Synthesis, characterization of ruthenium(II), nickel(II), palladium(II), and platinum(II) triphenylphosphine-based complexes bearing an ONS-donor chelating agent: Interaction with biomolecules, antioxidant, in vitro cytotoxic, apoptotic activity and cell cycle analysis

Article abstract of DOI:10.1016/j.jinorgbio.2021.111549

Four new transition metal complexes, [M(PPh₃)(L)].CH₃OH (M = Ni(II) (1), Pd(II) (2)) [Pt (PPh₃)₂(HL)]Cl (3) and [Ru(CO)(PPh₃)₂(L)] (4) (H₂L = 2,4-dihydroxybenzaldehyde-S-methyldithiocarbazate, PPh₃ = triphenylphosphine) have been synthesized and characterized by elemental analyses (C, H, N), FTIR, NMR (¹H, ³¹P), ESI-MS and UV–visible spectroscopy. The molecular structure of (1) and (2) complexes was confirmed by single-crystal X-ray crystallography. It showed a distorted square planar geometry for both complexes around the metal center, and the H₂L adopt a bi-negative tridentate chelating mode. The interaction with biomolecules viz., calf thymus DNA (ct DNA), yeast RNA (tRNA), and BSA (bovine serum albumin) was examined by both UV–visible and fluorescence spectroscopies. The antioxidant activity of all compounds is discussed on basis of DPPH^? (2,2-diphenyl-1-picrylhydrazyl) scavenging activity and showed better antioxidant activity for complexes compared to the ligand. The in vitro cytotoxicity of the compounds was tested on human (breast cancer (MCF7), colon cancer (HCT116), liver cancer (HepG2), and normal lung fibroblast (WI38)) cell lines, showing that complex (1) the most potent against MCF7 and complex (4) against HCT116 cell lines based on IC₅₀ and selective indices (SI) values. So, both complexes were chosen for further studies such as DNA fragmentation, cell apoptosis, and cell cycle analyses. Complex (1) induced MCF7 cell death by cellular apoptosis and arrest cells at S phase. Complex (4) induced HCT116 cell death predominantly by cellular necrosis and arrested cell division at G2/M phase due to DNA damage.

Full text of DOI:10.1016/j.jinorgbio.2021.111549

Journal of Inorganic Biochemistry 223 (2021) 111549
Contents lists available at ScienceDirect
Journal of Inorganic Biochemistry
journal homepage: www.elsevier.com/locate/jinorgbio
Synthesis, characterization of ruthenium(II), nickel(II), palladium(II), and
platinum(II) triphenylphosphine-based complexes bearing an ONS-donor
chelating agent: Interaction with biomolecules, antioxidant, in vitro
cytotoxic, apoptotic activity and cell cycle analysis
,
,*
Shadia A. Elsayeda *, Hagar E. Badra, Armando di Biaseb, Ahmed M. El-Hendawya
a Chemistry Department, Faculty of Science, Damietta University, New Damietta 34517, Egypt
b
`
Dipartimento di Chimica, Universita degli Studi di Milano, via C. Golgi 19, 20133 Milano, Italy
A R T I C L E I N F O
A B S T R A C T
Four new transition metal complexes, [M(PPh3)(L)].CH3OH (M = Ni(II) (1), Pd(II) (2)) [Pt (PPh3)2(HL)]Cl (3)
and [Ru(CO)(PPh3)2(L)] (4) (H2L = 2,4-dihydroxybenzaldehyde-S-methyldithiocarbazate, PPh3 = triphenyl-
phosphine) have been synthesized and characterized by elemental analyses (C, H, N), FTIR, NMR (1H, 31P), ESI-
MS and UV–visible spectroscopy. The molecular structure of (1) and (2) complexes was confirmed by single-
crystal X-ray crystallography. It showed a distorted square planar geometry for both complexes around the
metal center, and the H2L adopt a bi-negative tridentate chelating mode. The interaction with biomolecules viz.,
calf thymus DNA (ct DNA), yeast RNA (tRNA), and BSA (bovine serum albumin) was examined by both
UV–visible and fluorescence spectroscopies. The antioxidant activity of all compounds is discussed on basis of
DPPH• (2,2-diphenyl-1-picrylhydrazyl) scavenging activity and showed better antioxidant activity for complexes
compared to the ligand. The in vitro cytotoxicity of the compounds was tested on human (breast cancer (MCF7),
colon cancer (HCT116), liver cancer (HepG2), and normal lung fibroblast (WI38)) cell lines, showing that
complex (1) the most potent against MCF7 and complex (4) against HCT116 cell lines based on IC50 and selective
indices (SI) values. So, both complexes were chosen for further studies such as DNA fragmentation, cell
apoptosis, and cell cycle analyses. Complex (1) induced MCF7 cell death by cellular apoptosis and arrest cells at S
phase. Complex (4) induced HCT116 cell death predominantly by cellular necrosis and arrested cell division at
G2/M phase due to DNA damage.
Keywords:
Metal complexes
Cytotoxicity
ct DNA
tRNA
BSA
Apoptosis
1. Introduction
different types of cancer in clinics and is considered as a precursor for
other metallotherapeutic drugs such as oxaliplatin, carboplatin [1].
cis-Platin is the first platinum-based drug used for the treatment of
Since the discovery of cis-platin, its efficacy is evidenced in diverse types
Abbreviations: 1HNMR, Proton nuclear magnetic resonance; 31PNMR, Phosphorus-31 nuclear magnetic resonance; ARP, Antioxidant reducing power; BSA, Bovine
serum albumin; ct DNA, Deoxyribonucleic acid from calf thymus; DMF, Dimethyl formamide; DMSO, Dimethyl sulfoxide; DPA, Diphenylamine; DPPH, 2,2-diphenyl-
1-picryhydrazyl; EB, Ethidium bromide; EC50, Effective concentration; ESI-MS, Electron Spray Ionization mass spectroscopy; Et2O, diethyl ether; F, Fluorescence; fa,
Fractional accessible protein fluorophore; FTIR, Fourier-transform infrared spectroscopy; H2L, 2,4-dihydroxybenzaldehyde-S-methyldithiocarabzate; HCT116,
Human colon cancer; HepG2, Human liver cancer; HPLC, High Performance Liquid Chromatography; HSA, Human serum albumin; IC50, Half maximal inhibitory
concentration; Ka, Effective quenching constant for the accessible fluorophore; Kapp, Apparent binding constant; Kb, Intrinsic binding constant; Kq, Dynamic or
collision quenching constant; Ksv, Stern-Volmer constant; KP1019, [indazolium trans-[tetrachloride bis-(Hindazole) ruthenate(III)]; KP1339, (sodium trans-[tetra-
chlorobis(1H-indazole)ruthenate(III)]); LMCT, Ligand-to-metal charge transfer; MCF7, Human breast cancer; MLCT, Metal-to-ligand charge transfer; MTT, 3-(4,5-
dimethylthiazolꢀ 2-yl)-2,5-diphenyltetrazolium bromide; NA, Nucleic acid; NAMI-A, [imidazolium trans- [tetrachlorido(DMSO)(imidazole)ruthenate(III)]; PBS,
Phosphate buffered saline; PI, Propidium iodide; PDT, photodynamic therapy; PPh3, Triphenylphosphine; SAR, Structure activity relationship; SI, Selective index;
tRNA, Transfer ribonucleic acid; WI38, Human normal lung fibroblast;
* Corresponding authors.
ε, Extinction coefficient..
E-mail addresses: shadia.elsayed@du.edu.eg (S.A. Elsayed), amelhendawy@du.edu.eg (A.M. El-Hendawy).
https://doi.org/10.1016/j.jinorgbio.2021.111549
Received 7 April 2021; Received in revised form 12 July 2021; Accepted 15 July 2021
Available online 21 July 2021
0162-0134/© 2021 Elsevier Inc. All rights reserved.

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
of cancer including ovarian, colorectal, lung, and bladder cancers [2].
On the other hand, its toxicity, serious side effects, and drug resistance
property have limited its usage in the clinic [3,4]. From this point, re-
searchers dedicated their efforts to developing platinum and non‑pla-
tinum metal-based drugs using the advantage of cis-platin (clinical use)
with new strategies to improve the pharmacological properties and
elucidating the mechanism of action of the developed drugs [5]. In
addition to cis-platin and its derivatives, many metal complexes were
found to be potential anticancer agents, such as ruthenium [6], palla-
dium [7], and nickel [8]. The structure-activity relationship (SAR) is an
important factor to be taken into consideration while designing an
anticancer drug [9]. It explains the relationship between the molecular
structures and their biological activity [10], which enables determining
the structure or chemical group responsible for the biological effect
[11]. Then, the structure can be modified to enhance the bioactivity of
the compounds. Due to the resemblance between the coordination ge-
ometry of Pt(II), Pd(II) and Ni(II) complexes, palladium and, nickel
complexes have received considerable interest, due to the lesser side
effect and resistance therapies [7,12]. Platinum is most effective than
palladium due to ligand exchange kinetics, as palladium complexes
readily dissociate in solution (105 times faster than Pt), giving very
active species that are unable to reach the biological target such as DNA
and RNA and reduce the biological activity [13]. Hence, the structure
reactivity relationship plays an important role to design more stable Pd
(II) and Ni(II) complexes by introducing strong chelating agents, a bulky
ligand such as PPh3 [7,14], or strongly coordinating nitrogen ligand
[15] to overcome the rapid aquation process of theses complexes. From
square planar to octahedral complexes (Platinum to ruthenium), inor-
ganic and organometallic ruthenium (II/III) complexes are believed to
exhibit promising antitumor activity. Ruthenium(III) complex, sodium
trans-[tetrachlorobis(1H-indazole) ruthenium(III)]) (KP1339) is
currently developed for clinical investigation [16–18], and Ru(II)-
polypyridyl compound (TLD ꢀ 1433) has recently entered a human
clinical trial as PDT (photodynamic therapy) agent for bladder cancer
[19].
and protein may induce organ toxicity and other undesirable side ef-
fects, i.e. cisplatin can induce in vivo nephrotoxicity and genotoxicity
[35]. This may be attributed to excessive generation of free radicals by
the cisplatin inside the cell as well as intra-strand cross-link DNA by the
cisplatin in healthy cells [36]. That interaction may cause DNA damage
in vital organs, subsequently the appearance of secondary cancers [37].
So, it is important to innovate new metal-based anticancer drugs with
antioxidant properties to reduce organ toxicity through the free radical.
Also, the in vivo studies must be excessively done on the new metal-based
anticancer agents to avoid organ toxicity danger.
In this paper, due to the interesting properties of both dithiocarba-
zate and PPh3 ligand, we have synthesized and characterized four new
complexes of 2,4-dihydroxybenzaldehyde-S-methyldithiocarabzate
(H2L) with Ni(II), Pd(II), Pt(II), and Ru(II) bearing PPh3 ligand; [Ni
(PPh3)(L)].CH3OH (1), [Pd (PPh3)(L)].CH3OH (2), [Pt (PPh3)2(HL)]Cl
(3) and [Ru(CO) (PPh3)2 (L)] (4). We report their solution stability, calf
thymus (ct DNA)/yeast RNA (tRNA) and BSA binding properties, in vitro
anticancer activity against human normal and cancer cell lines.
Furthermore, DNA fragmentation, apoptotic activity, and cell cycle ar-
rest indicating that (1, 4) are promising metallotherapeutic candidates.
2. Experimental
2.1. Materials
The starting materials, NiCl2.6H2O, K2PdCl4, K2PtCl4 (Alfa Aesar),
RuCl3.xH2O (Pressure Chemicals), 2,4-dihydroxybenzaldehyde (Fischer
Scientific), Deoxyribonucleic acid sodium salt from calf thymus (ct
DNA), yeast tRNA (Sigma), bovine serum albumin BSA (Biomark,
98.5%) and HPLC grade solvents were used as received. The complexes
[Ni(PPh3)2Cl2] [38], [M(PPh3)2Cl2] (M = Pd(II), Pt(II)] [39], [RuHCl
(CO)(PPh3)2] [40] and S-methyldithiocarbazate [41] were prepared as
described in literature. All chemicals used for DNA fragmentation and
flow cytometry are from Sigma. The reagents, Roswell Park Memorial
Institute (RPMI)1640 medium, MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-
diphenyltetrazolium bromide) and dimethyl sulfoxide (DMSO) (Sigma
Co., St. Louis, USA), Fetal Bovine serum (GIBCO, UK). The cell lines,
Human (WI38), Colorectal carcinoma (HCT116), Mammary gland
breast cancer (MCF7) and Hepatocellular carcinoma (HePG2) were ob-
tained from ATCC via Holding company for biological products and
vaccines (VACSERA), Cairo, Egypt.
The unique properties of ruthenium complexes come from their
chemistry, it has variable oxidation states (+II to +IV), slow ligand-
exchange kinetics as well as octahedral geometry that allows the
structure diversity [20]. It has been also reported that ruthenium com-
plexes are less toxic, highly efficient, and selective against specific types
of tumors due to their strong affinity to bind to biomolecules (DNA/RNA
and bovine serum albumin (BSA) [18,21–23]. The mechanism of action
of ruthenium complexes to exert the antitumor effect mainly depends on
the nature of the ligand, complexes, and uncoordinated sites present in
the coordination sphere of the metal center [24]. Mechanism of action
may include inhibition of metastasis [25], binding to DNA and proteins
[26], apoptotic cell death, and production of reactive species [27].
Ligand design also has great importance in the development of
anticancer agents. S-alkyl and S-aryl dithiocarbazate Schiff bases are an
important class of biologically active chelating agents. They contain NS-
donors (hard/soft atoms) within a thioamide (thione-thiol) moiety
similar to those of thiosemicarbazones [28]. Their flexible properties
can be also modified incorporating different substituents [29]. This
characteristic improves their chelating ability by forming a stable
chelate with five or six-membered rings when linked to the metal ion
improving their structural stability as well as biological activity [30,31].
On the other hand, phosphine-based ligands are ligands of biological
interest including antitumor, antibacterial, and antifungal properties
[32]. They can form complexes with varieties of metal ions, such as Ni
(II), Pd(II), Pt(II), and Ru(II) with valuable bioactivities [7,26,33]. In
particular, Pd(II)/Ni(II)-PPh3-based complexes, to reduce their higher
lability to ligand exchange of Ni(II) and Pd(II) centers [7].
2.2. Methods
FT-IR spectra were recorded in the range of 4000–400 cmꢀ 1 on
JASCO 4100 FTIR spectrophotometer from KBr pellet. NMR (1H and 31P)
spectra were acquired on Bruker 400 using d6-DMSO as a solvent. ESI-
MS spectra were performed on Thermo Fisher LCQ Fleet ion trap mass
spectrometer equipped with HPLC UltiMate™ 3000 system. Elemental
analyses were done at the Microanalysis unit, Cairo University, Egypt.
The UV–Vis spectra of the compounds (in DMSO) were recorded by
JASCO V 630 using a quartz cuvette with 1 cm path-length at 298 K in
the range of 200–900 nm. Spectrofluorimeter (model 6285, UK) was
used for fluorescence excitation and emission spectra (200–700 nm).
Molar conductance values were determined using 10ꢀ 3 M solution of
complexes in DMF using CM-1 K portable conductivity meter. Micro-
plate reader (ELX800, USA) is used for cytotoxic study in MTT assay. The
single-crystal X-ray diffraction experiment was performed on a Bruker
Smart APEX II CCD diffractometer with graphite monochromated Mo-K
α
radiation (l = 0.71073 Å) [42] (Section 1.1, Electronic Supporting In-
formation (ESI†).
Nucleic acids (DNA/RNA) and proteins are attractive targets for
potential therapeutics. So the interaction of transition metals with those
biomolecules is essential to design any efficient metal-based anticancer
drugs [34]. However, the binding of transition metals with nucleic acids
2

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
2.3. Synthesis
concentrated to one third volume then left for slow evaporation and the
yellow precipitate was filtered off, washed with diethyl ether, then dried
in vacuo. Yield 75%. M. p.:170–172 ◦C. Anal. Calc. for
2.3.1. Preparation of 2,4-dihydroxybenzaldehyde-S-methyldithiocarbazate
ligand (H2L)
C45H39ClN2O2P2PtS2, Calcd.: C, 54.24; H, 3.95; N, 2.81%. Found. C,
An ethanolic solution of 2,4-dihydroxybenzaldehyde (1.38 g, 10
mmol, 10 mL) and S- methyldithiocarbazate [41] (1.22 g,10 mmol, 10
mL) were mixed and refluxed for 4 h in presence of drops of glacial
acetic acid. The resulting yellow solution was concentrated to one-half
volume and allowed to cool to room temperature. The yellow crystal-
line product was filtered, washed twice with ethanol, and dried under
vacuo. Yield: 71%. M. p.: 210–212 ◦C. Anal. Calc. for C9H10N2O2S2 (%):
C, 44.6; H, 4.1%; N, 11.6%. Found: C, 44.5; H, 4.0; N 11.4%. FT-IR
54.0; H, 3.63; N 2.51%. FT-IR (cmꢀ 1) in KBr:
ν(O H) 3283 b; ν(C=N)
–
–
–
–
ν
1603 s;
1252 s;
ν
(C=N)new 1556 s;
ν
(N N) 1060 s;
ν
(C S) 839 s;
(C O)phenolic
– –
(Pt P) 565 w; ν(Pt O) 504 w; (Pt N) 430 m; ν(PPh3) 998,
–
ν
ν
747 m. 1H NMR (400 mHz, DMSO‑d6, δ ppm, J Hz): δ 2.34 (s, 3H, SCH3);
δ 6.43(s,1H, H3); δ 6.22(d, J = 2.16 Hz, 1H, H5); δ 6.44(d, J = 4.4 Hz,
1H, H6); 7.18–7.70 (m, 30H, Ph); δ 8.53(s, 1H, —CH=N); δ 10.60(s, 1H,
4-OH); δ 10.72(s,1H,2-OH). 31P NMR (162 mHz, DMSO‑d6, δ ppm): δ
2.84 (d, 2JPP = 81.85 Hz), δ 23.76 (d, 2JPP = 70.60 Hz). ESI-MS (m/z):
(cmꢀ 1) in KBr:
ν
(O H) 3429 b, 3296 m;
ν
(N H) 3118 w;
ν
(C=N)
Calcd.: 960.15 ([Pt(HL)(PPh3)2]+), Found: 962.64 ([Pd(HL)(PPh3)2
+
–
–
1629 m;
ν
(C O) 1218 m,
2H+]). UV–Vis (DMSO, 1.66 × 10ꢀ 5 M): λmax (nm) ( , Mꢀ 1. cmꢀ 1): 370
ε
–
–
–
ν
(N N) 1097 m;
ν
(C=S) 1025 m;(C S) 844
m. 1H NMR (400 mHz, DMSO‑d6, δ ppm, J Hz): δ 2.52 (s, 3H, SCH3), δ
6.29 (s, 1H, H3), δ 6.33(d, J = 1.2 Hz, 1H, H5), δ 7.47(d, J = 8.0 Hz, 1H,
H6); δ 8.4(s, 1H, —CH=N); δ 10.07(s, 1H, 4-OH); δ10.25(s, 1H, 2-OH); δ
(9217), 417 (8675). Molar conductivity (10ꢀ 3 M, DMF, ΛM): 42
Ω
ꢀ 1cm2molꢀ 1
.
13.22 (s, 1H, NH). UV–Vis (DMSO, 1.66 × 10ꢀ 5 M): λmax (nm) (
ε
, Mꢀ 1
.
2.3.2.4. [Ru(CO)(PPh3)2 (L)] (4). The ligand (H2L) (0.242 g, 1 mmol)
was dissolved in benzene-ethanol mixture (5:1 v/v) and [RuHCl(CO)
(PPh3)3] [40] (0.952 g, 1 mmol) was added. The mixture was heated
refluxed for 5 h. The resulting orange solution was concentrated under
reduced pressure then petroleum ether was added to collect the pre-
cipitate. The yellowish orange product was filtered off, washed with
Et2O, and dried under vacuo. Yield 62%. M. p.: 160–162 ◦C. Anal. Calc.
for C46H38N2O2S2 RuP, Calcd.: C, 61.8; H, 4.28; N, 3.13%. Found. C,
–
cmꢀ 1): 312 (6807), 361 (35240) 377(25783).
2.3.2. Preparation of complexes
2.3.2.1. [Ni(PPh3)(L)].CH3OH (1). To a methanolic solution of H2L
(0.242 g, 1 mmol), [NiCl2(PPh3)2] [38] (0.653 g, 1 mmol) was added.
The mixture was refluxed for 3 h and the resulting orange solution was
concentrated to one third volume followed by addition of small amount
of diethylether (1 mL). The orange crystals suitable for X-ray crystal-
lography were collected by filtration, carefully washed by methanol and
diethylether then dried in vacuo. Yield: 78%. M. p.: 140–142 ◦C. Anal.
Calc. for C28H27N2NiO3PS2 (%): Calcd: C, 56.7; H, 4.6; N, 4.7%. Found:
–
61.60; H, 4.02; N, 3.20%. FT-IR (cmꢀ 1) in KBr:
ν
(O H) 3387 b;
ν
(C=N)
–
–
–
1625 s;
ν
(N N) 1031 s;
ν
(C S) 850 w;
ν
(C O)phenolic 1216 m;
– – –
(Ru P) 587w; ν(Ru O) 515 w; ν(Ru N) 462 w; ν(PPh3) 1031, 744 m;
ν
ν
(C=O) 1937 s. 1H NMR (400 mHz, DMSO‑d6, δ ppm, J Hz): δ 2.51 (s,
3H, SCH3); δ7.236–7.45 (m, 30H, Ph); δ 7.51 (s, 1H, H3); δ7.591(d, 1H,
H6); δ 8.24 (s, 1H, —CH=N); δ10.12 (s, 1H, 4-OH). 31P NMR (162 mHz,
DMSO‑d6, δ ppm): δ 29.31(s, 1P), 36.26 (s, 2P). ESI-MS (m/z): Calcd.:
894.08 [Ru(L)(CO)(PPh3)2], Found: [Ru(L)(CO)(PPh3)2+ H+]. UV–Vis
C, 56.6; H, 4.4; N 4.5%. FT-IR (cmꢀ 1) in KBr:
ν
(O H) 3386 b;
ν
(C=N)
– –
ν(C=N)new 1585 m; ν(N N) 1025 m; ν(C S) 845 m;
1618 s;
– – –
–
ν
ν
ν
(C O)phenolic 1228 m; ν(Ni P) 587w; ν(Ni O) 532 m; (Ni N) 437w;
(PPh3) 1069 m,745 m. 1H NMR (400 mHz, DMSO‑d6, δ ppm, J Hz): δ
(DMSO, 1.66 × 10 ꢀ 5 M): λmax (nm) (
ε
, Mꢀ 1. cmꢀ 1): 350 (11446), 395
2.50 (s, 3H, SCH3); δ 6.23(d, J = 2.2, 1H, H5); δ 5.78(d, 1H, H3); δ 7.24
(13313). Molar conductivity (10ꢀ 3 M, DMF, ΛM): 10 Ωꢀ 1cm2molꢀ 1
.
–
–
(d, J = 2.0, 1H, H6); δ7.244–7.7(m, 5H, Ph); δ 8.589(s, 1H,— CH N); δ
9.88(s, 1H, 4-OH). ESI-MS (m/z): Calcd.: 560.03 ([Ni(L)(PPh3)]), Found:
2.4. Solution stability
582.79 ([Ni(L)(PPh3) + Na]+). UV–Vis (DMSO, 1.66 × 10ꢀ 5 M): λmax
(nm) (
ε
, Mꢀ 1. cmꢀ 1): 306 (30662), 367 (28132), 384 (32349) and 408
Study the stability of the complexes in aqueous media is essential to
evaluate the biological activity of any metallodrug [43]. A stock solution
of 10ꢀ 3 M of each complex was prepared in DMSO (to improve solubi-
lity). The sample solutions were prepared by dilution of stock with
phosphate-buffered saline (PBS): pH 7.4, 137 mM NaCl, 2.7 mM KCl, 10
mM Na2HPO4 and 1.8 mM K2HPO4). The stability of complexes (1.66 ×
10ꢀ 5 M) was carried out in (1.6% DMSO/PBS v/v). The absorption
spectra of the complexes were measured with increasing time (over 24
h) at room temperature.
(24518). Molar conductivity (10ꢀ 3 M, DMF, ΛM):2.0 Ωꢀ 1cm2molꢀ 1
.
2.3.2.2. [Pd(PPh3)(L)].CH3OH (2). The ligand (H2L) ((0.12 g, 0.5
mmol) was dissolved in methanolic solution containing KOH (5 mL, 0.5
mmol). [PdCl2(PPh3)2] [39] (0.35 g, 0.5 mmol) was added to the above
solution. The reaction mixture was refluxed for 3 h during which the
orange precipitate obtained was filtered off, wash with methanol, fol-
lowed by diethylether, and then dried in vacuum. The filtrate was left at
room temperature for slow evaporation, orange crystals suitable for X-
ray crystallography were obtained, washed with methanol and dieth-
ylether. Yield: 81%. M. p.: 156–158 ◦C. Anal. Calc. for
2.5. Biological applications
C
28H27N2O3PPdS2, Calcd.: C, 52.46; H, 4.25; N, 4.37%. Found. C, 52.42;
H, 4.02; N 4.25%. FT-IR (cmꢀ 1) in KBr:
ν
2.5.1. Interaction with biomolecules
–
(O H) 3412,3283 b; ν(C=N)
The interaction studies of biomolecules (ct DNA, tRNA and BSA)
were performed in Tris-HCl buffer (pH = 7.2, 5 mM Tris-HCl + 50 mM
NaCl). The ctDNA and tRNA concentrations were determined spectro-
– – – –
ν(N N) 1072 s; ν(C S) 839 s; ν(C O)phenolic 1215 s; ν(Pd P)
1600 s;
588 w;
ν
(PPh3) 979 s,749 m. 1H NMR
– –
(Pd O) 530 w; ν(Pd N) 476 m; ν
(400 mHz, DMSO‑d6, δ ppm, J Hz): δ 2.56 (s, 3H, SCH3); δ 4.12 (s,1H, OH
(methanol)); δ 5.99 (s,1H, H3); δ 6.24 (d, J = 2.16 Hz, 1H, H5); δ 7.44 (d,
J = 4.4 Hz, 1H, H6); δ 7.55–7.63 (m, 15H, Ph); δ 8.57(s, 1H, —CH=N); δ
9.96(s, 1H, 4-OH). ESI-MS (m/z): Calcd.: 607.99 ([Pd(L)(PPh3)]),
Found: 609.01 ([Pd(L)(PPh3) + H+]). UV–Vis (DMSO, 1.66 × 10ꢀ 5 M):
photometrically at 260 nm (
ε
= 6600 Mꢀ 1 cmꢀ 1 for ct DNA) and (
ε =
7700 Mꢀ 1 cmꢀ 1 for tRNA) [44], whereas the concentration of BSA was
determined at 280 nm (
ε
= 66,433 Mꢀ 1 cmꢀ 1). The required concen-
trations of the test samples were prepared by dissolving the compound in
a minimum amount of DMSO, then completed to the required volume
with Tris buffer (1% DMSO/buffer). All the measurements were carried
out at room temperature.
λmax (nm) (
ε
, Mꢀ 1. cmꢀ 1): 308 (13012), 350 (9759), 396 (10542). Molar
conductivity (10ꢀ 3 M, DMF, ΛM): 2.0 Ωꢀ 1cm2 molꢀ 1
.
2.3.2.3. [Pt(PPh3)2(HL)]Cl (3). To an ethanolic solution of H2L (0.12 g,
0.5 mmol), [PtCl2(PPh3)2] [39] (0.395 g, 0.5 mmol) was added. The
mixture was refluxed for 3 h. and the resulting yellow solution was
2.5.1.1. Absorption studies. The binding affinity of biomolecules
(ctDNA, tRNA) towards metal complexes was investigated using
UV–visible spectroscopy. Various concentrations of ctDNA (0–140 μM)/
3

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
(a)
(b)
Fig. 1. Molecular structures for (a) [Ni(PPh3)(L)].CH3OH (1) and (b) [Pd(PPh3)(L)].CH3OH (2).
tRNA (0–45
μ
M) were added to a fixed concentration of the test com-
DMSO. The resulting solution was incubated in dark at room tempera-
pound 70 M for ct DNA and 45
μ
μ
M for tRNA. The absorption spectra
ture for 30 min before spectrophotometric measurement at 517 nm. The
were measured at 300–500 nm. Tris buffer was used as a blank, and an
equivalent amount of ct DNA/tRNA was added to the reference cell to
cancel the absorbance of ctDNA/tRNA.
DPPH• scavenging activity percent was determined using Eq. (1):
Ac ꢀ A
%DPPH scavenging activity =
s × 100
(1)
Ac
Where Ac and As are the absorbances in the presence and absence of
the test compound, respectively. The antioxidant activity is expressed
based on IC50 value and other antioxidant parameters: antioxidant
reducing power (ARP), effective concentration (EC50), stoichiometry,
and the number of reduced DPPH have been also determined [48].
2.5.1.2. Emission study. As the complexes has no fluorescent properties,
fluorescence emission spectra of the ligand and its complexes were
studied by pretreating the nucleic acid (ctDNA or tRNA) with ethidium
bromide (EB) ([ctDNA] = 50
μM, [tRNA] = 80
μM, [EB] = 5
μM in Tris-
HCl buffer. Different concentrations of test compounds (0–100
μM) were
added to the EB/DNA adduct. The emission was measured at 300–600
nm and the reduction in fluorescence intensity was recorded at 590 nm.
The quenching constant was evaluated by employing the Stern-Volmer
equation [45].
2.7. In vitro cytotoxic activity assay
The cytotoxic activity of the synthesized compounds was evaluated
to determine the inhibitory effects of compounds on cell growth using
MTT assay as depicted in (Section 1.2, ESI†).
2.5.2. BSA interaction studies
2.5.2.1. Absorption studies. The BSA-complexes interaction was also
2.8. Anticancer mechanism
determined spectrophotometrically by keeping the concentration of BSA
constant (15 μM) with adding different concentrations of test compound
2.8.1. Quantitative analysis of DNA fragmentation
(0–30 μM). The absorption spectra were recorded in the range 300–500
DNA fragmentation by the complexes (1) on MCF7 and (4) on
HCT116 cell lines was analyzed calorimetrically by diphenylamine
(DPA) reaction as explained in Section 1.3, ESI†).
nm after 5 min incubation period.
2.5.2.2. Emission study. The quenching emission of BSA tryptophan
residues was depicted using the ligand and complexes as quenchers. To a
2.8.2. Cell apoptosis by flow cytometry
fixed concentration of BSA (50 μM) in Tris-HCl buffer, varying concen-
The cell death mode of MCF7 cells by complex (1) and HCT116 cells
by complex (4) line has been performed by annexin V-FITC/PI (propi-
dium iodide) assay (Section 1.4, ESI†).
trations of quencher (0–50 μM) were added, and the emission intensity
was measured after a 5 min incubation period. The fluorescence emis-
sion spectra were scanned from 300 to 500 nm (excitation at 295 nm).
To determine the quenching parameters of the complexes for BSA, Stern
Volmer equations [46] were used.
2.8.3. Cell cycle analysis
The effect of complexes (1) and (4) on the DNA content by cell cycle
progression was assessed using MCF7 and HCT116, respectively (Section
1.5, ESI†).
2.6. Antioxidant activity by DPPH• (2,2-diphenyl-1-picrylhydrazyl)
radical scavenging assay
3. Results and discussion
The free radical scavenging activity was determined as reported in
the literature [47]. Various concentrations of the test compounds
3.1. Synthesis
(25–600 μM) and reference compound (ascorbic acid as a standard)
were prepared in 2.0 mL of DMSO. Then mixed with 1 mL of DPPH in
Synthesis and characterization details of the ligand and its complexes
4

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
O1–Ni–S1 176.74 (6) and P–Ni–N1 176.44 (6)◦ near the ideal angle
180◦. Similar related bond angles (Table 1) are found for palladium(II)
complex (2). Bond angles with a square planar coordination environ-
ment of Ni(II) and Pd(II) are in agreement with that found in the liter-
ature [50,51]. The donor atoms O1, N1, and S1 form the tridentate
ligand (H2L), and the phosphorous atom P of PPh3, occupies the square
plane. For complexes (1) and (2), the data depicted in Table 1 of bond
lengths M – X (M = Ni(II), Pd(II); X = O, N, S, P) are similar to those
reported for related nickel(II) and palladium(II) complexes [50,52] The
Table 1
Selected bond lengths (Å) and bond angles (◦) for nickel(II) (1) and palladium(II)
(2) complexes.
(1)
(2)
Bond lengths (Å)
M–S1
2.1475(6)
1.8536(16)
1.8808(18)
2.2173(6)
1.275(3)
2.2455(6)
2.0192(5)
2.0112(16)
2.2759(6)
1.275(2)
M–O1
M–N1
M–P
C2–N2
C2–S1
–
C2–N2 bond (1.275(3)Å)(1), (1.275(2)Å)(2) as the usual C N bond
–
1.743(2)
1.752(2)
length [53,54]. The bond length C2–S1 1.743(2) Å for (1) and 1.752(2)
Å for (2) is the same for the single bond C2–S2 1.752(2)(1) 1.7514(9)Å
(2). This is close to that found for similar related O,N,S- donor ligands
chelated in their thiolate form [50,54], where the thiol form is also
characterized by the fact that the hydrazinic nitrogen N1 is not bonded
to any hydrogen atom. The selected bond parameters are listed in
Table 1.
C2–S2
1.752(2)
1.7514(9)
Bond angles (◦)
O1–M–P
88.97(5)
89.89(2)
86.64(5)
94.54(7)
176.74(6)
176.44(6)
91.28(4)
92.314(19)
84.12(5)
92.38(1)
175.45(5)
175.91(5)
P–M–S1
S1–M–N1
N1–M–O1
O1–M–S1
P–M–N1
The data of specific contacts O∙∙∙H, S┄H and C┄H of the intermo-
lecular hydrogen bonding interactions characterized in the lattice for
complexes (1) and (2) are presented in Table 2. Each molecule of the
complex is connected with a strong hydrogen bond between O3 of
CH3OH solvent and H2 of the free hydroxyl group (dH(H2⋯O3) = 2.005
Å (1), 1.971 Å (2)). The distances dH(O┄H) are shorter than the
maximum values of 2.72 Å for the van der Waals radii of hydrogen and
oxygen atoms and considered for any contact [55]. The molecules are
interconnected via S2 atom from one molecule and H24(1) or H18(2) of
the other neighboring one, with a weaker interaction C18–H24(H18)⋯
∙S2 dH(H24(H18)⋯∙S2) = 2.924 Å (1), 2.97 Å (2). This is similar to that
observed in related O,N,S- metal complexes [56–58]. The molecules
form parallel planes which are constructed through C10 (PPh3) in a
plane and H3(L2ꢀ ) of the other plane the contact C3–H3⋯C10
(dH(H3⋯C10) = 2.802 Å (1), 2.857 Å (2)). In each complex ((1) or (2)),
the molecules are also specifically contacted through C28(CH3OH) and
H2(2-OH), dH(H2⋯C28) = 2.854 Å (1) or through C13(PPh3) and H3A
(CH3OH), dH(H3A⋯C13) = 2.748 Å (2). These lattice contacts are
shown in (Fig. S1a,b, ESI†).
Table 2
Hydrogen bonding parameters for complexes (1) and (2).
–
–
(Å)
–
D
H⋅⋅⋅A
Symmetry
D
H
H⋅⋅⋅A
D⋅⋅⋅A
D
H⋅⋅⋅A
(Å)
(Å)
(◦)
(1)
(i) O┄H and S┄H
contacts
O2–H2⋯O3
x,3/2–y,–1/2
+ z
0.775
0.93
2.005
2.924
2.762
3.684
165.66
139.91
C18–H24⋯S2
x,3/2–y,1/2
+ z
(ii) C┄H contacts
O2–H2∙∙∙C28
C3–H3∙∙∙C10
x,y,z
0.775
0.93
2.854
2.802
3.568
3.668
154.39
155.04
2–x,2–y,1–z
(2)
(i) O∙∙∙H and S∙∙∙H
contacts
O2–H2∙∙∙O3
x,3/2–y,–1/2
+ z
0.81
0.93
0.82
0.93
1.971
2.97
2.771
3.707
3.509
3.726
168.97
137.16
155.1
C18–H18∙∙∙S2
(ii) C∙∙∙H contacts
O3–H3A∙∙∙C13
x,y,z
3.2.2. Vibrational spectra
The infrared spectral data of the ligand and its complexes have been
identified and presented in the experimental Section 2.3. The FTIR
spectrum of the free ligand (H2L) (Fig. 2a), showed its distinctive bands
–1 + x,3/
2.748
2.857
2–y,–1/2 + z
1–x,1–y,1–z
C3–H3∙∙∙C10
156.2
at 3429 and 3118 cmꢀ 1 which are assigned to
ν
(N H)
ν(C=S)
– –
(O H) and ν
stretching vibrations, respectively. In addition, the presence of
have been mentioned in the experimental Section 2.3 (Synthesis). All
synthesized compounds are stable in air, insoluble in water, but fairly
soluble in DMSO and DMF, and partially soluble in CH2Cl2 and CH3CN.
The complexes (1), (2), and (4) showed lower molar conductance values
(2.0–10.0 Ωꢀ 1cm2molꢀ 1) which indicated the non-electrolytic nature of
the complexes, whereas complex (3) has a value of 42 Ωꢀ 1cm2molꢀ 1
indicating 1:1 electrolytic behavior [49].
stretching vibration at 1097 cmꢀ 1, suggests the presence of thione form
(HN–C = S) of the free ligand (Schem1a). This phenomenon is also
ꢀ 1
–
supported by the absence of
ν
(S H) at ca. 2500 cm [48,59]. In the
– –
FTIR spectra of complexes (1), (2) and (4), the ν(O H) and ν(N H)
–
bands disappeared, and the ν(C O) band shifted to higher frequency
[60,61] (1227–1252 cmꢀ 1) as compared to free ligand (1218 cmꢀ 1).
This result ascertained the involvement of phenolic oxygen (2-OH) in
coordination after deprotonation [60]. The coordination via azomethine
nitrogen ν(C=N) was indicated by the shift of this band to lower fre-
3.2. Characterization
quency in the spectra of the complexes in the (1600–1625 cmꢀ 1) region.
–
ν(N N) to lower frequency
This is also supported by the shift of
3.2.1. Molecular structure
(1027–1060 cmꢀ 1) compared to those of the parent ligand (1097 cmꢀ 1).
Suitable crystals for X-ray diffraction analysis for nickel(II) and
palladium(II) complexes [M(L)(PPh3)3].(CH3OH) (M = Ni (1), M = Pd
(2)) were obtained by slow evaporation of their methanolic solutions.
Crystallographic data of complexes (1) and (2) are listed in (Table S1,
ESI†) and their molecular structures are shown (Fig. 1a and b). The
ligand 2,4-dihyroxybenzaldehyde S-methyl dithiocarbazate (L2ꢀ ) co-
ordinates to Ni(II) or Pd(II) centers in a bi-negative tridentate manner
through phenolic oxygen O1, azomethine N1, and thiolate sulfur S2,
forming stable five- or six-membered chelate rings. The nickel or
palladium center adopts a slightly distorted square planar geometry
with bond angles; O1–Ni–P 88.97 (5)◦, P–Ni–S1 89.89 (2)◦, S1–Ni–N1
86.64 (5)◦ and N1–Ni–O1 94.54 (7)◦ of the sum 360◦ and the angles;
The absence of
ν
(C=S) band at 1025 cmꢀ 1 in the ligand is replaced by a
lower frequency band (839–850 cmꢀ 1) for
ν(C – S) [62] in complexes
along with the appearance of a new strong azomethine band in
‾
1556–1580 cmꢀ 1 region due to the formation of (C N – N = C – S )
–
–
moiety (Scheme 1c), this confirms that the ligand coordinates to the
metal center through the deprotonated thiol sulfur (Scheme 1b). This
concludes that the ligand acts as a bi-negative tridentate ONS-donor
through deprotonated phenolic oxygen, azomethine nitrogen, and thi-
olate sulfur as shown in (Fig. 2b, as a representative example).
Another coordination mode was exhibited by platinum(II) complex
– –
(3), where there is no significant shift in ν(O H) bands, while ν(N H)
5

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
H2L
100
90
80
70
60
50
NH
OH
N-N
C=S
C=N
4000
3500
3000
2500
Wavenumber
2000
-1
(cm
1500
)
1000
500
M-N
(b)
(4)
100
80
60
40
20
OH
M-O
C-S
N-N
PPh
3
C=N
CO
4000
3500
3000
2500
Wavenumber (cm )
2000
1500
1000
500
-1
Fig. 2. FT-IR spectra of (a) ligand (H2L) and (b) [Ru(CO)(PPh3)2 (L)] complex (4).
3
O-
HO
HO
4
2
OH
N
OH
HO
OH
N
PPh3
PPh3
S-
SH
CH3
S
S
OC
+X
8
PPh3
1
5
N
N
CH3
CH3
S
+ 2H+
N
HO
N
S
Ru
N
O
HO
6
7
H
S
Cl
PPh3
Pt
S
(c)
(a)
N
(b)
N
CH3
N
S
X = [MCl2(PPh3)2] (M = Ni(II), Pd(II)) and [RuHCl(CO)(PPh3)3]
H3C
S
(a)
(b)
Scheme 1. Tautomeric forms of H2L ligand; (a) thione and (b) thiol; (c)
ionic form.
Fig. 3. Proposed structure of (a) [Pt(PPh3)2(HL)]Cl (3) and (b) [Ru(CO)
(PPh3)2(L)] (4).
band disappeared, and a shift to a lower frequency for ν(C=N) was found
–
together with new bands display for ν(C=N) and ν(C S) as in complexes
data are constituent with a bi-negative tridentate behavior as ONS-
chelating donor for all complexes except for that for platinum(II) (3),
where NH signal disappeared and the 2-OH signal remain unaltered.
This indicates that the ligand coordinates to the Pt(II) center in a mono-
negative bidentate manner through thiol sulfur and azomethine nitro-
gen as NS-donor (Fig. 3a). The triphenylphosphine protons are located
in their expected position at δ7.18–7.70 ppm [63]. The 31P NMR spec-
trum of Pt(II) complex (3), shows two doublets at δ 2.70 and 23.48,
which indicates the presence of two magnetically different P-atoms co-
ordinated to platinum(II), corresponding to four satellite signals of the
Pt(II) ion [69]. For Ru(II) complex (4) (Fig. 3b) two signals at 29.0 and
36.33 ppm are shown, indicating the presence of two coordinated tri-
phenylphosphine groups in cis-configuration [70].
(1), (2) and (4). This confirms the mono-negative bi-dentate (HL)
manner of the ligand as NS-donor through azomethine nitrogen and
thiol sulfur (Scheme 1b). In all complexes, the new bands observed at
1091–1097 and 692–694 cmꢀ 1 are designated to
ν(P-Ph) stretching vi-
brations [63], while the bands appeared at 565–586 cmꢀ 1 due to
504–532 and 430–476 nm are due to ν(M – P), ν(M – N) and ν(M – N),
respectively [64,65]. In Ru(II) complex (4), the strong band observed at
1937 cmꢀ 1 is characteristic for
ν(CO). The presence of CH3OH molecule
in the structure of the complex (2) is defined by the band at 3597 cmꢀ 1
and that of complex (1) is obscured by the broad band at ca. 3500 cmꢀ 1
.
The FTIR spectra of the complexes (1)–(3) are presented in (Fig. S2a–c,
ESI†).
3.2.3. NMR spectra
3.2.4. Mass spectra
All 1H NMR spectral data and their assignments of H2L ligand and its
complexes are stated in the experimental section (2.1) and (Fig. S3a–e,
ESI†). The 1H NMR spectrum of the ligand shows singlets at δ13.22,
10.25, 10.07, 8.40, 6.29, and 2.52 due to NH, 2-OH, 4-OH, CH = N, 3-H,
Electron Spray Ionization mass (ESI-MS) is a valuable technique for
molecular weight determination in coordination compounds. The ESI-
MS spectra of the complexes (1)–(4) are shown in (Fig. S4a–d, ESI†).
Their molecular ion peaks [M+] are in good agreement with the pro-
posed chemical structure. The positive ion ESI-mass spectrum of [Ni(L)
(PPh3)].CH3OH complex (1) shows fragmentation patterns attributed to
subsequent degradation of the complex. The signal at m/z = 582.79
(Calcd. 583.02, 100%) and 298.92 (Calcd. 297.94, 12%) ascribed to [Ni
(L)(PPh3) + Na]+ and [Ni(L) + H]+, respectively. In addition, the signal
located at m/z = 1144.23 (Calcd. 1120.06) with relative abundance
100%, attributes to [Ni(L)(PPh3)]2 + Na]+, and assigned to association
of two molecules. The positive ion ESI-mass spectrum of [Pd(PPh3)
(L)].CH3OH complex (2) shows molecular ion peak at m/z = 609.01
(Calcd. 607.99) with a relative abundance of 100% corresponds to [Pd
and S-CH3 protons, respectively [48,60]. The two doublets occurred at δ
–
6.33 (J 1.2 Hz) and δ 7.47 (J = 8.0 Hz) are assigned to 5-H and 6-H
–
respectively. The absence of SH signal at ~4.0 ppm, confirms the thi-
one moiety of the ligand (Scheme 1a). In 1H NMR spectra of the com-
plexes (1), (2) and (4), the NH and 2-OH signals disappeared upon
coordination indicating the deprotonation before the coordination.
Furthermore, the coordination of azomethine nitrogen is confirmed by
the downfield or upfield shift (8.24–8.61 ppm) of azomethine proton
(CH=N) [66]. The splitting of the CH resonance of azomethine protons
may be due to the nuclear quadrupolar effect [67,68]. Thus, the 1H NMR
6

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
during the first hour near 360 and 400 nm attributed to fluctuation or
modification of the original compounds, then the data obtained up to 24
h shows a decrease in absorbance with no significant changes in band
positions indicating substantial stability of the complexes in buffer-
DMSO solution [74,79]. Complex (3) shows a slight spectral change at
λ = 430 nm which may simply result from the solvent exchange (DMSO
or H2O) with chloride ion.
1.0
(H L)
2
(1)
(2)
(3)
(4)
0.8
0.6
0.4
0.2
0.0
4. Biological applications
4.1. Interaction with biomolecules (ct DNA and tRNA and BSA)
In classical studies, DNA was known as the major target to design any
anticancer drugs.Recently, RNA also not less important than DNA in that
purpose, as the RNA itself is synthesized from DNA during the tran-
scription process [80] which in turn produces proteins in the body by
structures call ribosomes [81]. This concludes that both DNA and RNA
possess an important role in the drug design field and they are also the
key factor for many deadly diseases. Thus, in this context, we have
studied the binding properties of nucleic acid (ct DNA, tRNA) and BSA
with our complexes (1)–(4) compared to the parent ligand (H2L) using
absorption and emission studies.
300
400
500
600
Wavelenght (nm)
Fig. 4. Electronic spectra of ligand (H2L) and complexes (1)–(4) in 1.6 × 10ꢀ 5
M DMSO.
(L)(PPh3) + H]+, while in negative ion mode the base peak signal
showed up at 607.32 with relative abundance 100% corresponds to [Pd
(PPh3)(L)-H]ꢀ and the second fragment located at 345.16 (345.91, 20%)
due to [Pd(L)-H]ꢀ fragment. The positive ion ESI-mass spectrum of [Pt
(HL)(PPh3)2]Cl complex (3), shows its molecular ion peak at m/z =
962.64 (Calcd. 960.15 with relative abundance 10%) is assigned to [Pt
(PPh3)2(HL)]+. The other fragment observed at m/z = 698.01 (100%,
Calcd. 698.06) is attributed to [Pt(HL)(PPh3)]+ arising from the loss of
one PPh3 ligand. The positive ion ESI-mass spectrum of [Ru(CO)
(PPh3)2(L)] complex (4) show the first signal at m/z = 895.15 with
relative abundance 100% (Calcd. 894.08) assigned to [Ru(CO)
(PPh3)2(L) + H]+. The second signal appeared at m/z = 633.0 (8%,
Calcd. 631.99) is attributed to [Ru(CO)(PPh3)(L) + H]+ which corre-
spond to the cation formed by missing of PPh3 ligand.
4.1.1. Absorption studies
4.1.1.1. Ct DNA/tRNA binding studies. Electronic absorption titration of
the ligand and its complexes (1)–(4) has been carried out using UV–vi-
sible absorption spectra of the tested compounds in the absence and
presence of nucleic acid in the range of 200–600 nm. With the incre-
mental addition of ct DNA/tRNA to a fixed concentration of the com-
pounds, the absorption band observed at 350–420 nm decreased with
6–20% hypochromism. This confirms that the compounds undergo
interaction with the nucleic acids via intercalation mode [82]. This
mode of interaction is typically due to the π-π stacking interaction of
aromatic chromophore of the ligand with the base pairs of the nucleic
acid, which result in conformational alternation on the molecules of
DNA [83]. The binding strength of the compounds with nucleic acid has
been determined based on the intrinsic binding constant (Kb) using the
Wolfe–Shimer Eq. (2) [84,85]:
3.2.5. Electronic spectra and magnetic susceptibility measurements
The magnetic susceptibility measurements revealed that all com-
plexes are diamagnetic with square planar geometry for Ni(II), Pd(II),
and Pt(II) complexes which assigned to their (d8) configuration [71] and
low spin (d6, S = 0) for Ru(II) complex [48,54]. The UV–visible spectra
[Nucleic acid]
[Nucleic acid]
1
of the ligand and its complexes (1)–(4) were measured for 1.6 × 10ꢀ 5
M
ꢀ
)
ꢀ
)
ꢀ
)
=
+
…
(2)
εa
ꢀ
εf
εb
ꢀ
εf
Kb εb
ꢀ
εf
in DMSO solution at room temperature in the 200–600 nm range. The
electronic spectrum of the ligand shows absorption bands in 290–377
Where [Nucleic acid] is the molar concentration of ct DNA or tRNA,
εa is the apparent extinction coefficient of the complex bound to nucleic
acid (Aobs/[compound]), εf and εb are the extinction coefficients of the
free complex and that fully bound to DNA, respectively. The electronic
spectral changes upon addition of the nucleic acid to the complexes (1)–
(4) are presented in Fig. 5 and Fig. S6a, ESI† for the ligand. The equi-
librium binding constant (Kb) was obtained from the plot of [DNA] or
nm which are assigned to π-π* and n-π* transitions that resulted from
azomethine and thiocarbonyl moieties [72,73]. In the electronic spectra
of the complexes, these bands were shifted into higher wavelength
(redshift) 307–450 nm along with the appearance of a new band in
385–420 nm due to metal-to-ligand charge transfer (MLCT) due to the
coordination of metal (II) ion with the azomethine nitrogen and thiol
sulfur [74,75]. The d-d transition bands of the complexes obscured due
to their relatively low extinction coefficient compared to ligand-to-metal
charge transfer (LMCT) or MLCT [76]. The electronic spectra measure-
ments are shown in (Fig. 4 and Table S2, ESI†).
[RNA]/(εa - εf) vs [DNA] or [RNA] (Fig. 5) and dividing slope over the
intercept. The intrinsic binding constant (Kb) of the compounds was
found to be in the order 104 Mꢀ 1 as depicted in Table 3. The Kb values for
the complexes are of the order (4) > (3) > (1) > (2) > (H2L). This in-
dicates that the binding affinity of complexes is increased than that of
the ligand to ~2.5–8.0-fold for ct DNA and 12–24-fold for tRNA. The
binding affinities of tRNA to the metal complexes are greater than those
of the DNA (4.5–8.9 × 104 Mꢀ 1), which was expected due to striking
differences in the 3-dimensional morphology of ct DNA and tRNA.
Moreover, in contrast to DNA, the minor groove of RNA helix is wider
(shallow minor groove), which is more accessible for specific binders, to
form hydrogen bonds to the hydroxyl and phosphate groups with the
aromatic chromophore ligand [86]. The binding constant (Kb) values of
the compounds are 4.60 (4) > 3.40 (3) > 2.02 (1) > 1.38 (2) > 0.58
(H2L) × 104 Mꢀ 1 for ct DNA and 8.90 (4) > 6.90 (3) > 5.40 (1) > 4.50
(2) > 0.37 (H2L) × 104 Mꢀ 1 for tRNA (Table 3), which are similar to
3.2.6. Solution stability
Study the stability of metal complexes in solution is an essential step
while evaluating their interaction with biomolecules and anticancer
activity [43]. As the anticancer agents are usually introduced into the
bloodstream, it is worth studying their stability in aqueous media. For
example, it is reported that hydrolysis has a positive influence on the
anticancer activity of ruthenium(II) and platinum(II) complexes by
activating the DNA binding [77,78]. Hence, the solution stability of the
complexes (1)–(4) was studied by UV–Vis spectroscopy in phosphate-
buffered saline (pH = 7.4) containing 1% DMSO over a time period of
24 h (Fig. S5, ESI†). The complexes displayed an increase of absorbance
7

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
2.0
1.8
1.6
1.4
1.2
1.0
(a) ct DNA
7
6
5
4
3
2
(b) tRNA
(1)
1.0
0.8
0.6
0.4
0.2
1.2
1.0
0.8
0.6
0.4
0.2
(1)
0
3
6
9
12
15
0.5
1.0
1.5
2.0
2.5
3.0
3.5
[DNA]
[RNA] 10-5
M
350
(2)
400
450
500
550
600
350
400
450
500
550
600
Wave lenght (nm)
Wavelength (nm)
0.6
0.5
0.4
0.3
0.2
0.1
4.0
3.6
3.2
2.8
2.4
1.4
1.2
1.0
0.8
0.6
0.4
0.2
5.0
4.5
4.0
3.5
3.0
2.5
2.0
(2)
0
1
2
3
4
5
[RNA] 10-5
M
0
1
2
3
4
5
6
7
8
-5
[DNA] 10
M
300 350 400 450 500 550 600
Wavelenght (nm)
300 350 400 450 500 550 600
Wavelengh
0.8
80
60
40
20
0.8
8
7
6
5
4
3
2
(3)
(3)
0.6
0.4
0.2
0.0
0.6
0.4
0.2
0
10
20
30
40
50
-5
[DNA] 10
M
0
1
2
3
4
5
[RNA] 10-5
M
350
400
450
500
550
600
350
400
450
500
550
600
Wavelength (nm)
Wavelength(nm)
2.2
2.0
1.8
1.6
1.4
1.2
1.0
1.4
1.2
1.0
0.8
0.6
0.4
0.2
0.8
0.7
0.6
0.5
0.4
0.3
0.2
0.1
(4)
2.4
2.0
1.6
1.2
0.8
(4)
0
5
10
20
M
25
15-6
0
1
2
3
4
[DNA] 10
-5
[RNA] 10
M
350
400
450
500
550
600
350
400
450
500
550
600
Wavelenght (nm)
Wavelenght (nm)
Fig. 5. Electronic spectral changes of the complexes (1)–(4) in Tris-HCl buffer (pH =7.2) upon addition of different concentrations of (a) ctDNA, (b) tRNA. Inset: plot
of [DNA] or [RNA]/(εa εf) vs [DNA] or [RNA].
-
those obtained for related nickel(II) and palladium(II)-PPh3 base com-
plexes of hydrazone complexes [8,74].
biology, and pharmacology due to their significant importance in
pharmacology as it consists of 76% similarity with HSA [87]. The
electronic absorption study is a very useful tool to evaluate the
quenching type (static or dynamic) as well as the structural changes of
BSA by the effect of interaction with small molecule drugs [88]. This
study has been performed for BSA without and with different
4.1.1.2. Bovine Serum Albumin (BSA) binding studies. Study the inter-
action of small molecules with bovine serum albumin (BSA) have
recently received much attention from researchers in chemistry,
8

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
Benesi–Hildebrand Eq. (3) [88]:
(A∞ ꢀ Ao)
Table 3
Ct DNA and tRNA binding constants (Kb, Ksv, Kq and Kapp) obtained from ab-
sorption and fluorescence spectroscopy for the ligand and its complexes.
1
=
+ 1…
(3)
(Ax ꢀ Ao) Kb[com pound]
Compound
Absorption
Emission spectroscopy
spectroscopy
Where Ao is the absorbance of free BSA, Ax, the absorbance of BSA
upon addition of different concentrations of the compounds, and A∞ is
the fully bound BSA with the compounds. The binding constants for the
ligand and its complexes can be obtained from the linear relation be-
tween (A∞–Ao)/(Ax–Ao) against (1/[compound]) (Fig. 6, Fig. S6b, ESI†)
and intercept to slope ratio. The obtained binding constants of the
compounds are (1.70–4.29) × 104 Mꢀ 1 and in the order of (4) > (3) >
(1) > (2) > (H2L). The same as ctDNA and tRNA, the compounds follow
the same order and the complex (4) had the highest binding affinity with
the protein in a spontaneous fashion.
(Kb) × 104
Ksv (Mꢀ 1) × 103
Kq (Mꢀ 1Sꢀ 1) ×
Kapp
105
×
1011
ct
tRNA
ct
tRNA
ct
tRNA
ct DNA
DNA
DNA
DNA
H2L
(1)
(2)
(3)
(4)
0.58
2.02
1.38
3.4
0.37
5.4
4.5
6.9
8.9
2.95
3.76
ꢀ 1.97
8.4
2.65
2.82
–
6.69
9.46
1.34
1.7
1.2
1.28
–
3.04
4.3
2.7
3.7
–
7.9
8
ꢀ 0.89
3.8
4.6
12
5.45
It is worth noting that the process of dynamic quenching occurs only
in the excited state, where the lifetime is extremely short, the quencher
(compounds) and fluorophore (BSA) come into contact during the
temporary existence of the excited state, and it has no role in the ab-
sorption spectrum [92].
concentrations of the test compounds (ligand and its complexes) in Tris-
HCl buffer (pH 7.2) in the range 200–320 nm. The absorption spectra of
complexes (2)–(4) are presented in Fig. 6 (as representative examples),
the ligand and complex (1) are presented in (Fig. S6b, ESI†) and the
binding constant data are depicted in Table 4. It has been observed that
increasing the concentration of the compounds results in increasing the
absorbance intensity with hyperchromism from 5.30–62.5% for all
compounds with blue-shift (4–5 nm) for the complexes. This informs
that the interaction between BSA and the test compounds is static and
the complexes are in the ground state [89]. These data may also explore
that the structural changes have occurred as a result of non-covalent
binding mode (electrostatic or via hydrogen bonding interaction be-
tween the compounds and BSA). In complex (2), an isosbestic point (not
sharp) was observed at ca. 293 nm, which indicated that the interaction
between BSA and the complex attained equilibrium [90,91]. The bind-
ing strength of the compounds with BSA has been determined using
4.1.2. Emission studies
Fluorescence quenching is considered an important technique to
study the binding affinity of the drug and biomolecules (DNA, RNA and
proteins). The binding affinity of the synthesized compounds cannot be
directly detected by emission spectra, as they don't show any fluores-
cence in presence of ct DNA or tRNA, hence ethidium bromide (EB)
displacement assay has been employed to investigate the binding af-
finity of ctDNA and tRNA to metal complexes. Ethidium bromide (EB) is
the most widely applied fluorophore that strongly intercalates between
base pairs of DNA and RNA to flourish a fluorescence that can be
measured at 600 nm [93].
25
20
15
10
5
1.0
(2)
0.8
0.6
0.4
[BSA]
0.2
0
0.0
1
2
3
4
260
280
Wavelength (nm)
300
1/[complex] 105 M-1
1.0
0.8
0.6
0.4
0.2
0.0
(3)
60
40
20
0
[BSA]
260
280
300
320
0
10
20
30
40
1/[complex] 104 M-1
Wavelength (nm)
2.0
30
(4)
25
20
15
10
5
1.6
1.2
0.8
0.4
0.0
0
[BSA]
260
0
5
10 15 20 25 30 35 40 45
1/[complex] 104 M-1
280
Wavelenght (nm)
300
320
Fig. 6. (Left): Absorption spectra of BSA (15
μ
M) in absence and presence of increasing amounts complexes (2)–(4) (0–30
μ
M) in Tris-HCl buffer pH = 7.2 at 25 ◦C.
Arrow indicates the changes in absorbance upon increasing the compound concentration. (Right): Benesi – Hildebrand linear plot [(A∞-Ao)/(Ax-Ao)] versus 1/
[compound] (Right).
9

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
Table 4
KEB[EB] = Kapp[Com plex]50%
(6)
The BSA binding constants and parameters (Kb, Ksv, Kq, Ka, and n) for the ligand
and its complexes.
Where KEB is 1.0 × 107 and [EB] = 5
μM and [Complex]50% is the
Complex
Absorption
Emission spectroscopy
complex concentration at a 50% reduction of the fluorescence intensity.
spectroscopy
It has been found that all binding constant values (Ksv, Kq, and Kapp
)
Stern-Volmer
Modified
Scatchard
follow the order: (4) > (3) > (1) > (H2L) as shown in Table 3. The
complex (2) is not in order, as the enhancement is observed in Fig. 5b.
These results reveal that, among all compounds, the Ru(II) complex (4)
possesses the strongest affinity to ctDNA and tRNA. By comparing with
some Schiff base metal complexes of triphenylphosphine
[26,48,54,74,95], the reported binding constants are in the order
103–104 Mꢀ 1 range which is similar or lower than those obtained in the
current study.
Stern Volmer
(MSV)
(Kb) × 104
Ksv
Kq
Ka
Kb
n
H2L
(1)
1.7
–
–
–
–
–
3.57
1.50
1.50
×
1.20 × 104
4.4
×
1.35
× 104
1012
1.00
×
105
5.1
×
(2)
(3)
(4)
3.55
3.6
1.00
1.14 × 104
1.19 × 104
1.50 × 104
1.3
× 104
1012
1.60
×
104
7.1
×
4.1.2.2. BSA binding studies. The binding properties of BSA with the
synthesized compounds have been also studied using fluorescence
spectroscopy to confirm whether the BSA-complex formation follows a
static or a dynamic quenching mechanism. The emission spectrum of
BSA shows its characteristic peak at ~340 nm (excited at 290 nm) that is
associated with the existence of tryptophan residues (Trp134 and Trp
213). Trp 134 is located on the surface of the protein, while Trp213
residue present within a hydrophobic binding pocket of the molecules.
Both residues have an intrinsic fluorescence [101,102]. The successive
addition of the compounds to BSA in buffer solution results in quenching
in fluorescence intensity in the complexes (1)–(4). This quenching is
usually assigned to static quenching mechanism as the changes occur in
BSA secondary structure as a result of binding to the compounds, and
this indicates that the formation of a ground state new complex between
BSA and metal complexes rather than dynamic collision [103].
Conversely, in the addition of different amounts of H2L to BSA solution,
the intrinsic fluorescence of protein is enhanced at 450 and 485 nm,
where their interaction may occur in different a different mechanism
[104]. Emission spectra of the ligand (H2L) and complexes (1)–(4) are
shown in (Fig. S6d, ESI†).
1.60
1.16
1.37
× 104
1012
–
104
1.1
×
4.29
–
106
4.1.2.1. Ct DNA and tRNA binding studies.. If the test compound can
displace ethidium bromide from ctDNA/tRNA-EB adduct, the fluores-
cence intensity will be quenched (due to the formation of a non-
fluorescent complex), and the interaction between the small molecule
and the nucleic acid will be intercalation [94,95]. On the other hand, if
the complex failed to displace EB from nucleic acid-EB, the surface or
groove binding may take place. In the present study, while adding
increasing amounts of the complexes to nucleic acid-ethidium bromide
(NA-EB) adduct, the fluorescence intensity at 580 nm is reduced for the
H2L and its (1, 3, 4) complexes (Fig. 7 and H2L (Fig. S6c). Conversely,
the palladium complex (2) showed a different mode of interaction,
where the fluorescence intensity is enhanced upon addition of the
complex (Fig. 7), this indicates that this complex (2) interacted with
ctDNA-EB adduct via another way rather than intercalation (interact via
non-competitive inhibition) and a negative binding constant was ob-
tained [96–98]. Furthermore, in the case of yeast tRNA (Fig. 7), the
emission intensity is decreased at the beginning within the concentra-
The Stern-Volumer (Ksv) and quenching rate (Kq) constants were
obtained from Stern-Volumer Eqs. (4) and (5), respectively. Both values
(Ksv and Kq) were obtained from the plot of Fo/F versus [Q] for complexes
(1)–(3), but not for H2L and complex (4) due to the negative slope and
nonlinearity, respectively. So, the quenching process was further
analyzed using the modified Stern-Volmer Eq. (7) [105] below for the
complexes (1)–(4) (Fig. 8).
tions (5–25
μM) with 17% hypochromism and bathochromic shift of 4
nm, then the enhancement takes place with the concentrations (30–75
μ
M) with 10% hyperchromism and bathochromic shift 7 nm, which also
confirms that interaction of tRNA with complex (2) occurs differently
(intercalation along with surface binding)[98], for the differential
structure of DNA and RNA [99].
Fo
1
1
=
+
(7)
(Fo ꢀ F) faKa[Q] fa
Where Fo and F were mentioned above, fa is the fractional accessible
protein fluorophore, [Q] is the molar concentration of the quencher and
Ka is the effective quenching constant for the accessible fluorophore,
which is similar to the association binding constants for the quencher
acceptor system. It can be obtained from the intercept (1/fa) to slope (1/
faKa) ratio from straight line obtained after plotting Fo/Fo-F against 1/
[Q], as shown in Fig. 8. This linear relationship confirms a static
quenching mechanism has taken place for the interaction of complexes
with BSA. The quenching constant Ka for the complexes follow the
following order (Table 4):
The quenching efficiency can be measured from Stern-Volmer Eq. (4)
[46]:
Fo/F = 1 + Ksv[Q],
(4)
Where Fo and F stand for the fluorescence intensities before and after
the addition of quencher, respectively, Ksv is Stern-Volmer rate constant,
and [Q] is the concentration of the quencher. Ksv is obtained from the
slope of the linear relation between Fo/F and [Q] and the Ksv values were
presented in Table 3.
The dynamic or collision quenching constant (Kq) can be calculated
from Ksv using the Eq. (5):
Additionally, the binding constant (Kb) and the number of binding
sites (n) in static quenching can be calculated using Scatchard Eq. (8)
[106,107] for static quenching,
Log10 [(Fo ꢀ F)/F ] = Log10 Kb + nLog10 [Q]
Where Fo and F are defined the same as above, the values of Kb and n
values are given in Table 4 which obtained from the inverse logarithm of
the intercept and the slope, respectively from the interpolations of Log
[(Fo – F)/F] versus Log[Q]. The binding affinity is found in the order (4)
> (1) > (3) ~ (2) to show that the complex (4) still has the highest
binding constant with nucleic acids and protein as shown in Table 4. For
Ksv = Kqτo
,
(5)
Where τo is the lifetime of the fluorophore in absence of the
quencher, which is often approximate at 10ꢀ 8 s [100]. It was found that
the calculated biomolecular rate constant (Ksv, Table 3) is higher than
(2.0 × 1010 Mꢀ 1Sꢀ 1), the maximum collisional quenching of different
types of biopolymer quencher, which confirm the results obtained from
absorption spectra, the quenching takes place through NA-complex
formation [82].
(8)
Furthermore, the apparent binding constant (Kapp) can be evaluated
using the Eq. (6).
10

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
70
60
50
40
30
20
10
70
60
50
40
30
1.12
1.10
1.08
1.06
1.04
(b) tRNA
(a) ct DNA
(EB+DNA)
(1)
(1)
(EB+RNA)
1.02
0.05 0.10 0.15 0.20 0.25 0.30 0.35
-4
[Q] 10
M
560
580 600
Wavelength (nm)
620
560
580
Wavelength (nm)
600
620
1.00
0.95
0.90
0.85
0.80
70 (2)
60
(2)
6
4
2
0
50
0
2
4
[Q] 10
6
M
8
10
-5
40
30
20
10
0
550
575 600
Wavelength(nm)
625
560
580
600
Wavelength (nm)
620
60
1.8
(EB+DNA)
(EB+RNA)
70
1.7
1.6
1.5
1.4
1.3
1.2
1.1
1.0
(3)
(3)
50
40
30
20
10
60
50
40
30
20
10
0.0
0.2
0.4
0.6
-4
0.8
1.0
1.2
[Q] 10
M
575
600
Wavelength (nm)
625
560
580 600
Wavelength (nm)
620
2.0
60
(EB+RNA)
1.8
1.6
1.4
1.2
1.0
60
(EB+DNA)
(4)
(4)
50
50
40
30
20
10
40
30
20
10
0
0.0
0.2
0.4
[Q]
0.6
0.8
M
1.0
1.2
560
580
600
620
560
580
600
620
Wavelenght (nm)
Wavelength (nm)
Fig. 7. Fluorescence quenching curves of EB- bound; (a) ct DNA and (b) tRNA in presence of complexes (1)–(4);[EB] = 5 μМ, [DNA] = [RNA] = 80 μМ, [complex] =
0–140
μ
M. Inset: plot of Fo/F vs. [Q].
all complexes, the number of the binding sites (n) on BSA was in the
range (1.16–1.37) as shown in Table 4 and Fig. S6e. which indicates the
existence of a single available binding site for the complexes [108].
solvent. In this method, the chromogen free radical (DPPH•) reacts with
the test compounds (antioxidant). Due to accepting a proton or electron
donation, the purple color of the chromogen changes into yellow in the
solution, and this change was followed spectrophotometrically at 517
nm. The reaction can be simply presented as DPPH• + HA → DPPH-H +
A•. The reduction of color depends on the concentration of the com-
pounds which led to the enhancement of radical inhibition. As shown in
Table 5. The DPPH scavenging activity of the complexes is higher than
that of the free ligand indicating that the chelation to the metal ion
enhances the activity, and the radical scavenging activity of the test
compounds as well as ascorbic acid is concentration-dependent as
shown in complex (4) (Fig. 9 as an example), and Fig. S7, ESI†). The
higher antioxidant activity results were taken as a lower value of IC50 of
order (Table 5) which of the order: AA >4 > 3 > 1 > 2 > H2L. The
4.2. Antioxidant activity
The antioxidant activity of the ligand and its complexes has been
evaluated using DPPH radical assay. In DPPH, scavenging activity, the
antioxidant scavenges the free radical by donation of hydrogen, which
can be observed visually, as the color becomes lighter (purple to yellow).
The DPPH assay is one of the most useful methods used for evaluating
the antioxidant activity of the compounds depending on the electron/
proton donation capacity. This assay can be affected by numerous fac-
tors, such as the concentration of hydrogen, metal ions, amount of
11

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
prompted us to evaluate their anticancer activity against human cancer
cell lines; Colorectal carcinoma Colon cancer (HCT116), Hepatocellular
carcinoma (HepG2), and mammary gland breast cancer (MCF7) along
with the human lung fibroblast (WI38) as a non-tumor cell line and
compared to cis-platin as positive control by MTT assay. The cell
viability percentage was plotted against the concentration of the com-
pounds after 24 and 48 h of treatment (Fig. S8, ESI†). As shown in these
figures, the cell viability decreased in a concentration-dependent
manner and incubation period as well as the type of cancer cell
[109,110]. The data are presented based on IC50 (the half-maximum
inhibitory concentration) and selectivity index (SI) = IC50 of normal
cell/IC50 of cancer cell) [111] as shown in (Figs. 10 and 11, and
Table S3, S4), and respectively. The IC50 values showed the higher ac-
tivity of metal complexes in a 48 h period, this may be attributed to the
high kinetic stability of the complexes in culture media [112]. It has
been shown that the complex (4) showed the lowest IC50 (highest
(4
3
1
(2 )
>
50
35
(2)
(1)
40
30
20
10
0
30
25
20
15
10
5
0
0
5
10
15
20
25
30
0
5
10
1/[Q]
15
20
25
30
4
-1
1/[Q] 10
M
16
12
8
20
16
12
8
(4)
(3)
4
4
0
0
0
5
10
1/[Q]
15
20
25
30
0
5
10
15
20
25
30
4 -1
1/[Q] 10 M
cytotoxic activity) on HepG2 (26.14 ± 2.0
1.9 M) and the highest selectivity index, 3.75 and 4.45, respectively.
On the other hand, nickel(II) complex (1) showed the lowest IC50 on
MCF7 (14.35 ± 1.4 M) highest selectivity index 5.59. The cytotoxic
activity of the compounds was found to follow the order on the basis of
μM) and HCT116 (22.02 ±
μ
Fig. 8. Modified Stern –Volmer plot for the quenching of BSA fluorescence
complexes (1)–(4).
μ
Table 5
IC50
:
Antioxidant parameters in DPPH scavenging activity data of the ligand and its
HepG2: (4) > (3) > (1) > (2) > (H2L)
complexes.
Compound
HCT116: (4) > (3) > (1) > (H2L) > (2)
EC50
ARPa
Stoichiometrya
No of reduced DPPH
a
MCF7: (1) > (4) > (3) > (H2L) ≈ (2)
IC50
a
In comparing ruthenium(II) complexes with platinum(II) complexes,
it is known that ruthenium complexes have several characteristic
properties over the platinum complexes such as, octahedral geometry,
which induces the structure diversity as compared to square planar
platinum(II) complexes, multiple accessible oxidation states (+II to +IV)
under physiological conditions, and kinetic lability of ruthenium(II)
species which facilitate the ligand exchange reaction and leads to more
rapid interaction with biological targets molecules [113,114]. In addi-
tion to the concentration of the tested compound and the incubation
period, the anticancer activity may also be affected by the structure-
activity relationship [115]. For instance, among the three square
planar complexes (1)–(3), the Pt(II) complex exhibited a better cytotoxic
H2L
(1)
408
235
270
91
4.08
2.35
2.7
0.25
0.43
0.37
1.09
1.82
2.32
8.16
4.7
0.12
0.21
0.18
0.55
0.91
1.16
(2)
5.4
(3)
0.91
0.55
0.43
1.82
1.1
(4)
55
Ascorbic
acid
43
0.86
a
EC50 (μmol of antioxidant/μmol DPPH), ARP = 1/EC50, stoichiometry = 2 ×
EC50, no. of reduced DPPH = 1/stoichiometry.
complex (4) exhibited the highest scavenging activity among all com-
pounds (IC50 = 55 μM), which is similar to that reported for potent
activity (in terms of IC50 values) towards HePG2 (28.94 ± 2.4
μM) and
antioxidants of ruthenium(II) and vanadium(IV) complexes containing
similar O,N,S-donor ligand [48]. The other antioxidant parameters;
EC50, ARP, stoichiometry and number of reduced DPPH• radicals have
been calculated, and we found that the lower IC50 and EC50 values, the
greater ARP values [48].
HCT116 (31.11 ± 2.8 M) after 48 h of treatment, while it showed
μ
moderate activity against MCF7 (44.76 ± 3.4
μM), but the Ni(II) com-
plex (1) exhibited a high selectivity index to all tested cancer cell lines
(1.59–3.03), and presented a safely profile towards the normal cell line
(WI38, IC50 = 80.30 ± 4.2 μM). This may attribute to the structure di-
versity of cationic Pt(II) complex (3), where, the ligand acts as a mono-
negative bidentate rather than bi-negative tridentate in the rest of the
complexes. Among all the compounds, Ru(II) and Ni(II) complexes dis-
played interesting activity towards cancer cells as their selectivity
indices (3.46–5.59) are higher than 3.0, comparable to that of cisplatin
4.3. In vitro cytotoxic activity (MTT assay)
The interesting results collected from ct DNA, tRNA, BSA binding
affinity, and antioxidant properties of the synthesized compounds have
0
25
50
1
(a) 1.2
(b) 500
(4)
H2L
75
1.0
100
2
400
(1)
(2)
0.8
300
3
(3)
0.6
200
(4)
4
0.4
AA
5
100
0.2
0
0.0
H2L (1) (2) (3) (4) AA
500
550
600
650
Compounds
Wavelength (nm)
Fig. 9. (a) Changes in UV–Vis spectrum of DPPH (100
μM) in the presence of complex (4) (25–100 μM) after 30 min incubation in DMSO at room temperature, (b)
DPPH scavenging activity of the compounds on the bases of IC50
.
12

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
HePG2
24 h
WI38
100
80
60
40
20
0
100
80
60
40
20
0
48 h
24 h
48 h
H2L
(1)
(2)
(3)
(4) Cisplatin
H2L
(1)
(2)
(3)
(4) Cisplatin
Compound
Compound
HCT116
MCF7
80
60
40
20
0
80
60
40
20
0
24 h
48 h
24 h
48 h
H2L
(1)
(2)
(3)
(4) Cisplatin
H2L
(1)
(2)
(3)
(4) Cisplatin
Compound
Compound
Fig. 10. Cytotoxicity (IC50 values) of the (Ligand (H2L), complexes (1)–(4) and cisplatin) at 24 and 48 h against WI38, HepG2, HCT116, and MCF7 cell lines.
24 h
48 h
4
3
2
1
0
6
5
4
3
2
1
0
HCT116
HePG-2
MCF7
HCT116
HePG-2
MCF7
H2L
(1)
(2)
(3)
(4) Cisplatin
H2L
(1)
(2)
(3)
(4) Cisplatin
Compound
Compound
Fig. 11. Selective indices of the ligand and complexes towards cancer cell lines at 24 and 48 h.
(< 1.5). This provide a high selectivity towards particular human cancer
cell lines [116]. It is worth noting that the selectivity of the compounds
towards cancer cells can be determined by the selectivity index (SI),
which becomes an essential prerequisite to estimate the capability of the
test compound to kill the cancerous cells without harming the healthy
one [117]. The higher the SI, the higher selectivity of the compounds. It
has been reported that the compounds which have SI ≥ 3 are considered
highly selective towards a specific cell line, so it becomes a curial
requirement for the development of a chemotherapeutic agent with
good selectivity towards cancer cells [118,119].
The complexes provide comparable cytotoxic levels with some other
reported complexes for the selected tumor cell lines. It has been reported
that a series of Ni(II) triphenylphosphine complexes with thioamide
derivatives [120] or aroylhydrazone [95] has shown IC50 of 20–40 and
13

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
cancer [123]. To distinguish between apoptotic and necrotic cell death,
cell cycle assays by Annexin has been carried out by following cyto-
metric technique. Based on ct DNA/tRNA and BSA interaction studies,
along with antioxidant and cytotoxic properties of the synthesized
compounds, complexes (1) and (4) exhibited better activities compared
to their parent ligand and other complexes. So, both complexes were
further selected for investigating their efficacy in cell death in MCF7 and
HCT116 cells, respectively. The induction of apoptosis was studied by
the colorimetric diphenylamine (DPA) assay and flow cytometric anal-
ysis of DNA contents.
40
30
20
10
0
4.4.1. DNA fragmentation
The DNA fragmentation was quantitatively determined using the
colorimetric diphenylamine assay (DPA) [124]. The complexes (1) and
(4) were selected for DNA fragmentation analysis. The fragmentation
percentage data were presented with respect to the control (untreated
cells) in Fig. 12 and Table S5. The breast cancer cells (MCF7) were
treated with complex (1) at IC50 = 14.35 ± 1.4 μM, it was found that the
DNA fragmentation percent was 34.09 ± 1.86% (control, 5.24 ±
0.29%). In the case of treatment of colon cancer cells (HCT116) with
complex (4) at IC50 = 22.02 ± 1.9 μM, the fragmentation percent was
29.75 ± 1.62 (control, 4.5 ± 0.25). This concludes that the fragmenta-
tion percent of each complex compared to its selected cell lines increased
ca. 6.5-fold compared to control cells for 48 h. These findings may
suggest that complexes promote cell death via DNA damage [125].
7
7
6
6
1
1
F
F
1
1
4.4.2. Apoptosis analysis by flow cytometry
C
C
M
T
T
M
Flow cytometry is considered one of the most effective methods to
analyze cell death and evaluate the therapeutic efficacies of anti-cancer
drugs [126]. Most of the anticancer agents inhibit cancer cell prolifer-
ation by induction of apoptosis in tumor cells [127]. To determine the
cellular DNA content and the growth inhibition mechanism on tumor
cells, the flow cytometry technique has been employed.
/
C
C
)
H
H
1
/
(
)
4
(
Fig. 12. DNA fragmentation using DPA assay for complex (1) on MCF7, and
complex (4) on HCT116, MCF7, and HCT116 cell lines as control.
4.4.2.1. Cell apoptosis. The apoptotic cell death in MCF7 and HCT116
was investigated by flow cytometry using co-staining of annexin-V FITC
and (PI). Generally, the cells are distributed in four different quadrants
(Q1-Q4) as shown in Fig. S9, ESI†, Q1 (live cells, lower left quadrant),
Q2 (early apoptosis, lower right quadrant), Q3 (late apoptosis, upper
right quadrant) and Q4 (necrosis, upper left quadrant). The MCF7 cells
were treated with complex (1) at IC50 concentration for 48 h. In control
(untreated MCF7 cells), the living cell % was 98.15% and apoptotic %
was 1.03%. On the treatment of MCF7 cells with the complex (1), the
apoptotic percent increased to 24.58% (2.33% early and 22.25% late
apoptosis), i.e. 23.86-fold (control, 1.03%), this indicates that the
complex can induce cell death via apoptosis. It has been also noted that
32.6 μM, respectively against MCF7 cell line and they showed higher
toxicity to cancer cells than cis-platin, which support the Ni(II) complex
possesses the potential to act as effective metal-based anticancer drugs.
4.4. Apoptotic activity
Apoptosis and necrosis are two major forms of cell death [121,122]
Apoptosis (or programmed cell death) plays an essential role in regu-
lating cell death by controlling the number and size of the cells. Necrosis
(sudden cell death), usually causes spillage of cell contents into sur-
rounding tissues, consequently, cell damage takes place. Inappropriate
regulation of apoptosis can result in various forms of diseases including
MCF7
MCF7 (control)
(a)
(b)
HCT116
30
20
10
0
HCT116 (control)
HCT116 /(4)
24.58
25
20
15
10
5
MCF7/ (1)
22.25
19.11
9.71
5.77
4.16
2.33
2.25
1.03
0.82
1.61
0.66
0.37
0.68
0.52
0.16
0
Total ap. Early ap. Late ap. Necrosis
Apoptosis/necrosis
Total ap. Early ap. Late ap. Necrosis
Apoptosis/necrosis
Fig. 13. Apoptosis/necrosis cell percentages of: (a) MCF7 with complex (1), (b) HCT116 with complex (4) after 48 h incubation period at IC50 values.
14

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
Fig. 14. Percentage of cell populations among cell phases in: (a) untreated (control) and treated cells MCF7 with complex (1), (b) untreated (control) and treated
HCT116 with complex (4) at IC50 concentrations for 48 h.
the number of cells in Q4 was increased to 9.71 (control, 0.82%)
demonstrated that the cell death mode can, to a slight extent, takes place
via necrosis. While, the treatment of HCT116 cells with complex (4) at
IC50 concentration for 48 h, the apoptosis percent increased to 5.77%
(1.61% early and 4.16% late apoptosis), i.e. increased 8.49-fold
compared to untreated cells (0.68%), which indicates that the cells
may be undergoing apoptosis [128]. It has been also observed that the
HCT116 cells increased in necrosis quadrant to 19.11%, by 8.49-fold
compared to untreated cells (2.25%), and this result following early
reports [129] which demonstrated that Ru(II) complexes can majorly
induce cell death through necrosis. Apoptotic cell % is shown in Fig. 13
and (Fig. S9 and Table S6, ESI†).
consists of some phases such as quiescent (G0), interphase (gap1 (G1),
synthesis (S), and gap2 (G2)), and mitotic phase (M) [130]. During the
cell dividing progression, the cells undergo a transition through (G1) →
(S) → (G2) → (M) phases to synthesize DNA, cell division followed by
mitosis process. However, if the cells enter the inactive phase (G0 or Pre-
G1 phase), this means that the cells enter a resting state (quiescent),
where there is no dividing, no proliferation [126].
The effect of complexes (1) and (4) have been studied on the cell
cycle of MCF7 and HCT116 cells, respectively. Complex (1) exhibited a
significant increase at the pre-G1 phase (~32.44% versus untreated
cells) (Fig. 14, Fig. S10 and Table S7). Earlier reports indicated that
disruption of the pre-G1 phase in the cell cycle might be one of the
reasons for apoptosis [131]. The MCF7 cells were arrested in the S phase,
since there is an increase in the number of cells by ~14.12% (49.38%,
control 35.26%), but an insignificant change (~2.01%) was found at the
G2/M phase. This observation along with the DNA fragmentation
studies may suggest that the nickel(II) complex (1) can stop the MCF7
4.4.2.2. Cell cycle analysis. Based on the inhibitory effect on cell pro-
liferation, the effect of complex (1) on the cell cycle of MCF7 cells was
evaluated. The analysis of the cell cycle phases was carried out by flow
cytometry at IC50 concentration for 48 h. Cell cycle phase distribution
15

S.A. Elsayed et al.
Journal of Inorganic Biochemistry 223 (2021) 111549
proliferation by induction of apoptotic cell death. On the other hand, on
the treatment of the HCT116 cells with complex (4), there was a
decrease in the number of cells in G1 phase (38.49%, control 49.13) and
S phase (41.32, control 46%) as shown in Fig. 14. However, the number
of cells increased in the G2/M phase (20.19, control 4.87%) which
means cell cycle arrest at the G2/M phase. That increased in G2/M leads
to the accumulation of the cells in this stage indicating the inhibition of
HCT116 cells from replication again and this may be due to the strong
interaction between the complex (4) and the DNA that was previously
confirmed by ct DNA binding assay. From the former data, we observed
that the complex (4) can interact with the cellular protein, as observed
from BSA binding affinity, along with strong interaction with nucleic
acids which may break the DNA and make unpacked DNA rather than
fragmentation, which prevent cancer replication and arrest the cell cycle
at G2/M. Moreover, the HCT116 cells treated with ruthenium(II) com-
plex have increased the pre-G1 population which may appear from
loosely packed DNA that happened and resulted in a high increase in cell
percentage at necrotic quadrant, when analyzed by annexin assay,
revealed the induction of necrotic cell death.
scavenging activity and showed that the complexes have better antiox-
idant activity compared to the parent ligand. Anticancer activity against
four human cell lines has been demonstrated, indicating that all com-
plexes are toxic to cancer cell lines and safe towards the normal one.
Finally, complexes (1) and (4) were selected for DNA fragmentation,
apoptotic activity and cell cycle analysis. Our data revealed, complex (1)
induced apoptotic cell death of the MCF7 cells by fragmentation of the
DNA after binding with it, increased pre-G1 phase and arrested the cell
cycle at S phase. Also, complex (1) has antioxidant activity that helped
in scavenging the excess ROS and preventing healthy cell damage along
with inhibiting the cancer cell replication. For complex (4), it exhibited
a good anti-proliferative effect against HCT116 cells that arrested at the
G2/M phase. Necrosis induction has also occurred, that may be due to
the intercalation and surface binding with DNA strands which can un-
pack the two strands leading to sudden cell death. Also, the increased
safety of the complex (4) on the healthy cells was confirmed by its potent
antioxidant activity and elevated SI values.
Declaration of Competing Interest
Finally, our data revealed the apoptotic cell death of the MCF7 cells
treated with complex (1) was confirmed by high DNA fragmentation
percentage and the good binding affinity with biomolecules (DNA, RNA,
and BSA). When complex (1) made strong binding with nucleic acid
resulting DNA damage and hence the S phase in the cell cycle was
arrested, as increased its percentage, and the pre-G1 phase has appeared
with high percentage as a result of apoptotic cell death. The MCF7
apoptotic cell death was also confirmed by the annexin V/PI stain, after
treated with complex (1), with an increase in the total apoptosis %
23.86-fold than control. Cancer cells induced high reactive oxygen
species (ROS) as increased metabolism due to increased proliferation
rate [132]. The antioxidant potency of the complex (1) can scavenge the
elevated ROS and inhibited the cancer cell replication and that increased
the safety of that complex on healthy cells which is confirmed by the
high SI and good cytotoxicity results.
The authors declare that they have no known competing financial
interests or personal relationships that could have appeared to influence
the work reported in this paper.
Acknowledgments
Authors acknowledge support from the Ministry of Higher Education
of Egypt for CIQAP (CP3-016-MAN) project research equipment funds.
Special thanks go to Prof. Alessandro Caselli, Prof. of Inorganic Chem-
`
istry, Dipartimento di Chimica, Universita degli Studi di Milano, Milano,
Italy, for giving us the opportunity to carry out the X-ray analysis and
Mass spectra of complexes.
Appendix A. Supplementary data
On the other hand, the treatment with complex (4) induced an anti-
proliferative effect against HCT116 cells via making intercalation and
surface binding with DNA strands which can unpack the two strands
leading to replication inhibition which confirmed by cell cycle arrest at
the G2/M phase. Also, the necrotic cell death that appears in annexin V/
PI stain data may be attributed to high binding affinity with bio-
molecules, which can make irreversible binding, and results in the
sudden death of the cancer cells (necrotic cell death). However, the DNA
fragmentation data of the complex (4) may due to the intercalation with
DNA that led to unpacked strands rather than truly fragmentation
because DNA fragmentation didn't contribute to necrotic cell death. One
of the probable hypotheses of the appearance of the pre-G1 phase is the
accumulation of dead cells that are affected by the complex (4) treat-
ment through necrotic death [133,134]. The accumulated dead cells
were due to sudden unpacked DNA strands and they may reduce the
fluorescence intensity of the G1 population during cell cycle analysis
leading to the appearance of the pre-G1 phase [135,136] Concurrently,
the high antioxidant activity of complex (4) has increased its safety on
healthy cells confirmed with high SI and cytotoxicity values.
Supplementary data to this article can be found online at https://doi.
org/10.1016/j.jinorgbio.2021.111549.
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