Ruthenium complexes of ferrocene mannich bases: DNA/BSA interactions and cytotoxicity against A549 cell line

Article abstract of DOI:10.1016/j.jphotochem.2015.02.010

Two different series of ruthenium complexes, ML₃ and MLL′₂ where M = Ru(III)/Ru(II), L = ferrocenyl amino acid mannich base conjugates and L′ = 1,10-phenanthroline have been synthesized and characterized by spectroscopic methods. The

Full text of DOI:10.1016/j.jphotochem.2015.02.010

Journal of Photochemistry and Photobiology A: Chemistry 305 (2015) 1–10
Contents lists available at ScienceDirect
Journal of Photochemistry and Photobiology A:
Chemistry
journal homepage: www.elsevier.com/locate/jphotochem
Ruthenium complexes of ferrocene mannich bases: DNA/BSA
interactions and cytotoxicity against A549 cell line
Pulipaka Ramadevi a, Rinky Singh a, Sarmita S. Jana b, Ranjitsinh Devkar b,
Debjani Chakraborty a,
*
a
Department of Chemistry, Faculty of Science, The M.S. University of Baroda, Vadodara-390002, Gujarat, India
Division of Phytotherapeutics and Metabolic Endocrinology, Department of Zoology, Faculty of Science, The M.S. University of Baroda, Vadodara-390002,
b
Gujarat, India
A R T I C L E I N F O
A B S T R A C T
Two different series of ruthenium complexes, ML3 and MLL02 where M = Ru(III)/Ru(II), L = ferrocenyl
amino acid mannich base conjugates and L0 = 1,10-phenanthroline have been synthesized and
characterized by spectroscopic methods. These ferrocenyl ligands and their ruthenium complexes have
been investigated for their interactions with DNA and BSA (bovine serum albumin) employing steady-
state fluorescence quenching measurements, UV–vis spectroscopy and DNA viscosity measurements.
High binding constants obtained from the DNA binding studies (Kb = 104 – 106 Mꢀ1) prompted the in-vitro
cytotoxicity assay of complexes on A549 human lung carcinoma cells (employing MTT assay). The IC50
Article history:
Received 4 September 2014
Received in revised form 30 January 2015
Accepted 12 February 2015
Available online 25 February 2015
Keywords:
Ferrocenyl mannich base
Ruthenium complexes
DNA/BSA binding interactions
Cytotoxicity
values (within the range of 46
well known ruthenium complex NAMI-A currently under phase II clinical trials which has IC50 values in
the range of 550 M–750 M for various cancer cell lines. Interaction of these complexes with A549 cells
mM–422 mM) obtained herein were found to be lower than those of the
m
m
has been further scrutinized using acridine orange (AO)/ethidium bromide (EB) dual staining technique
to indicate apoptosis as the mode of cell death.
A549 human lung carcinoma
Apoptosis
ã 2015 Elsevier B.V. All rights reserved.
1. Introduction
[2], mefloquine [3] and artemisinin [4]. Moreover ferrocene
derivatives have been used as scaffolds to design new molecules
Ferrocene is one of the members of the well known
organometallic family- “metallocenes” in which an iron atom is
flanked by two cyclopentadienyl (Cp) rings. The unique structural
and conformational properties of ferrocene, an atomic ball bearing
character, are characterized by the parallel alignment of the two
cyclopentadienyl rings and the free rotation of the rings around the
axis penetrating their centers. Its remarkable stability and
fascinating chemistry has attracted the attention of the scientific
and technical community. Owing to the favorable electronic
properties of ferrocene and its easy functionalization their
applications have been explored in a wide range of scientific
areas ranging from catalysis to the design of new nonlinear optic
materials to new biologically active compounds. Several structural
modifications of established drugs with ferrocenyl moiety have
been reported, such as ferrocene fluconazole [1], ferrocene aspirin
that recognize cations, anions, organic molecules, nucleoba-ses,
dinucleotides and amino acids [5–10]. Seio et al. [11] have reported
new DNA binding molecules utilizing structural and conforma-
tional properties of ferrocene. Their design concept is based on the
fact that the distance between the two cyclopentadienyl rings of
ferrocene, ca. 3.3 Å is close to the distance between two aromatic
rings stacked with each other. In addition, it is well-known that the
minor groove of DNA can accommodate stacked aromatic rings, as
established by the structural studies of natural or synthetic
molecules that recognize the minor groove. Therefore, it was
expected that ferrocene derivatives could be a new type of DNA
binding molecules if appropriately designed aromatic rings were
attached to the Cp rings. Recent studies on interaction of ferrocenyl
ligands with DNA has shown binding constants to be in the order of
103–105 [12] agreeing with the above mentioned fact. The stability
of ferrocene in aqueous and aerobic media has made ferrocenyl
compounds very popular molecules for such biological applica-
tions. Furthermore such favorable characteristics of ferrocene led
to the design of ferrocenyl derivatives that function as highly
* Corresponding author. Tel.: +91 9537521703.
E-mail address: debchak23@gmail.com (D. Chakraborty).
http://dx.doi.org/10.1016/j.jphotochem.2015.02.010
1010-6030/ã 2015 Elsevier B.V. All rights reserved.

2
P. Ramadevi et al. / Journal of Photochemistry and Photobiology A: Chemistry 305 (2015) 1–10
sensitive detectors of proteins or as reporters of protein activity.
Studies on ferrocenyl conjugates with amino acids and end-labeled
ferrocenyl di- and tripeptides have demonstrated distinctive
electrical, structural, and medicinal properties [13]. Many
researchers have shown interest in design of unnatural ferrocenyl
amino acids and peptides which further have been studied for their
biomedical applications. Modification of proteins by incorporating
such unnatural ferrocenyl amino acids helps the study of protein
structure, activity and interaction with other biomolecules [14].
The ferrocenyl compounds are known to hold good potential as
anticancer agents. This can be exemplified by the results obtained
by the group of Gérard Jaouen, with the development of ferrocifens
(i.e., ferrocene-modified tamoxifens), which exhibit strong anti-
proliferative effects not only in hormone dependent but also in
hormone-independent breast cancer cells [15]. A recent review by
Ornelas throws light on the various applications of ferrocene and
its derivatives in cancer research [16].
drugs, DNA interactions of antitumor ruthenium agents are of a
great interest [22,23].
Complexes of 1,10-phenanthroline (phen) have been shown to
exhibit interesting clinical activities including antitumoural,
antimycobacterial, antifungal and antimicrobial activities. More-
over, these compounds can be used as DNA intercalating agents
and metal synthetic nucleases [24].
The ferrocene amino acid derivatives have been coordinated to
Ru3+ and [Ru(phen)2]2+ and investigated for their interaction with
DNA and bovine serum albumin. Results obtained in the DNA
binding studies prompted the in-vitro cytotoxicity evaluation of
ruthenium complexes on A549 human lung carcinoma cells using
MTT assay. The interaction of these complexes with A549 cells has
been scrutinized using AO/EB staining technique to assess the
mechanism of cell death viz. apoptotic or necrotic.
2. Experimental
Here we are reporting a series of amino acid conjugated
ferrocene mannich bases. These ferrocenyl amino acids have been
targeted as it has been shown that tethering biologically active
groups to the ferrocenyl unit increases their potency, possibly due
to the combined action of the organic molecule with Fenton
chemistry of the Fe-center [17,18]. Since DNA represent very
important potential target of cancer diagnostic and chemotherapy
drugs, before going to expensive cell line study it was decided to
preliminarily explore the anticancer potency of the compounds by
determining their DNA interaction because ‘The control over DNA
can control the cancerous cell growth’. Like DNA, proteins are also
considered to be one of the prime molecular targets for diagnostic
and imaging agents, and so equal attention has been paid on
designing novel probes for proteins [19–21].
2.1. Reagents and materials
All the chemicals and solvents used for synthesis and
characterization of ligands and complexes are of analytical grade
and were used as purchased. Ferrocene carboxaldehyde and DAPI
stain were purchased from Sigma–Aldrich. Amino acids, CT-DNA,
Tri–sodium citrate and EB (ethidium bromide) were purchased
from SRL (Sisco Research Laboratory, Mumbai, India). RuCl3. 3H2O
and BSA (bovine serum albumin) were purchased from Hi media.
All the solvents used in the present studies were purchased from
Merck and are of analytical grade.
2.2. Methods and instrumentation
Several ruthenium compounds have been shown to inhibit DNA
replication, possess mutagenic activity, induce SOS repair, and
reduce RNA synthesis, which are all consistent with DNA binding of
these compounds in vivo. Thus, by analogy to platinum antitumor
1H NMR spectra of the ligands were recorded on Bruker
400 MHz NMR Spectrophotometer. ESI Mass spectra of the ligands
and complexes were recorded on Applied Biosystem API 2000
Fig. 1. (i) Dry MeOH, NaOH reflux, NaBH4, stirring at r.t. for 1.5 h, (ii) RuCl3.3H2O, DMF and reflux for 6 h, (iii) Ru(1,10-phenanthroline)2Cl2. 2H2O, EtOH and reflux for 7 h under
N2.

P. Ramadevi et al. / Journal of Photochemistry and Photobiology A: Chemistry 305 (2015) 1–10
3
Mass spectrometer. Infrared spectra (400–4000 cmꢀ1
)
were
The overall synthetic route of the ligands and complexes has
been presented schematically in Fig. 1.
recorded on PerkinElmer RX-1 FTIR with samples prepared as
KBr pellets. UV spectra were recorded in DMSO solution at
concentrations in the range 10ꢀ6–10ꢀ3 M on PerkinElmer Lambda-
35 dual beam UV–vis spectrophotometer. C, H and N elemental
2.5. DNA binding experiments
analysis were performed on
a
PerkinElmer 240B elemental
2.5.1. UV absorption studies
analyzer. Fluorescence spectra were recorded in solution on JASCO
FP-6300 fluorescence spectrophotometer.
The interaction of compounds with CT DNA has been studied
with UV spectroscopy in order to investigate the possible binding
modes to CT DNA and to calculate the binding constants (Kb).
Absorption studies were performed with fixed compound con-
centrations while varying the CT-DNA concentration within. Stock
solution of the ligands and complexes was diluted with tris buffer
2.3. Synthesis of ligands
The ferrocenyl amino acids L1–L4 were synthesized according
to a procedure reported by Goswami et al. [25].
to get the desired ligand concentration (75
mM) and complex
Amino acid (2.76 mmol) and NaOH (2.76 mmol) in dry methanol
(5 ml) were stirred for 30 min to get a homogeneous solution. A
methanolic solution (5 ml) of ferrocene carboxaldehyde
(2.76 mmol) was added dropwise to the above solution, which
was refluxed for 90 min, cooled, and treated with sodium
borohydride (5.2 mmol) with constant stirring. The solvent was
evaporated, the resulting mass was dissolved in water and acidified
with dilute HCl, and the solution pH was maintained within 5–6.
The ligand that precipitated as a yellow solid was filtered,
thoroughly washed with water and cold methanol, and finally
dried in vacuum. Yields of all the four ligands L1–L4 were in the
range of 80–90%.
concentration (37.5 M–50 M). While measuring the absorption,
m
m
equal increments of CT-DNA were added at different ratios to both
the compound solution and the reference solution to eliminate the
absorbance of CT-DNA itself.
2.5.2. Competitive binding studies with EB using fluorescence
spectroscopy
The competitive binding study of each compound with EB has
been investigated with fluorescence spectroscopy in order to
examine whether the compound can displace EB from DNA–EB
complex. The DNA–EB complex was prepared by adding EB
(33.3 mM) and DNA (20 mM) in tris buffer. The binding mode of
complexes C5–C8 with CT-DNA was studied by adding a certain
amount of a solution of each complex step by step into the solution
of the DNA–EB complex. The influence of the addition of each
complex has been obtained by recording the variation in the
fluorescence emission spectra of the DNA–EB complex. The
fluorescence intensities of EB bound to CT-DNA were measured
at 609 nm (524 nm excitation) after addition of different concen-
trations of the complexes at different ratios.
2.4. Synthesis of complexes
2.4.1. [Ru(L)3]
RuCl3 3H2O (0.2 mmol) and the ferrocenyl ligands (0.6 mmol)
ꢁ
were taken in 5 ml DMF and refluxed for 6 h. After reflux the
reaction mixture was cooled to r.t. and acetone (5 times the volume
of DMF) was added followed by overnight cooling at 0 ꢂC. The
brown precipitates of the complexes were then filtered off, washed
with acetone and ether and dried in oven at 50 ꢂC. Yields of all the
four complexes C1–C4 were in the range of 50–60%.
2.5.3. Competitive binding studies with DAPI using fluorescence
spectroscopy
The competitive binding study of C1–C4 with known groove
binder DAPI has been investigated with fluorescence spectroscopy.
2.4.2. [Ru(L0)2(L)]ClO4 (L0 = 1,10-phenanthroline)
[Ru(L0)2]Cl2
ꢁ
2H2O (0.2 mmol) and the ferrocenyl ligands
The DNA–DAPI complex was prepared by adding DAPI (5
mM) and
(0.2 mmol) were taken in 5 ml ethanol and refluxed for 7 h under
nitrogen. After reflux the reaction mixture was cooled to r.t. and
saturated aqueous solution of sodium perchlorate was added drop-
wise and stirred for 2 h followed by overnight cooling at 0 ꢂC. The
brown precipitates of the complex were then filtered off, washed
with water and ether and dried. Yields of all the four complexes
C5–C8 were in the range of 55–60%.
DNA (20 M) in tris buffer. The binding mode of complexes C1–C4
m
with CT-DNA was studied by adding a certain amount of a solution
of each complex step by step into the solution of the DNA–DAPI
complex. The influence of the addition of each complex has been
obtained by recording the variation in the fluorescence emission
spectra of the DNA–DAPI complex. The fluorescence intensities of
DAPI bound to CT-DNA were measured at 451 nm (340 nm
Fig. 2. Titration plot of complexes (a) C1 and (b) C5 with DNA. Insets: Plot of [DNA]/(eA–ef) versus [DNA] for (a) C1–C4 and (b) C5–C8. (For interpretation of the references to
color in the text, the reader is referred to the web version of this article.)

4
P. Ramadevi et al. / Journal of Photochemistry and Photobiology A: Chemistry 305 (2015) 1–10
excitation) after addition of different concentrations of the
complexes at different mixing (r) ratios.
instruments, Inc., Winooski, VT). The IC50 values were determined
by plotting the percentage viability versus concentration on a
logarithmic graph and reading off the concentration at which 50%
of cells remained viable relative to the control. Each experiment
was repeated at least three times to obtain mean values.
2.5.4. Viscosity measurements
Cannon–Ubbelohde viscometer maintained at
a constant
temperature of 32.0 ꢃ 0.1 ꢂC in a thermostat was used to measure
the relative viscosity of DNA (200
mM) solutions in the presence of
2.8. AO/EB staining technique (induction of apoptosis)
complexes C1–C8 (with [complex]/[DNA] ratio of 0, 0.04, 0.08, 0.12,
0.16, 0.20 and 0.24) in Tris–HCl buffer (pH 7.2). Digital stopwatch
with least count of 0.01 s. was used for flow time measurement
with accuracy of ꢃ0.1 s. The flow time of each sample was
measured three times and an average flow time was calculated.
A549 cells were grown in triplicates using a 12 well tissue
culture plate and allowed to acclimatize overnight. Next day, cells
were treated with C3 (100 mg/ml) and C5 (250 mg/ml) and
incubated for 16 h. After the incubation, cells were stained with
acridine orange and ethidium bromide dyes for 5 min in dark and
immediately washed three times with PBS. The cells were then
suspended in PBS and photographed on confocal LSM-710 fluores-
cence microscope [29].
Data are presented as (h/h0) h is
1/3 versus [complex]/[DNA], where
the viscosity of DNA in the presence of complex and h0 is the
viscosity of DNA alone. Viscosity values were calculated from the
observed flow time of DNA-containing solutions (t) corrected for
that of the buffer alone (t0),
h
= (t ꢀ t0)/t0 [26].
3. Results and discussion
2.6. BSA binding experiments
3.1. Characterization
2.6.1. Steady-state fluorescence spectroscopy
The protein-binding study was performed employing steady
state fluorescence spectroscopy. Tryptophan fluorescence quench-
ing experiments were carried out using bovine serum albumin
3.1.1. NMR spectroscopy
The 1H NMR spectra of ligands L3 and L4 show 9 proton
multiplet in the
d
range of 4–5 ppm that can be ascribed to the
= 1–2 pertains to the
(BSA, 8.3
m
M) in buffer (containing 15 mM tri-sodium citrate and
ferrocenyl ring protons. The singlet signals at d
150 mM NaCl at pH 7.0). The quenching of emission intensity of the
tryptophan residues of BSA at 343 nm was monitored in the
presence of increasing concentrations of ligands L1–L4 (0–
166 mM) and complexes C1–C8 (0–30 mM) as quenchers [27].
Fluorescence spectra were recorded from 300 to 500 nm at an
excitation wavelength of 296 nm.
N—H and COO—H proton of the amino acid. The spectrum of L3
(Fig. S1,a), apart from the above mentioned signals, also shows a 6
proton singlet at
(CH3)2 of leucine. Whereas the spectrum of L4 (Fig. S1,b) shows 5
proton multiplet in the range of 7–8 ppm ascribable to the
aromatic protons of the tryptophan and a one proton singlet at
= 11 ppm attributed to the indoyl N—H.
d= 1.2 ppm owing to two methyl groups—CH
d
d
2.7. Cytotoxicity
3.1.2. Mass spectrometry
Standard 3-(4,5-dimethylthiazole)-2,5-diphenyltetraazolium
bromide (MTT) assay was used [28]. A549 cells (5.0 ꢄ103 cells
wellꢀ1) were placed in 96-well culture plates (Tarson India Pvt.,
Ltd.) and grown overnight at 37 ꢂC in a 5% CO2 incubator.
Compounds to be tested were then added to the wells to achieve
The ESI-MS spectra of the ligands L1–L4 and their complexes
C1–C8 showed molecular ion peaks at m/z values equivalent to
their molecular weights. The m/z values of all the complexes are in
well agreement with the proposed composition (Fig. 1) and have
been tabulated in Table S1. All the mass spectra have been provided
as Supplementary material Fig. S2. Furthermore the composition
and purity of the complexes have been confirmed by their C, H, N
elemental analysis, data of which has been provided in Table S1.
final concentrations ranging from 10 to 500 mg/ml. Control wells
were prepared by addition of culture medium without the
compounds. The plates were incubated at 37 ꢂC in a 5% CO2
incubator for 24 h. Upon completion of the incubation, MTT dye
solution (prepared using serum free culture medium) was added to
each well to a final concentration of 0.5 mg/ml. After 4 h of
incubation with MTT, the culture media was discarded and the
wells were washed with Phosphate Buffer Saline (Hi-Media, India
Pvt., Ltd.), followed by addition of DMSO to dissolve the formazan
crystals so formed and subsequent incubation for 30 min. The
optical density of each well was measured spectrophotometrically
at 563 nm using Biotek-ELX800MS universal ELISA reader (Bio-Tek
Table 1
Kb values of ligands L1–L4 and their ruthenium complexes C1–C8.
Compound
lmax
Kb (Mꢀ1
)
L1
L2
L3
L4
C1
C2
C3
C4
C5
C6
C7
C8
208
208
209
207
207
208
207
207
264
264
265
266
6.7 ꢄ104
2.6 ꢄ104
1.4 ꢄ104
1.3 ꢄ104
2.4 ꢄ105
1.7 ꢄ105
5.3 ꢄ104
6.6 ꢄ104
5.3 ꢄ106
3.8 ꢄ 105
2.5 ꢄ105
1.1 ꢄ106
Fig. 3. Plot of Fluorescence emission intensity I versus wavelength
complex at different concentrations of C5. Inset: Stern–Volmer quenching plot of
CT-DNA–EB for C5–C8.
l for CT DNA–EB

P. Ramadevi et al. / Journal of Photochemistry and Photobiology A: Chemistry 305 (2015) 1–10
5
3.1.3. FTIR spectroscopy
other hand intercalation generally results in hypochromism and a
red shift (bathochromism) of the absorption band due to a strong
stacking interaction between an aromatic moiety of the ligand and
the base pairs of the DNA [33]. The UV spectra of ligands L1–L4
(10ꢀ9 M), complexes C1–C4 (10ꢀ9 M) and C5–C8 (10ꢀ6 M) have
been recorded in absence and presence of varying CT-DNA
concentration (1–50 ꢄ10ꢀ6 M) within. The absorption bands of
the ligands L1–L4 (Fig. S4) and complexes C1–C4 (Fig. 2a) centered
at 207–208 nm showed significant hypochromism and a slight red
shift, speculative of primarily groove binding nature of the
compounds whereas the absorption bands of complexes C5–C8
(Fig. 2b) centered at 264–266 nm also exhibited hypochromism
but with negligible red shift. DNA intercalators show a much larger
bathochromic shift and hypochromism of the spectral bands,
although the intercalative mode of binding of the complexes with
DNA cannot be ruled out completely [25].
The FTIR spectra of the complexes C1–C8 displayed character-
istic strong stretching bands at 1520–1580 cmꢀ1 and weaker bands
at 1490–1514 cmꢀ1 due to asymmetric and symmetric carboxylate
(COOꢀ) stretch respectively which were found as strong bands in
the fingerprint region at 1580–1610 cmꢀ1 in the spectra of free
ligands L1–L4. Moreover the distinct broad band at ꢅ3450 cmꢀ1
owing to the O—H stretching of free carboxylic acid group found in
the ligand is completely lost in the IR spectra of the complex
indicating complexation of the ligand with metal via the
carboxylate oxygen. Furthermore the medium secondary amine
N—H stretching bands found in the spectra of the free ligands L1–
L4 in the region of 2900–3000 cmꢀ1 was found to have a positive
shift in the region of 3080–3200 cmꢀ1 in C1–C8 indicating
complexation of the ligand with metal via the nitrogen of
secondary amine (mannich base). The presence of perchlorate as
the counter ion in the complexes C5–C8 is indicated by its nCl—O
stretching band in the range of 625–635 cmꢀ1 [30]. All the
important stretching values have been tabulated in Table S2.
The magnitude of binding strength to CT-DNA may be
determined through the calculation of binding constant Kb which
is obtained by monitoring the changes in the absorbance of the
compounds with increasing concentrations of CT-DNA. Kb is given
3.1.4. Electronic spectra
by the ratio of slope to the y intercept in plots [DNA]/(eA
–
ef) versus
The electronic absorption spectra of the ligands L1–L4 and
complexes C1–C8 in DMSO solution were recorded in the region
200–900 nm. The electronic spectra of free ligands displayed
[DNA] (Fig. 2, insets) according to Eq. (1) [34]
½DNAꢆ
½DNAꢆ
1
¼
þ
(1)
ð
eA
ꢀ
ef
Þ
ð
eb
ꢀ
ef
Þ
Kbðeb
ꢀ ef Þ
intense absorption bands at 207–209 nm ascribed to
p–p* intra
where [DNA] is the concentration of DNA in base pairs,
e
A = Aobsd/
ligand transition of the cyclopentadienyl rings of ferrocene (Fig. S3,
a) which were observed unchanged in the spectra of the complexes
C1–C4 (Fig. S3,b). The intra ligand transition bands of complexes
C5–C8 were observed at longer wavelength region at 278–282 nm
(Fig. S3,d) due to coordination with Ru(II). In addition all the
complexes showed peaks in the region 382–488 nm corresponding
[compound], ef is the extinction coefficient for the unbound
compound (L1–C8) and eb is the extinction coefficient for the
compound in the fully bound form. The binding constant Kb values
(Table 1) for the ligands are in the range of 6.7 ꢄ104–1.3 ꢄ104 Mꢀ1
indicative of strong binding of the ligands with ligand L1 showing
the highest binding constant. The Kb values of metal complexes
showed 10–100 folds higher binding efficacy compared to the
ligands. In general, complexes C5–C8 with two 1,10 phenanthroline
moieties bound to the metal center, exhibit stronger binding
to dp–p* MLCT transitions (Fig. S3,c and d). Furthermore,
complexes C5–C8 showed Ru(II) centered distinct d–d bands in
the visible region 680–700 nm (Fig. S3,d). The absorption peak
values have been tabulated in Table S3.
interactions (105–106 Mꢀ1) than complexes C1–C4 (104–105 Mꢀ1
)
due to (i) intercalation facilitated by the presence of phenanthro-
line ligands by insertion of the complex into the adjacent base pairs
of DNA and (ii) groove binding of the ferrocenyl moiety. Complex
C5 with a tyrosine substituted ferrocenyl moiety bound to the Ru
(II) center shows the highest binding constant value due to
additional hydrogen bonding interactions between —OH group of
tyrosine and DNA nucleobases which are accessible both in major
groove and minor groove and is also observed in the case of the free
ligand L1. Complex C8 with a tryptophan substituted ferrocenyl
moiety bound to Ru(II) center shows a similar binding constant
value as C5 which also may be due to additional hydrogen bonding
interaction between —NH group of tryptophan and DNA nucle-
obases. The titration curves for ligands L1–L4, complexes C2–C4
3.2. DNA binding studies
3.2.1. Electronic absorption titration
The presence of ground state interactions between the
biological macromolecule DNA and compounds under study have
been detected using absorption spectroscopy. DNA can provide
three distinctive binding sites (groove binding outside of DNA
helix, along major or minor groove, electrostatic binding to
phosphate group and intercalation), a behavior important for the
biological role of antibiotic and anticancer drugs in vivo [31]. The
binding efficiency of metal complex to DNA can be effectively
investigated employing electronic spectroscopy since the observed
changes in the spectra may give evidence of the existing
interaction mode [32]. Any interaction between the compounds
(LI–L4 and C1–C8) and DNA is expected to perturb the ligand
centered transitions of the compounds. Binding with DNA via non-
intercalative binding modes, such as electrostatic forces, van der
Waals interactions, dative bonds, hydrogen bonds and hydropho-
bic interactions generally results in increase in absorption intensity
(hyperchromism) upon increasing the concentration of CT-DNA
owing to the degradation of the DNA double helix structure. On the
and C6–C8, as well as Plot of [DNA]/(eA–ef) versus [DNA] for L1–L4
have been provided as Supplementary material in Fig. S4.
3.2.2. Competitive binding studies with ethidium bromide using
fluorescence spectroscopy
To further examine the mode of binding of the compounds with
DNA, via intercalation or groove binding, a competitive binding
study with two dyes: EB and DAPI have been carried out using
steady state fluorescence spectroscopy. Ethidium bromide (=3,8-
diamino-5-ethyl-6-phenyl-phenanthridinium bromide) is a phe-
Table 2
nanthridine fluorescence dye and is
a typical indicator of
KSV values of ruthenium complexes C5-C8.
intercalation, forming soluble complexes with nucleic acids and
emitting intense fluorescence in the presence of CT DNA due to the
intercalation of the planar phenanthridinium ring between
adjacent base pairs on the double helix [35,36]. Addition of a
second molecule, which may bind to DNA more strongly than EB
results in a decrease of the DNA-induced EB emission [37]. The
Compound
KSV (Mꢀ1
)
C5
C6
C7
C8
8.3 ꢄ103
7.3 ꢄ103
6.5 ꢄ103
1.2 ꢄ104

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P. Ramadevi et al. / Journal of Photochemistry and Photobiology A: Chemistry 305 (2015) 1–10
Fig. 4. Plot of DNA-DAPI competitive binding titration curve with C1.
emission spectra of DNA–EB (lex = 546 nm, lem = 610) in the
absence and presence of increasing amounts of ligands and
complexes have been recorded. Addition of the ligands L1–L4 or
the complexes C1–C4 did not have any kind of effect on the
emission intensity or nature of the emission of DNA–EB complex.
On the other hand addition of complexes C5–C8 resulted in a
significant decrease of the intensity of the emission band at 609 nm
indicating the competition of the compounds with EB in binding to
DNA (Fig. 3). The observed quenching of DNA–EB fluorescence
suggests that they displace EB from the DNA–EB complex and
interact with DNA by intercalation. The planar phenanthroline
ligands of complexes C5–C8 seems to have facilitated intercalation
resulting in partial replacement of EB from DNA–EB complex
resulting in the observed quenching of fluorescence.
The relative binding of complexes to CT-DNA was determined
by calculating the quenching constant (KSV) from the slopes of
straight lines obtained from the Stern–Volmer Eq. (2) [38]:
I0
I
¼ 1 þ Ksv½Qꢆ
(2)
where I0 and I are the emission intensities in the absence and the
presence of the quencher (C5–C8) respectively, [Q] is the
concentration of the quencher and KSV is the Stern–Volmer
constant which can be obtained from the slope of the plot of I0/I
versus [Q].
The Stern–Volmer quenching plots (Fig. 3, inset) illustrate that
the quenching of EB bound to DNA by C5–C8 is in good agreement
(R = 0.93–0.99) with the linear Stern–Volmer equation and the
Fig. 5. Schematic presentation of DAPI displacement from DNA helix by quencher molecule followed by fluorescence quenching and corresponding energy diagrams.

P. Ramadevi et al. / Journal of Photochemistry and Photobiology A: Chemistry 305 (2015) 1–10
7
Fig. 6. Effect of increasing amounts of the complexes C1–C8 and ethidium bromide
(EB) on the relative viscosity of CT-DNA (200
[Complex]/[DNA] = 0, 0.04, 0.08, 0.12, 0.16, 0.20 and 0.24.
m
M) in Tris–HCl buffer at 32 (ꢃ0.1) ꢂC.
Fig. 8. Stern–Volmer plots for the quenching of BSA fluorescence by C1–C8.
DNA–DAPI fluorescence. The initial fluorescence enhancement can
be attributed to partial overlap of the electronic states of the
quencher molecules (C1–C4) and DNA–DAPI complex leading to
partial stabilization of the ground state complex and an increase in
the value of the Franck–Condon factor. The fluorescence intensity
is proportional to the overlap of vibrational wavefunctions and
consequently to the Franck–Condon factor. A quencher molecule
when approaches a DNA–DAPI complex, it forms an intermediate
complex, which is temporarily more stable than the original DNA–
DAPI complex. This shifts the ground electronic state to left and
down, and as a result the overlap between the vibrational functions
in the excited and ground electronic states increases due to the
Frank–Condon principle (greater the overlap between the vibra-
tional functions of excited and ground states of the complex,
greater is the Franck–Condon factor), leading to the increase in
fluorescence signal. Further addition of the quencher leads to the
complete displacement of DAPI from the DNA helix and fluores-
cence depletion [41]. This phenomenon has been explained
schematically in Fig. 5. The quenching of the DNA–DAPI fluores-
cence is conclusive of the fact that complexes C1–C4 are replacing
DAPI from the minor grooves of DNA and themselves getting
bound. Moreover a small hump can be seen forming slowly on
increasing concentration of the compounds in titration curves 7–
11 (Fig. 4) marking the initiation of a new peak formation which is
indicative of a ground state complex formation of the compounds
with DNA. Thus it can be concluded that complexes C5–C8 with
planar phenanthroline rings, bind to CT-DNA via intercalation
Stern–Volmer quenching constant KSV values are given in Table 2.
Complexes C8 and C5 show higher quenching constant values of
1.2 ꢄ104 Mꢀ1 and 8.3 ꢄ103 Mꢀ1 respectively indicating their
greater efficiency to replace EB and bind strongly with DNA which
is also evident from their higher DNA binding constant (Kb) values.
The titration curves for C6–C8 have been provided as Supplemen-
tary material in Fig. S5.
3.2.3. Competitive binding studies with DAPI using fluorescence
spectroscopy
DAPI (40,6-diamidino-2-phenylindole) is
a classical minor
groove binder to DNA molecule. It is also shown to bind specifically
to GC regions by intercalation; however, the minor binding to AT
regions is 2 orders of magnitude stronger than the intercalative
binding mode. The fluorescence of DAPI increases approximately
30 times when 20-fold excess (base pairs) of DNA is added to the
solution of the dye [39,40]. The fluorescence spectra of a mixture of
DNA–DAPI solution with increasing concentration of C1–C4 have
been recorded (Fig. 4). The addition of aliquots of the complexes
cause an initial slight fluorescence enhancement which on further
addition of greater amounts showed subsequent quenching of the
Table 3
KSV,kq, Ka and n values for ligands and complexes.
Compound
KSV (Mꢀ1
)
kq (Mꢀ1 sꢀ1
)
Ka (Mꢀ1
)
n
L1
L2
L3
L4
C1
C2
C3
C4
C5
C6
C7
C8
4.5 ꢄ103
4.2 ꢄ103
2.5 ꢄ103
8.9 ꢄ103
3.2 ꢄ104
5.3 ꢄ104
4.2 ꢄ104
5.6 ꢄ104
4.1 ꢄ104
3.8 ꢄ 104
2.8 ꢄ 104
6.4 ꢄ104
4.5 ꢄ1011
4.2 ꢄ1011
2.5 ꢄ1011
8.9 ꢄ1011
3.2 ꢄ1012
5.3 ꢄ1012
4.2 ꢄ1012
5.6 ꢄ1012
4.1 ꢄ1012
3.8 ꢄ 1012
2.8 ꢄ 1012
6.4 ꢄ1012
1.8 ꢄ 103
1.0 ꢄ103
1.1 ꢄ103
3.0 ꢄ103
5.5 ꢄ104
8.1 ꢄ105
5.2 ꢄ104
2.8 ꢄ 105
2.8 ꢄ 106
7.3 ꢄ106
5.8 ꢄ 106
9.6 ꢄ106
0.98
0.8
0.9
0.87
1.05
1.2
1.02
1.14
1.41
1.28
1.51
1.48
Fig. 7. Plot of Fluorescence emission intensity versus wavelength for BSA at
different concentrations of C1.

8
P. Ramadevi et al. / Journal of Photochemistry and Photobiology A: Chemistry 305 (2015) 1–10
Table 4
IC50 values of complexes C1–C8 obtained from MTT assay on A549 cells.
Compound
IC50 values
g/ml)
(
m
(mM)
C1
C2
C3
C4
C5
C6
C7
C8
180
125
50
105
250
390
300
275
145
105
46
80
266
422
337
286
whereas complexes C1–C4 without the phenanthroline rings
prefer groove binding.
3.2.4. Viscosity measurements
Fig.10. % Cell viability in presence of complexes C1–C8 for A549 human lung cancer
cell lines. Each point is the mean ꢃ standard error obtained from three independent
experiments.
In order to further confirm the modes of binding of complexes
C1–C8 to CT-DNA, viscosity measurements of DNA solutions were
carried out in presence and absence of these complexes. The
viscosity of DNA is sensitive to length changes and is regarded as
the least ambiguous and the most critical clues of a DNA binding
mode in solution [42]. In general, intercalating agents are expected
to elongate the double helix to accommodate the ligands in
between the base pairs, leading to an increase in the viscosity of
DNA. In contrast, a complex that binds exclusively in the DNA
grooves typically causes less pronounced (positive or negative) or
no changes in DNA solution viscosity [31,43]. The effects of
complexes C1–C8 and classical intercalator EB on the viscosities of
CT-DNA solution are shown in Fig. 6. For complexes C5–C8 with
increasing [complex]/[DNA] concentration ratios, the relative
viscosities of CT-DNA increased gradually indicative of character-
istic intercalative mode of binding which is in accordance with
previous findings. In contrast with increasing [complex]/[DNA]
concentration ratios for complexes C1–C4 no significant change in
the relative viscosity of CT-DNA solution was observed which ruled
out intercalative binding mode of complexes and is consistent with
the DNA groove binding as indicated by DNA–DAPI competitive
binding studies.
3.3. BSA binding studies
3.3.1. Steady-state fluorescence spectroscopy
BSA is extensively studied, due to its structural homology with
human serum albumin (HSA). HSA contains one tryptophan
located at position 214, while BSA has two tryptophan residues
at positions 134 and 212 along the chain. BSA solutions exhibit a
strong fluorescence emission with a peak at 343 nm, due to the
tryptophan residues, when excited at 296 nm [44,45]. Neither the
ligands nor the complexes show any emission in the range 300–
450 nm on excitation at 296 nm. Addition of increasing concen-
trations of the ligands (L1–L4) and complexes (C1–C8) to a solution
of BSA results in a decrease of the fluorescence intensity as shown
for C5 in Fig. 7 because of their binding to BSA which may change
the protein conformation, subunit association or denaturation
leading to changes in the tryptophan environment of BSA.
The values of the Stern–Volmer quenching constant (KSV, Mꢀ1
)
and the quenching rate constant (kq, Mꢀ1 sꢀ1) for the ligands L1–L4
and the complexes C1–C8 interacting with BSA as calculated by
Stern–Volmer quenching equation (Eq. (2)) and the corresponding
Stern–Volmer plots (Fig. 8) are cited in Table 3 which suggest good
binding propensity of the complexes with BSA. The Kq values
(>1011 Mꢀ1 sꢀ1) are greater than diverse kinds of quenchers for
biopolymer fluorescence (1010 Mꢀ1 sꢀ1) indicating the existence of
static quenching mechanism [46]. The titration curves for ligands
L1–L4 and complexes C2–C8 are provided as Supplementary
material in Fig. S6 and the Stern–Volmer plots of ligands L1–L4 are
provided as Supplementary material in Fig. S7.
The association binding constant Ka (Mꢀ1) and the number of
binding sites per albumin (n) can be calculated by double
logarithm equation (Eq. (3)) [47]
logðI0 ꢀ IÞ
¼ logKa þ nlog½Qꢆ
(3)
I
The plot of log [(I0 ꢀ I)/I] versus log [Q] for all the ligands and
complexes are linear (Fig. 9) and the Ka and n values for ligands L1–
L4 and for complexes C1–C8 have been obtained from the intercept
and slope, respectively (Table 3). The double log plot for ligands
L1–L4 is provided as Supplementary material in Fig. S8.
In general, the values of quenching rate constants (Kq) and
association binding constants (Ka) are higher for complexes C1–C4
as compared to the ligands indicating complexation to ruthenium
increases their BSA binding affinities. Additionally higher values
for C5–C8 compared to C1–C4 reveal that the coordination of
Fig. 9. Double logarithmic plot for the quenching of BSA fluorescence by C1–C8.

P. Ramadevi et al. / Journal of Photochemistry and Photobiology A: Chemistry 305 (2015) 1–10
9
Fig.11. Confocal images of A549 cells treated with (100
mg/ml) C3 and (250 mg/ml) C5 for 16 h followed by dual staining with AO/EB. Here “L” stands for live cells, “NF” stands
for nuclear fragmentation and “AB” stands for apoptotic bodies. (For interpretation of the references to color in the text, the reader is referred to the webversion of this article.)
ferrocenyl amino acids to ruthenium in the presence of 1,10-
phenanthroline leads to enhanced affinity to BSA. The n values for
the ligands as well as the complexes average out to be 1 which
suggests that there is only one binding site available on the protein.
Moreover the linear nature of the double logarithm plots of L1–L4
and C1–C8 indicates that only one of the tryptophan residues on
BSA protein is interacting with the compounds [48].
Ethidium bromide (EB) is only taken up by cells when cytoplasmic
membrane integrity is lost, and stains the nucleus red. EB also
dominates over AO. Thus live cells have a normal green nucleus;
early apoptotic cells have bright green nucleus with condensed or
fragmented chromatin; late apoptotic cells display condensed and
fragmented orange chromatin; cells that have died from direct
necrosis have a structurally normal orange nucleus [52]. The cells
were treated with C3 at 100 mg/ml and with C5 at 250 mg/ml
3.4. Cytotoxicity
concentration for 16 h. The confocal images of control cells show
that the live cells are stained with AO and hence emit green
fluorescence whereas the treated cells bearing yellow nucleus
indicating early apoptosis (Fig. 11). Moreover distinct nuclear
fragmentation can also be seen within the treated cells and the
presence of apoptotic bodies makes it evident that the mode of cell
death is apoptosis.
In-vitro cytotoxicity tests were performed on the human lung
carcinoma (A549) cell line. The cell viabilities (%), obtained for
A549 cells with continuous exposure to the compounds for 24 h,
are depicted in Fig. 10. The cytotoxicities of the complexes were
found to be dose dependent, that is, the cell viability decreased
with increasing concentrations. The IC50 values of the complexes
have been tabulated in Table 4. The reported IC50 values of NAMI-A
which is a well known ruthenium complex currently under phase
4. Conclusion
2 clinical trials is in the range of 550
cancer cell lines [49]. Although the synthesized ruthenium
complexes are found to be less active compared to cis platin
(IC50 = 26 mM), they are much more active on A549 cancer cells as
compared to NAMI-A with complex C3 (IC50 = 46 mM) showing the
maximum potency.
m
M–750
m
M for various
The work discussed here thus focuses on the synthesis,
characterization and bioactivity of ferrocenyl amino acid mannich
base conjugates and their ruthenium complexes of: (1) M(L)3 type
where M = Ru(III), L = ferrocenyl amino acid mannich base con-
jugates and (2) M(L0)2L type where M = Ru(II), L0 = 1,10-phenanthro-
line. Literature has already shown that ferrocenyl compounds are
good DNA and protein binders which in the present study have
been found to be further enhanced on complexation with
ruthenium. This can be based on their better binding constants
obtained for interactions with DNA and BSA compared to those of
the free ligands. Moreover the ferrocenyl mannich base ligands and
their ML3 type Ru(III) complexes have been found to be good
groove binders unlike their mixed ligand Ru(II) complexes which
show intercalative mode of binding to DNA. Such impressive
binding efficacies of the ruthenium complexes containing ferro-
cenyl amino acids broaden their scope for in-vitro investigations as
potent anticancer agents following which we carried out the
cytotoxicity studies employing MTT assay. Lower IC50 values of the
Ru(III) complexes, C1–C4 can be either attributed to their reduction
to active Ru(II) species in the hypoxic medium within the tumor
cells or to the presence of three ferrocene moieties which have
been shown to be active against tumor cells. Furthermore the
mode of cell death, scrutinized using AO/EB dual staining
technique was found to be apoptosis also called as programmed
cell death which is the preferred mode of cell death over necrosis.
Experimental data on NAMI-A and Keppler like complexes have
led to the proposal that Ru(III) complexes are, in fact, prodrugs that
act by an “activation by reduction” mechanism, yielding more
active Ru(II) species. By analogy, lower IC50 values of C1–C4 as
compared to C5–C8 may be due to their reduction to active Ru(II)
species in the hypoxic medium within the tumor cells. Alterna-
tively the high anticancer activity of C1–C4 may be due to the
presence of more number of ferrocene moieties which are known
to have good anticancer activity [15,16,50]. The in-vitro anticancer
activity of the compounds was inconsistent with their DNA-
binding abilities. The complexes with only ferrocenyl ligands (C1–
C4) were found to be more cytotoxic than the complexes with both
ferrocenyl and phenanthroline ligands (C5–C8), whereas the DNA
binding order is the reverse i.e., C5–C8 > C1–C4. The different order
of DNA binding affinity and the in-vitro anticancer activity means
multiple targets and multiple mechanisms coexisted in the
anticancer process of the compounds. DNA binding and cleavage
need not be the only target and mechanism for cytotoxicity [51].
3.5. Induction of apoptosis
Acknowledgements
DNA-binding dyes AO and EB (Sigma, USA) were used for the
morphological detection of apoptotic and necrotic cells. Acridine
orange (AO) permeates all cells and makes the nuclei appear green.
The authors gratefully acknowledge the University Grants
Commission RFSMS-BSR fellowship [UGC No. F.4-1/2006 (BSR)/5-

10
P. Ramadevi et al. / Journal of Photochemistry and Photobiology A: Chemistry 305 (2015) 1–10
68/2007 (BSR)], New Delhi, for their financial assistance. The
authors are thankful to the Head, Department of Chemistry and
Department of Zoology, Faculty of Science, The Maharaja Sayajirao
University of Baroda for providing us with the necessary lab
facilities and required instrumentation facilities to carry out the
research work. Thanks are also due to Dr. M.N. Patel, Department of
Chemistry, Sardar Patel University, Vidyanagar for the DNA
viscosity measurements. The authors thank DBT-MSUB-ILSPARE
project, Dr. Vikram Sarabhai Science Block, M.S.University of
Baroda for providing the confocal LSM-710 fluorescence micro-
scope in order to procure the confocal images of AO/EB staining
technique.
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