al., 2014; Pichichero, Cicconi, Mattei and Canini, 2011). Chrysin is an efficient chemopreventive
agent (Rehman et al., 2013) that offer 3‒hydroxy‒4‒keto system as a O,O'–chelating motif for
RuII(η6–p–cymene) moiety which not only augment the bioavailability and multitargeted
approach but also synergizes it to act in concord at the target site (Grazul and Budzisz, 2009;
Kurzwernhart et al., 2012). Recently, reports have revealed that functionalization of chrysin
scaffold improves its aqueous solubility, protein interactions and cytotoxic activity (Singh, Kaur
and Silakari, 2014; Kurzwernhart et al., 2013).
Our interest stems in modulating the chrysin unit by incorporation of organochalcogen moiety
with Ru(II) metal centre which could synergize the effects of these artifacts by specifically
enhancing the selectivity towards biomolecules. Sulfur (S) and selenium (Se) containing
derivatives exhibit prevention against cancer mainly due to their functions in radical scavenging
and enzymatic decomposition of oxygen metabolites (Battin and Brumaghim, 2009). Indeed,
selenium is recognized as an essential micronutrient of particular interest owing to its ability to
participate in crucial redox reactions particularly implicated in antioxidant (Rahmanto and
4

Davies, 2012), chemopreventive (Janakiram et al., 2013) or apoptotic activities (Sanmartín,
Plano and Palop, 2008). Additionally, it has been suggested that mechanisms of action by which
seleno– complexes exert their anticancer acitivity includes induction of apoptosis, inhibition of
angiogenesis and modulation of AKT and COX pathway (Sanmartin, Plano, Sharma & Palop,
2012). In fact, incorporation of selenium in bioactive ligand is a successful strategy that
exhibited synergistic effects to chemotherapeutic drugs (Martins et al., 2013; Qi et al., 2012)
leading to inhibition of cancer cell growth in various xenograft of rat models for different cancer
phenotypes (Zeng, Cheng and Johnson, 2013; Nguyen et al., 2011).
Herein, we report synthesis and characterization of new organometallic Ru(II) complexes
[C31H35O4SRuCl], 1; [C31H35O4SeRuCl], 2; [C33H31O4SRuCl], 3; and [C33H31O4SeRuCl], 4 from
chrysin derivatives (L1‒L4) with the general formula (RE–chry) {where, RE = nBuS (L1); nBuSe
(L2); PhS (L3) and PhSe (L4)}. We have further carried out interaction studies of these
complexes by various spectroscopic studies in order to validate their chemotherapeutic
candidature. The complexes were screened for their cytotoxicity by SRB assay against different
cancer cell lines which revealed the superior cytotoxicity of complex 2 as compared to other
analogs on selective cancer phenotypes. Therefore, the key strategy of our current study provide
an important rationale for the design of new Ru(II) metallodrugs towards exploring the important
modulation in their chemotherapeutic candidature by tethering organochalcogen motif at
bioactive chrysin unit.
2. Experimental section
2.1. Materials
All chemicals viz., bis[(η6–p–cymene)dichloridoruthenium(II)], chrysin (TCI, India), 1,2–
dibromoethane, 1–bromobutane, bromophenyl (Spectrochem), K2CO3 (Fisher scientific),
selenium(CDH),
n–butanethiol,
sodium
methoxide,
thiophenol
(Aldrich),
and
tris(hydroxymethyl)aminomethane or tris buffer (Fisher scientific) were of AR grade and used as
received. Disodium salt of calf–thymus DNA (CT DNA) and human serum albumin (HSA) with
a molecular mass of 66478 Da were purchased from Sigma Aldrich and stored at 4 ºC. 7–(2–
Butylthio)ethoxy)–5–hydroxy–2–phenyl–4H–chromen–4–one (L1), 7–(2–butylselanyl)ethoxy)–
5–hydroxy–2–phenyl–4H–chromen–4–one
phenylthio)ethoxy)–4H–chromen–4–one
(L2),
5–hydroxy–2–phenyl–7–(2–
5–hydroxy–2–phenyl–7–(2–
(L3)
and
phenylselanyl)ethoxy)–4H–chromen–4–one (L4) were synthesized according to previously
5

reported procedures with slight modification (Lee,Park, Kim, Jung and Cho, 1995; Fonseca et
al., 2015). All the reactions were carried out in anhydrous conditions using dry solvents under
inert atmosphere.
2.2. Instrumental methods
Elemental analyses (C, H and N) were performed on Perkin Elmer series II 2400 CHNO Rapid
elemental analyzer. IR spectra for all derivatives were recorded from KBr pellets in the range
1
4000–400 cm–1 using Nicolet, Protege 460 FT–IR spectrometer. H and 13C NMR spectra were
recorded on a Bruker Spectrospin DPX–300 spectrometeroperating at 300.13 and 75.46 MHz,
respectively using CDCl3 as a solvent and tetramethylsilane (TMS) as internal standard, while
77Se NMR spectra were carried on Bruker–400 operating at 76.31 MHz in CDCl3 solvent. High
resolution mass spectra (HR–MS) were obtained on Bruker MS–632 mass spectrometer. Samples
were prepared at µM level in CH3CN solvent. UV–vis spectra were recorded at room
temperature on a Perkin–Elmer Lambda 35 spectrometer against a solvent blank reference in the
wavelength range of 250–650 nm. Fluorescence measurements were carried out on a HORIBA–
Jobin Yvon Scientific Fluoromax–4 equipped with quartz cells (1.0 cm), setting the widths of
both excitation and emission slits to 5 nm. CD spectra were measured on Jasco J–815–CD
spectropolarimeter (Jasco, Japan) at room temperature using a 1 cm quartz cuvette.
2.3. DNA Interaction Studies
2.3.1. UV–vis absorption studies
DNA concentration was determined spectrophotometrically by assuming ε260nm = 6600 M–1cm–1
(Marmur, 1961). The absorption titration of complexes in Tris–HCl buffer were performed by
using a fixed concentration of Ru(II) complexes (0.2 x 10–6 M) with increasing concentration of
CT DNA (0.1–1.0 x10–6 M).The intrinsic binding constant, Kb value of the complex to CT DNA
was determined by Wolfe–Shimer equation (1) (Wolfe, Shimer and Meehan, 1987).
[ DNA ]
[ DNA ]
1
(1)



 
 b  
K
b (  
)
f
a
f
f
a
where [DNA] represents the concentration of DNA and a, f, b, the apparent extinction
coefficient (Aobs/[M]), the extinction coefficient for free metal complex (M), and the extinction
coefficient for the complex (M) in the fully bound form, respectively. The slope and intercept of
the linear fit of the plot of [DNA]/|a–f| vs. [DNA], give 1/|a–f| and 1/Kb|a–f|, respectively.
The intrinsic binding constant, Kb can be obtained from the ration of the slope to the intercept.
6

2.3.2. Fluorescence spectral studies
Relative binding of Ru(II) complexes with CT DNA was studied by fluorescence spectral
method using CT DNA solution in Tris–HCl buffer (pH 7.2). In an experiment, complex solution
(0.2 x10–6 M) was added to the increasing concentrations of CT DNA (0–1.2 x 10–6 M). The
fluorescence intensity measured at excitation wavelength 270 nm in the range of 310–500 nm.
Binding constant, K of the metal complexes was determined using following Scatchard equation
(2 & 3) by emission titration (Cui et al., 2009).
CF = CT (I/Io–P) (1–P)
r/CF = K (n–r)
(2)
(3)
where, CF is the free metal complex concentration, CT is the total concentration of the probe
added, I and Io are fluorescence intensities in presence and absence of CT DNA, respectively and
P is the ratio of the observed fluorescence quantum yield of bound probe to that of the free
probe. The value P was obtained as the intercept by extrapolating from a plot of Io/I vs. 1/[DNA],
r denotes ratio of CB (CB = CT–CF) to the DNA concentration i.e., the bound probe concentration
to the CT DNA concentration, K is the binding constant and n is the number of binding sites.
2.4. HSA Interaction Studies
2.4.1. UV–vis absorption studies
The stock solutions of HSA were prepared by dissolving the solid HSA in 50 mM Tis–HCl, 100
mM NaCl buffer at pH 7.2. HSA concentration was determined from absorption spectra, using
the value of ε278nm = 36000 M–1 cm–1 (Samari, Hemmateenejad , Shamsipur, Rashidi and
Samouei, 2012) and absorbance of a 1 mg ml–1 solution at ~280 nm (λmax Trp–214).The
absorption titrationof complexes in Tris–HCl buffer (pH 7.2) were performed by using a fixed
concentration of HSA (2.0 x 10–6 M) with increasing concentration of complexes (0–4.0 x 10–6
M). The value of binding constants can be calculated from the double reciprocal plot of 1/A–Ao
vs. 1/Ccomplex by using the following equation (4) (Stephanos, 1996):
Ao



1
HSA
HSA


(4)
A  Ao

B K C Complex
B
Where Ao and A are the absorbance of HSA at ~280 nm, in the absence and presence of
complex, respectively. ɛHSA and ɛB are the molar extinction coefficient of HSA and the bound
complex, respectively, and l is the light path of the cuvette (1 cm).
7

2.4.2. Fluorescence spectral studies
Fluorescence measurements were carried with the excitation and emission wavelength set at 290
and 300–500 nm, respectively. For fluorescence measurements, fixed concentration of HSA (1.0
x 10–6 M) was titrated with varying concentrations (0–4.0 x 10–6 M) of complexes at 300 and 310
K. The intensity at 350 nm (Tryptophan) was used to calculate the binding constant, K. The
fluorescence quenching of HSA at different temperatures (300 and 310 K) were determined
using the Stern–Volmer equation (Soares, Mateus and de Freitas, 2007):
(5)
F
 1  K q  [Q ]  1  K SV [Q ]
F
where Fο and F are the fluorescence intensities in absence and presence of quencher,
respectively, [Q] is the quencher concentration, and Ksv is the Stern–Volmer quenching constant,
Kq is the biomolecular quenching rate constant and τo is the average lifetime of the fluorophore
in absence of quencher and its value is around 10–8 s for most biomolecules. Ksvvalues were
calculated from Fo/F vs. [Q] plot, whereas from the log (Fo/F–1) vs. log [Q] plot, the number of
binding sites (n) and the binding constants (K) were calculated.
2.4.3. Determination of thermodynamic parameters
The thermodynamic parameters were calculated from the van′t Hoff equation (Cheng, Fang, Bai,
Ouyang and Hu, 2017):
  H
 S
ln K 

(6)
RT
R
where K is the Lineweaver–Burk static quenching constant at corresponding temperature and R
is the gas constant, in which ∆H and ∆S of reaction was determined from the linear relationship
between ln K and the reciprocal absolute temperature. Furthermore, the free energy change (∆G)
was calculated by the equation:
 G   H  T  S   RT ln K
(7)
2.4.4. Three–dimensional (3D) fluorescence spectral studies
Three–dimensional fluorescence spectroscopy is a method for providing conformational and
structural information of proteins. The maximum emission wavelength and the fluorescence
intensity of the residues have a close relation to the polarity of their micro–environment. The
excitation and emission slits were set at 5 nm andrespective blanks were sbstracted.
8

2.4.5. CD spectral studies
The CD spectra of DNA were collected in the presence of 1–4 at molar ratio of 1:1 in the range
of 200–250 nm, with 20 nm/min scan speed and a response time of 2s. Similarly, the CD spectra
of HSA were collected in the presence of complexes at molar ratio of 1:1 in the range of 200–
250 nm. Measurements were taken at wavelengths between 200 and 250 nm.
2.5. Molecular docking studies
The rigid molecular docking studies were performed by using HEX 6.3 software, which is an
interactive molecular graphics program for the interaction, docking calculations, and to identify
possible binding site of the biomolecules (Mustard and Ritchie, 2005). Structures of the
complexes were sketched by CHEMSKETCH (http://www.acdlabs.com) and converted to pdb
format. The crystal structure of the B–DNA (PDB ID: 1BNA) and HSA (PDB ID: 1h9z) were
retrieved from the protein data bank (http://www.rcsb.org./pdb). Visualization of the docked
pose has been performed by using Pymol and Discovery Studio 3.5 softwares.
2.6. In vitro antitumor studies
The cell lines used for in vitro antitumor activity were HeLa (cervical), MIA–PA–CA–2
(pancreatic), MCF–7 (breast), Hep–G2 (Hepatoma), SK–OV–3 (ovarian). The cell lines were
grown in RPMI 1640 medium containing 10% fetal bovine serum and 2 mM L–glutamine. For
present screening experiment, cells were inoculated into 96 well microtiter plates in 90 µL at
5000 cells per well. After cell inoculation, the microtiter plates were incubated at 37 °C, 5%
CO2, 95% air and 100% relative humidity for 24 h prior to addition of experimental drugs.
Experimental drugs were solubilized in appropriate solvent to prepare stock of 10–2M
concentration. At the time of experiment four 10–fold serial dilutions were made using complete
medium. Aliquots of 10 µl of these different drug dilutions were added to the appropriate micro–
titer wells already containing 90 µl of medium, resulting in the required final drug
concentrations. After compound addition, plates were incubated at standard conditions for 48
hours and assay was terminated by the addition of cold TCA. Cells were fixed in situ by the
gentle addition of 50 µl of cold 30 % (w/v) TCA (final concentration, 10 % TCA) and incubated
for 60 minutes at 4°C. The supernatant was discarded; the plates were washed five times with tap
water and air dried. Sulforhodamine B (SRB) solution (50 µl) at 0.4 % (w/v) in 1 % acetic acid
was added to each of the wells, and plates were incubated for 20 minutes at room temperature.
After staining, unbound dye was recovered and the residual dye was removed by washing five
9

times with 1 % acetic acid. The plates were air dried. Bound stain was subsequently eluted with
10 mM trizma base, and the absorbance was read on an Elisa plate reader at a wavelength of 540
nm with 690 nm reference wavelength. Percent growth was calculated on a plate–by–plate basis
for test wells relative to control wells and was expressed as the ratio of average absorbance of the
test well to the average absorbance of the control wells*100. Using the six absorbance
measurements [time zero (Tz), control growth (C), and test growth in the presence of drug at the
four concentration levels (Ti)], the percentage growth was calculated at each of the drug
concentration levels. The dose response parameters were calculated for each test article. Growth
inhibition of 50 % (GI50) was calculated from [(Ti–Tz)/(C–Tz)] x 100 = 50, which is the drug
concentration resulting in a 50% reduction in the net protein increase (as measured by SRB
staining) in control cells during the drug incubation. The drug concentration resulting in total
growth inhibition (TGI) was calculated from Ti = Tz. The LC50 (concentration of drug resulting
in a 50% reduction in the measured protein at the end of the drug treatment as compared to that
at the beginning) indicating a net loss of cells following treatment is calculated from [(Ti–Tz)/Tz]
x 100 = –50.
3. Results and discussion
Organochalcogen (S/Se) functionalized chrysin ligands L1‒L4 and their corresponding RuII(η6–
p–cymene) complexes 1‒4 were synthesized and characterized by various spectroscopic
methods. The synthetic route for organochalcogen derivatives of chrysin ligands involved the
preparation of bromo–chrysin intermediate, L (Hu, Wang, Cheng, Pan and Ren, 2011). The
bromo–chrysin ligand, L was obtained by reacting chrysin with 1,2–dibromoethane in the
presence of K2CO3 as a base and acetone as solvent. Both sulphur and selenium functionalized
chrysin ligands, L1‒L4 were synthesized by reacting bromo–chrysin (L) with different
organochalcogenolate anions (Scheme I). In case of thio–functionalized chrysin ligands L1 &L3,
thiolate anions were generated via reaction of respective thiols with sodium under a nitrogen
atmosphere, while seleno‒tethered ligands L2 & L4, were obtained from organoselenolate anions
generated in situ from the respective diorganyl diselenide in the presence of sodium borohydride.
10

Scheme I. Synthetic route of modified chrysin ligands, L1–L4.
Organoruthenium(II)–arene complexes 1–4 were synthesized by reacting ligands L1‒L4 with
[Ru(η6–p–cymene)Cl2]2 salt in presence of NaOMe (used for deprotonation of the hydroxyl
group of chrysin moiety) (Scheme II). The complexes were found air stable and soluble in
common organic solvents. The proposed structures were formulated on the basis of various
spectroscopic techniques viz., IR, NMR (1H, 13C and 77Se) and HR–MS.
11

Scheme
II:
Synthesis
of
Ru(II)–arene
complexes
1–4.
The comparative IR spectra of the ligands, L1‒L4 and their corresponding organoruthenium(II)
complexes1–4 provided intricate details about the binding behavior and coordination mode with
the metal ion. IR spectra of ligands, L1‒L4 exhibited broad band at ~3456 cm–1 assigned to
hydroxyl ‒OH group which was found absent in complexes, implicating the coordination of
hydroxyl group with the metal ion via deprotonation (Raut and Wender, 1959). Another
important feature observed in the IR spectra of complexes 1−4 was significant lower frequency
shift (~15 cm–1) in the stretching vibrations of ν(C=O), which suggested the chelation of chrysin
unit of ligand as a O,O′‒bidendate donor to the Ru(II) centre (Pettinari et al., 2005).
The 1H NMR spectra of complexes1‒4 recorded in CDCl3 revealed the absence of –OH signal at
~δ 12.7 ppm indicating the deprotonation of phenolic –OH group on coordination to Ru(II)
centre. The spectra of complexes 1–4, further exhibited two signals as triplet centered at ~δ 4.20
and ~δ 2.90 ppm, corresponding to OCH2CH2 and OCH2CH2, respectively. Additionally, all
complexes exhibited signals at ~δ5.54 and ~δ5.26 ppm as doublet, corresponding to protons of
p–cymene moiety. However, complexes 1 and 2 exhibited multiplet signals in the range of δ
3.28–0.93 ppm assigned to αCH2, βCH2, γCH2 and‒CH3 protons of butylchalcogenide
substituent, merged with the characteristic CH3 and CH protons of p–cymene moiety. While, for
complexes 3 and 4the aromatic signatures of the phenylchalcogenide protons were found to be
merged in the range of δ 8.82–7.21 ppm (Fig. S1).
In13C NMR spectra of the complexes, appreciable downfield shift from 182.4 to 178.1 ppm of
C=O carbon was observed as a consequence of the coordination to Ru(II), while the O–C–Ar
12

carbons of the chrysin appeared in the range ofδ167.73–98.64 ppm. Furthermore, complexes 1–4
displayed signature carbons of p–cymene moiety at18.07, 24.08, 30.86, 78.91, 82.33, 90.60 and
97.01 ppm, implicating the coordination of Ru(II)–p–cymene fragment. Other peaks
corresponding to the OCH2CH2 and OCH2CH2 carbons were also observed at 32.7 and 69.0 ppm,
respectively in all the complexes 1–4 (Fig. S2).
In order to provide further evidence, 77Se NMR spectra of selenium bearing entities were
recorded in CDCl3. The ligands, L2 and L4 exhibited a sharp singlet at 144.14 and 271.52 ppm,
respectively which shifted marginally to 142.65 and 269.84 ppm in Ru(II) complexes 2 and 4
(Fig. S3), excluding any possibility of coordination of organoseleno moiety to Ru(II) metal
centre.
HR–MS spectra of the ligands, L1‒L4 displayed prominent peaks at m/z 419.06, 439.03, 371.13
and 391.10, respectively corresponding to the [M+H]+ molecular ion fragment (Fig. S4) whereas
the spectra of RuII complexes 1–4 were dominated by species assigned to [M‒Cl]+ at m/z 653.07,
673.03, 605.12 and 625.09, respectively (Fig. S5).
The UV–Vis spectra of complexes 1–4 were recorded in DMSO at room temperature exhibited
characteristic higher energy intense absorption band in the range of 272–276 nm which could be
attributed to the intraligand π→π* transitions. Other low energy broadband in the spectra of
complexes observed at 416‒475 nm arise from π→π* (MLCT) transitions due to the interaction
of filled metal orbitals of low spin d6 Ru(II) ion with the π* orbitals of the ligand.
3.1. DNA Interaction Studies
Genomic DNA is one of the primary intracellular target (Hosoya and Miyagawa, 2014) for
numerous FDA–approved inorganic (cisplatin, carboplatin, oxaliplatin etc.) and organic
(doxorubicin, gemcitabine, 5–fluorouracil etc.) chemotherapeutic drugs which exert their
anticancer effect by damaging the DNA replication (Kumar, Dasari and Patra, 2017;
Tanzadehpanaha et al., 2018). During the past decade, a significant amount of reserch work has
been directed on the DNA–binding ability of organometallic Ru(II) complexes (Caruso et al.,
2014; Frik et al., 2014) which have indicated differences in mode of DNA interaction depending
on their structure. DNA can provide three distinct binding modes for metal complexes viz.,
intercalation, groove binding and electrostatic binding to phosphate backbone of DNA. Usually,
intercalation within the base pairs of DNA results in hypochromism with or without red/blue
shift, while non–intercalative, electrostatic interaction with DNA double helix causes
13

hyperchromism (Liu et al., 2010).The interaction of complexes 1–4 with DNA causes electronic
perturbations in these complexes which can be observed by UV–vis absorption, fluorescence and
CD spectroscopic studies.
3.1.1. UV–vis absorption studies
The UV–vis spectra of RuII(η6–p–cymene)complexes 1–4 exhibited an intense absorption band
centered at ~274 nm attributable to π→π* intraligand (IL) transitions. On addition of increasing
concentration of CT DNA (0.1–1.0 x 10–6M) to a fixed concentration of complexes 1–4 (0.2 x
10–6M), significant hyperchromism along with blue shifts were observed (Fig. 2).
(a)
(b)
(c)
(d)
Fig. 2. Absorption spectra of complexes (a) 1, (b) 2, (c) 3 and (d) 4 (0.2 x 10–6 M) in the presence
of increasing concentrations of CT DNA (0.1–1.0 x 10–6 M), obtained in Tris–HCl buffer (pH
7.2).
The observed ‘hyperchromic effect’ along with a blue shift may be attributed to non–covalent
interaction preferably via external contact due to electrostatic binding to DNA helix (Long and
14

Barton, 1990; Pasternack, Gibbs and Villafranca, 1983). To quantify the extent of binding of
Ru(II) complexes 1–4 to CT DNA, the intrinsic binding constant, Kb values were determined by
monitoring the changes in the absorbance at the corresponding λmax with increasing
concentrations of CT DNA. Shifting in the position of absorption bands and binding constant, Kb
values were calculated from Wolfe–Shimer equation (1), and are summarized in Table 1. From
the binding constant values it is inferred that, among the chrysin Ru(II) complexes
organoseleno– derivatives 2 and 4 exhibited higher binding propensity towards CT DNA in
comparison to organothio– derivatives 1 and 3, following the order 2 > 4 > 1 > 3, thereby
suggesting significant effect of organoselenium moiety on DNA binding interaction. Moreover,
efficient binding of butyl derivatives in comparison to their phenyl derivatives is attributed to the
increased lipophilicity of the complex due to the presence of aliphatic chain (Batchelor,
Paunescu, Soudani, Scopelliti and Dyson, 2017).
Table 1: Binding constant values (Kb) for interaction of complexes 1–4 with DNA.
Absorption λmax (nm)
Complexes
Free
(nm)
272
Bound
(nm)
268
Δ
Hyperchromicity
Kb (M–1)
(nm)
(%)
28
4
5
5
6
3.86(±0.09) x 105
6.35(±0.12) x 105
1.73(±0.07) x 105
4.71(±0.11) x 105
1
2
3
4
274
269
24
273
268
30
276
270
36
3.1.2. Fluorescence spectroscopy
Further insight in the interaction mode of complexes with DNA, fluorescence studies were
carried out. The emission spectra of complexes 1–4 displayed an intense luminescence at ~350
nm in Tris–HCl buffer at room temperature in the absence of DNA when excited at 270 nm. The
concomitant addition of CT DNA (0–1.2 x 10–6 M) to a fixed concentration (0.2 x 10–6 M) of
complexes 1–4, resulted a significant enhancement in the fluorescence emission intensity with no
apparent change in the shape and position of the emission bands (Fig. 3).
15

(a)
(b)
(c)
(d)
Fig. 3. Fluorescence spectra of complexes (a) 1, (b) 2, (c) 3 and (d) 4 (0.2 x10–6 M) in the
presence of increasing concentrations of CT DNA (0–1.2 x 10–6 M) in Tris–HCl buffer (pH 7.2).
The observed enhancement in emission intensity is related to the extent with which that
complexes are protected inside the hydrophobic environment of DNA helix. Therefore, the
binding of complex to DNA restricts the complex mobility at the binding site due to the
inaccessibility of the solvent water molecules, leading to decrease in vibrational mode of
relaxation and thus avoiding the quenching effect of solvent molecules. The quantitative
assessment of DNA binding strength of the complexes 1–4 was ascertained by binding constant,
K as quatified by Scatchard equation (2 & 3) and were found to be 4.73 (±0.05) x 105, 7.68
(±0.11) x 105, 2.04 (±0.07) x 105 and 5.81 (±0.15) x 105 M–1, respectively, consistent with
absorption spectral studies.
16

3.1.3. Circular dichroism
Circular dichroic (CD) spectroscopy is a powerful technique for monitoring the morphological
and conformational transitions in proteins and nucleic acids upon interaction with small
molecules. The conformational changes in DNA structure upon interaction with complexes 1–4
was studied by circular dichroism (CD) spectra. The CD spectrum of CT DNA consists of a
positive band at 275 nm due to base stacking and a negative band at 245 nm, attributed to right–
handed helicity of B–DNA. Simple groove binding and electrostatic interaction of the complexes
with DNA showed less or no perturbations on the base stacking and helicity bands, while
intercalation can stabilize the helix conformation of B–DNA, and enhance the intensities of both
the CD bands (Norden and Tjerneld, 1982). In our experiments, the CD spectra of CT DNA in
presence of complexes 1–4 demonstrated decrease in the intensities of both the negative and
positive bands without any shift in the band position (Fig. 4). These results suggested that the
complexes induce distortion of the secondary structure of B–DNA without the alteration in its
conformation from B– to A–form or B– to Z–form. However, CD spectral changes clearly ruled
out the intercalative mode of binding and demonstrated non–covalent interaction between the
complexes and DNA, probably via electrostatic mode. Furthermore, the change in intensity
followed the order 2 > 4 > 1 > 3, which were consistent with the results obtained by UV–vis and
fluorescence spectroscopy.
Fig.4. CD spectra of CT DNA alone (blue), CT DNA + 1 (orange), CT DNA + 2 (green), CT
DNA + 3 (black) and CT DNA + 4 (red) in Tris–HCl buffer (pH = 7.2). [Complex] = 10–6 M,
[DNA] = 10–6 M.
17

3.2. HSA Interaction Studies
HSA is identified as the primary transporter of various pharmaceutical drugs particularly
following intravenous administration (Sarkar, 1989). It has been reported that interactions of
drug entities with HSA most likely involve non–covalent forces which increases their circulatory
half–lives in the blood and improves biodistribution throughout the body. Hence, interaction of
Ru(II) complexes with serum protein is recognized as an important aspectin determining their
pharmacological properties viz., the absorption, transportation, distribution, metabolism,
excretion and efficacy of these metal based drugs (Canovic et al., 2017; Karami, Lighvan,
Farrokhpour, Jahromi and Momtazi-borojeni, 2017). Keeping in view the immense importance
of drug‒HSA interaction in the area of pharmacology, the interaction of R(II)‒arene complexes
with HSA were carried out using UV–vis spectroscopy, fluorescence spectroscopy and circular
dichroism (CD) studies.
3.2.1. UV–vis absorption studies
HSA protein exhibited a characteristic absorption peak at~280 nm attributed to aromatic rings in
tryptophan (Trp–214) tyrosine (Tyr–411) and phenylalanine (Phe) amino acid residues. A fixed
concentration of HSA (2.0 x 10–6 M) was titrated with increasing concentration of complexes 1–
4(0–4.0 x 10–6 M) (Fig. 5). The complexes exhibited hyperchromism at λmax~280 nm along with
blue shift, indicating that more aromatic residues were extended into the aqueous environment
(Wang et al., 2011). Further, the observed “hyperchromism” suggested the interaction between
HSA and complexes due to the formation of ground state complex of the type HSA‒
Ru(II)complex most likely by electrostatic attraction between the Ru(II) complexes and HSA.
Overall, the changes in the absorbance spectra indicated that the microenvironment of the three
aromatic residues was altered causing prominent perturbation in tertiary structure of HSA.
Meanwhile, binding constant, Kb values for the complexes 1–4 were found to be 3.91(±0.12) x
104, 7.23(±0.09) x 104, 2.16 (±0.15)x 104 and 5.70(±0.07) x 104 M–1, respectively suggesting the
moderate binding propensity with HSA followed by conformational changes in its structure.
Since moderate affinity of a drug towards a protein helps in the diffusion of the drug from the
circulatory system to reach its target site, thus obtained Kb values for complexes fulfill this
criterion for their transport in the blood circulation and diffusion at the target site. Further,
binding affinity of 1–4 falls in the order 2 > 4 > 1 > 3, which supports that repalcing the phenyl
18

entity by butyl group at the chalcogen moiety of the chrysin increases the electron donating
ability of the substituent which consequently increases the protein binding ability of the complex.
(a)
(b)
(c)
(d)
Fig. 5. Absorption spectra of HSA (2.0 x 10–6 M) in the presence of increasing concentrations of
complexes (a) 1, (b) 2, (c) 3 and (d) 4 (0–4.0 x 10–6 M) in Tris–HCl buffer (pH 7.2). Arrows
indicate the absorbance changes upon increasing concentration of complex.
3.2.2. Fluorescence spectroscopy
HSA is a polypeptide chain with 585 amino acids, of the amino acids single tryptophan residue
(Trp–214) is mainly responsible for the majority of the intrinsic fluorescence of the protein. HSA
exhibits strong fluorescence emission peak at ~350 nm due to tryptophan residues, when excited
at ~290 nm (Lakowicz, 2006) wavelength. Quenching in the fluorescence emission spectra of
tryptophan in HSA are primarily due to changes in protein conformation, subunit association,
19

substrate binding or denaturation. The concomitant addition of RuII(η6–p–cymene) complexes 1–
4 (0.0–4.0 x 10–6 M) to a fixed amount of HSA (1.0 x 10–6 M), resulted in quenching of the
intrinsic fluorescence intensity of HSA at ~350 nm along with a blue shift of ~3 nm (Fig. 6). The
observed quenching may be attributed to changes in tryptophan environment of HSA due to the
binding of complexes to the albumin, leading to conformational alterations in the tertiary
structure of HSA (Rajendiran et al., 2007). Fig. 7 represents the Stern–Volmer quenching plot of
the complexes with HSA.
(a)
(b)
(c)
(d)
Fig. 6. Fluorescence spectra of HSA (1.0 x 10–6 M) in the presence of increasing concentrations
of complexes (a) 1, (b) 2, (c) 3 and (d) 4 (0–4.0 x 10–6 M), obtained in Tris–HCl buffer (pH 7.2)
upon excitation at 290 nm.
The quenching can be either via static or dynamic mechanism, which was validated by carrying
out temperature‒dependent fluorescent experiments. Static quenching refers to fluorophore–
20

quencher complex formation in the ground state, while in dynamic quenching the interaction
between fluorophore and quencher is during transient existence of the excited state
(Ramachandran, Raja, Bhuvanesh and Natarajan, 2012; Raja, Bhuvanesh and Natarajan, 2011).
If the quenching constant (Ksv) increased by the increase in temperature then mechanism of
quenching is dynamic since at a higher temperature molecules results in rapid diffusion, leading
to decrease in collision probability. On the other hand, if Ksv decreases by the increase in
temperature, then mechanism of quenching is static because the static quenching is caused by the
formation of complex and an increase in temperature lowers the stability of the complex and the
value of quenching constant.
Fig. 7. Stern–Volmer plot for quenching of the complexes 1 (cyan), 2 (green), 3 (red) and 4
(magenta) with HSA.
The fluorescence quenching experiments were performed at two different temperatures viz., 300
and 310 K. As depicted in Table 2, an inverse correlation between the Stern–Volmer quenching
constant (Ksv) values and temperature was noticed. Moreover, the biomolecular quenching rate
constant (Kq) values for all Ru(II) complexes were much greater than the maximum value of
dynamic quenching (2.0 x 1010 mol L‒1 s‒1), suggesting that the fluorescence quenching of HSA
was triggered viathe static quenching mechanismthrough the formation of a complex (Zhao, Liu,
Chi, Teng and Qin, 2010). The number of binding sites (n) were calculated ~1, which strongly
supported the existence of a unique binding site in HSA for all complexes. Comparing the
affinity of complexes 1‒4 for HSA, it is inferred that 2 exhibited higher affinity for HSA.
21

Further, the values of the quenching rate constant (Kq) and binding constant (K) for complexes
1‒4 were found to be similar to the values reported for ruthenium(II) complexes (Kljun et al.,
2013).
Table 2: Binding and thermodynamic parameters of complex–HSA system at different
temperatures obtained from fluorescence quenching experiments (pH 7.2).
T (K)
300
310
300
310
300
310
300
310
Ksv (M–1)
1.93 x 104
0.73 x 104
5.15 x 104
3.63 x 104
1.38 x 104
0.51 x 104
2.74 x 104
1.99 x 104
kq (M–1s–1)
1.20 x 1012
0.85 x 1012
4.45 x 1012
3.89 x 1012
0.91 x 1012
0.67 x 1012
3.51 x 1012
2.28 x 1012
K(M–1)
Complexes
1
n
3.90 x 104
2.07 x 104
13.54 x 104
11.21 x 104
1.89 x 104
0.73 x 104
9.68 x 104
7.53 x 104
0.97
1.34
0.80
1.02
2
3
4
3.2.3. Thermodynamic parameters
The noncovalent interactions with protein are of four types, which are hydrogen–bonding,
hydrophobic interaction, van der Waals interaction and ionic interaction (Moradi, Khorasani–
Motlagh, Rezvani, and Noroozifar, 2018). The signs and magnitude of thermodynamic
parameters viz., change in enthalpy (ΔH), entropy change (ΔS), and free energy change (ΔG), are
the main evidence to determine the interaction mode between Ru(II) complexes and HSA. The
thermodynamic parameters, ∆H > 0 and ∆S > 0 imply a hydrophobic interaction; ∆H < 0 and ∆S
< 0 reflect the van der Waals or hydrogen bond formation and ∆H < 0 and ∆S > 0 suggestive of
electrostatic interactions. The calculated thermodynamic parameters are summarized in Table 3,
showing the negative ∆H and positive ∆S values for the complexes which suggested that the
complexes bind to HSA through van der Waals force or hydrogen–bonding. Nevertheless
probability of binding of phenyl appended analogs 3 and 4 to HSA via hydrophobic interaction
cannot be ruled out (Fu et al., 2014). Moreover, all the complexes showed negative values of ∆G,
22

implicating the spontaneous binding process and the lower ∆G values for 2 supported its higher
binding affinity towards HSA.
Table 3: Thermodynamic parameters of complex–HSA interaction at pH 7.2.
∆G
(KJ mol–1)
–23.86
–24.43
–29.18
–29.82
–21.86
–22.39
–26.36
–26.95
∆H
∆S
T (K)
Complexes
(KJ mol–1K–1)
(J mol–1K–1)
300
310
300
310
300
310
300
310
–6.60
–9.91
–5.80
–8.43
57.53
64.23
53.53
59.76
1
2
3
4
3.2.4. Circular dichroism
CD measurements were performed in the presence of RuII(η6–p–cymene) complexes 1–4 to
examine its possible impact on the secondary structure of HSA. As depicted in Fig. 8, the CD
spectrum of HSA revealed two negative bands characteristic of α–helical proteins at 208 and 222
nm assignable to π→π* and n→π* transitions, respectively. The ellipticity of HSA decreased
moderately on the addition of complexes, indicating some loss of α–helical secondary structure
(Divsalar, Bagheri, Saboury, Mansoori–Torshizi and Amani, 2009). This may imply that
complexes could interact with the amino acid residues of the main polypeptide chain of HSA.
However, the shape and position of CD spectra remained almost unchanged in presence and
absence of complexes, suggesting that conformation of HSA was predominantly in α–helix even
after binding to the complexes.
23

Fig. 8. CD spectra of HSA alone (black), HSA + 1 (pink), HSA + 2 (red), HSA + 3 (blue) and
HSA + 4 (green) in Tris–HCl buffer (pH=7.2). [Complex] = 10–6 M, [HSA] = 10–6 M.
3.2.5. Three–dimensional (3D) fluorescence spectral studies
Three–dimensional (3D) fluorescence spectroscopy is an analytical technique that provides the
information about the conformational changes of fluorophore by changing excitation and
emission wavelengths simultaneously (Zhang et al., 2008). The presence of peak a reflects
Rayleigh scattering (λex = λem) while peak b denotes secondary scattering peak (λem = 2λex).
Peak 1 corresponds mainly to the spectral behavior of Trp and Tyr residues (λex ~ 280, λem ~ 346
nm). Any alteration in the microenvironment of Trp and Tyr residues of HSA upon interaction
with complex implicates an obvious change in either intensity or wavelength of the spectra. By
comparing the 3D fluorescence spectral changes of HSA in the absence and presence of Ru(II)
complexes 1–4, it was observed that the fluorescence intensities of peak 1 decreased
significantly indicating a strong quenching of fluorescence induced by Trp residue of HSA (Fig.
S6). However, the quenching phenomena was more pronounced in case of complex 2 indicating
its stronger affinity in comparison to other complexes and hence suggested more perturbation in
hydrophobic microenvironment near the tryptophan and tyrosine residues leading to
conformational changes in HSA.
24

3.3. Molecular docking studies
3.3.1. Molecular docking with DNA
Molecular docking studies of synthesized RuII(η6–p–cymene)complexes 1–4 with DNA duplex
of sequence d(CGCGAATTCGCG)2 dodecamer (PDB ID: 1BNA) were performed in an attempt
to provide vital information regarding the preferred orientation of the complexes inside the DNA
grooves. The favorable energy minimized docked poses of n–butylseleno complex, 2 revealed
that it interact adjacent to AT–rich sequence of the minor groove and was in close proximity to
base pairs T7, T8, A18, T19 and T20 of the targeted DNA (Fig. 9), while its phenylseleno–
analog, 4 showed the interaction in GC–rich region of minor groove adjacent to base pairs C9,
G10, C15 and G16 (Fig. S7). In general, electronegative AT sequences provide better van der
Waals contacts since they are narrower compared to GC regions and therefore expose better
fitting for docking of small molecules (Sahoo, Ghosh, Bera and Dasgupta, 2008). Moreover, the
stabilization could also be enhanced by extensive van der Waals interaction and hydrophobic
contacts in the groove region. Relative binding energy of the docked structures was found to be–
278.2, –350.9, –243.6 and –312.1KJ mol–1 for1–4, respectively, implicating the higher binding
affinity of n–butylseleno complex 2 as compared to other analogs.
Fig. 9. Molecular docked model of complex 2 in the minor groove of DNA dodecamer duplex
of sequence d(CGCGAATTCGCG)2 (PDB ID:1BNA).
25

3.3.2. Molecular docking with HSA
RuII(η6–p–cymene) complexes 1–4 have demonstrated higher binding affinity towards HSA
target, and albumins proteins have been identified as the main in vivo carrier molecules, so it was
imperative to carry out molecular docking studies of RuII(η6–p–cymene) complexes with HSA
(PDB ID: 1H9Z). Crystal structure descriptions of the 3D structure of HSA revealed a single
polypeptide chain of 585 amino acid residues comprises three structurally homologous domains
(I–III): I (residues 1–195) II (196–383) and III (384–585); each domain has two subdomains (A
and B) that assemble to form heart shaped molecule (He and Carter, 1992; Yamasaki, Chuang,
Maruyama and Otagiri, 2013). The principal region of drug binding sites of HSA are located in
hydrophobic cavities in subdomain IA and IIA, corresponding to site I and III, respectively and
tryptophan residue (Trp–214) of HSA in subdomain IIA. The molecular docking pattern of 2
with HSA indicated that complex was located in the hydrophobic cavity between subdomains
IIA and IIIA, implicating the existence of hydrophobic interactions which also results in the
fluorescence quenching of Trp–214.
Fig. 10. Molecular docked model of (a) interaction between complex 2 and amino acid residues of
HSA (b) complex 2 with HSA and (c) representation of 2 located within the hydrophobic pocket in
subdomain III A and II A of HSA.
26

As clearly depicted in Fig. 10, n–butylseleno complex 2 was in close proximity of the amino acid
residues such as Arg–197, Lys–205, Cys–200, Gln–204, Ala–201, Typ–148, Cys–246, Ala–194,
His–247, Asn–458 and Val–462. Our results further revealed that hydrophobic interactions
played a major role in 2–HSA interaction, however hydrogen bonding between Arg–197and
Lys–205 residues with O1/2 of O,O– chelating motif of 2 offered additional stability. Further,
molecular docking experiments were also performed with phenylseleno derivative, 4 which
revealed that complex was found in vicinity of residues viz., Lys–162, Ala–158, Lue–155 and
Lue–121 (Fig. 11). Nevertheless, complex 2 revealed more negative binding energy values
towards HSA target than complex 4 thereby showed more binding affinity which correlated well
with the in vitro binding experiments.
Fig. 11. Molecular docked model of complex 4 with HSA within the hydrophobic pocket
in subdomain I B and I A of HSA and its interaction with amino acid residues of HSA.
3.4. In vitro anticancer activity
The cytotoxicity profile of RuII(η6–p–cymene) complexes 1‒4 were carried out at different molar
concentrations in comparison to conventional standard drug Adriamycin (ADR), on selected
panel of cancer cell lines viz., HeLa (cervical), MIA–PA–CA–2 (pancreatic), MCF–7 (breast),
Hep–G2 (Hepatoma), SK–OV–3 (ovarian) by Sulforhodamine B (SRB) assay. Fig. 12 shows the
27

antitumor screening images of different human carcinoma cell lines which demonstrated the
good to moderate activity of complexes towards certain specific cancer cell lines.
Fig. 12. Images for the in vitro cytotoxicity of Ru(II) complexes 1‒4 on selected cancer cell lines
viz., HeLa (cervical), MIA–PA–CA–2 (pancreatic), MCF–7 (breast), Hep–G2 (Hepatoma), SK–
OV–3 (ovarian).
The anticancer screening results showed that all the complexes were effective against HeLa
(cervical) cancer cell lines at concentration of 10–4 M, although the complexes did not exhibit
any detectable cytotoxicity at concentration of 10–7 M as compared to Adriamycin. Moreover,
ourinitial cytotoxic screening data revealed that complex 2 exhibited comparatively better
selectivity and cytotoxicity against certain human cancer phenotypes viz., HeLa, MIA–PA–CA–
2and MCF–7 at 10–4 M which could be presumably due to its higher DNA/HSA binding affinity.
4. Conclusions
Considering the chemotherapeutic effects of chrysin ligand and biological effects of
organochalcogen functionalities (S/Se) we have designed hybrid organometallic Ru(II)
complexes and explored the influence of these motifs on the anticancer activity as well as on
28

their modes of action. This work features the design and synthesis of four new RuII(η6–p–
cymene) complexes 1–4 derived from organochalcogen (S/Se) functionalized chrysin ligand
pharmacophores which were thoroughly characterized by various spectroscopic and analytical
techniques. The comparative in vitro interaction studies of complexes 1–4with DNA/HSA targets
were carried by employing UV–vis absorption, fluorescence spectroscopy and circular dichroic
techniques to validate their chemotherapeutic candidature. Our results demonstrated that RuII(η6–
p–cymene) complexes 1‒4 exhibited high binding propensity towards CTDNA via electrostatic
mode and the observed trend in the binding interactions revealed that binding propensity of
complexes showed structural dependence i.e, seleno– derivatives constitute attractive scaffold
and higher binding affinity than their thio– derivatives. Further, mode of interaction of the
complexes with DNA/HSA was validated by molecular docking studies, which complemented
our spectroscopic observations. The cytotoxicity activity of RuII(η6–p–cymene) complexes 1‒4
were carried out by SRB assay against selected human cancer cell lines viz., HeLa (cervical),
MIA–PA–CA–2 (pancreatic), MCF–7 (breast), Hep–G2 (Hepatoma), SK–OV–3 (ovarian) which
demonstrated good activity of all complexes on HeLa cancer cell line, whereas moderate activity
on other cell lines, implicating the selective response towards different cancer phenotypes. The
results revealed superior cytotoxicity of complex 2 as compared to other complexes on selective
n
cancer phenotypes suggesting that tethering of Bu chain over its Ph ring could effectively tune
its biological properties.
Supplementary material
Additional supplementary material can be found in the online version of this article.
Acknowledgments
The author S. Yadav thanks Science & Engineering Research Board (SERB) a statutory body of
the Department of Science & Technology (DST) for the financial support under project grant
PDF/2016/002767. The authors gratefully acknowledge Prof. Yoshiaki Nishibayashi, University
of Tokyo, Japan for his valuable inputs in accomplishing this work. The authors are thankful to
Department of Chemistry, IIT Delhi for instrumentation facility and Anti–Cancer Drug
Screening Facility (ACDSF) at ACTREC, Mumbai for carrying out the in vitro anticancer
activity.
29