Synthesis, structures and anticancer potentials of

European Journal of Medicinal Chemistry 155 (2018) 609e622

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Research paper

Synthesis, structures and anticancer potentials of platinum(II) saccharinate complexes of tertiary phosphines with phenyl and cyclohexyl groups targeting mitochondria and DNA Veysel T. Yilmaz a, *, Ceyda Icsel a, Omer R. Turgut a, Muhittin Aygun b, Merve Erkisa c, Mehmet H. Turkdemir a, Engin Ulukaya c a b c

Department of Chemistry, Faculty of Arts and Sciences, Uludag University, 16059, Bursa, Turkey Department of Physics, Faculty of Sciences, Dokuz Eylul University, 35210, Izmir, Turkey Department of Medicinal Biochemistry, Faculty of Medicine, University of Istinye, 34010, Istanbul, Turkey

a r t i c l e i n f o

a b s t r a c t

Article history: Received 24 April 2018 Received in revised form 12 June 2018 Accepted 13 June 2018

A series of new Pt(II) saccharinate complexes containing PR3 ligands (PPh3, PPh2Cy, PPhCy2 and PCy3) with progressive phenyl (Ph) replacement by cyclohexyl (Cy) were synthesized and structurally characterized by IR, NMR, ESI-MS and X-ray diffraction. The anticancer activity of the complexes was tested against human breast (MCF-7), lung (A549), colon (HCT116), and prostate (DU145) cancer cell lines as well as against normal bronchial epithelial (BEAS-2B) cells. Trans-configured complexes 1, 3 and 5 emerged as potential anticancer drug candidates. The mechanism of action of the potent complexes was then investigated in detail. The three complexes interacted with DNA by groove binding and with HSA via hydrophobic IIA subdomain. Furthermore, the complexes cleaved plasmid DNA efficiently. Cellular uptake studies in MCF-7 cells showed that the biologically active complexes were mainly localized in cytoplasm. The cytotoxic activity was a function of the lipophilicity and cellular accumulation of the complexes. As determined by M30, Annexin V and Caspase 3/7 activity assays, the complexes induced apoptosis in MCF-7 and HCT116 cells. Mechanistic studies showed that the potent complexes cause excessive generation of reactive oxygen species (ROS) and display a dual action, concurrently targeting both mitochondria and genomic DNA. © 2018 Elsevier Masson SAS. All rights reserved.

Keywords: Platinum(II) Saccharinate Tertiary phosphine Anticancer activity Apoptosis

1. Introduction In the last two decades, investigations to discover new chemotherapeutic metal-based complexes alternative to the clinicallyused platinum based anticancer drugs such as cisplatin, carboplatin and oxaliplatin have received great attention to overcome their limitations and to increase their selectivity [1e6]. It is well established that these platinum based agents target DNA. They hydrolyze in the aqueous solution giving cationic species, which bind to DNA covalently forming intrastrand cross-links [7,8]. The formation of stable complexes between DNA and these compounds blocks DNA replication and transcription, consequently triggering death of cancer cells. On the other hand, metal complexes interacting with DNA non-covalently were also shown to have potential as

* Corresponding author. E-mail address: [email protected] (V.T. Yilmaz). https://doi.org/10.1016/j.ejmech.2018.06.035 0223-5234/© 2018 Elsevier Masson SAS. All rights reserved.

promising anticancer drugs [9e11]. The selection of planar ligands or functionalization of ligands with planar groups often facilitates strong interactions of metal complexes with DNA through intercalation or groove binding. The use of the platinum complexes prescribed in the treatment of cancer is limited by their severe side effects [12]. The nanofunctionalization of the platinum-based drugs has received much attention due to controlling their delivery to targets, enhancing their efficacy and reducing the side effects [13e15]. In other words, the main aim of nanoformulations is to increase their selectivity for cancer cells in order to achieve an optimum pharmacological profile. The nanoscale drug formulations containing the clinically used platinum drugs exhibited promising in vitro and in vivo pharmacological properties so that some of these formulations are currently in clinical trials [16]. Our research group has long been involved in synthesis and biological activity of various metal complexes interacting with DNA non-covalently. One of the ligand we used is a well-known artificial

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sweetener, saccharin (sacH, o-sulphobenzimide). SacH readily loses the imine hydrogen in solutions. The corresponding anion saccharinate (sac) behaves as a versatile polyfunctional ligand, owing to the presence of the negatively charged imino nitrogen atom and the carbonyl and sulfonyl oxygen atoms, and forms metal complexes from mononuclear to coordination polymers [17]. Recently, we have focused on the synthesis of palladium and platinum complexes of sac [18e25]. In vitro and in vivo cytotoxic activities of some of these complexes have also been evaluated with encouraging results [24e28]. In addition, several Pt(II) complexes of sac with potent anticancer activity were reported by other research groups [29,30]. In this work, triphenylphosphine (PPh3) and its three derivatives bearing one, two, and three cyclohexyl (Cy) groups were selected as ancillary ligands. In fact, only one example of platinum(II) complex of sac with PPh3, namely cis[PtCl(sac)(PPh3)2] [31] was reported earlier. Therefore, in order to fill the gap in this field, we decided to design and synthesize a set of platinum complexes of sac with the tertiary phosphine ligands (PR3) with progressive replacement of the phenyl (Ph) group by cyclohexyl (Cy) group (Scheme 1). In vitro cytotoxicity of the complexes were screened against several human cancer cell lines. The accumulation of the complexes in the cells were explored. In addition, we also included studies concerning the apoptosis induction ability of the most cytotoxic complexes, their implications in ROS production, change of mitochondrial potential, and damage to genomic DNA. Moreover, the interaction of the complexes with DNA and protein were studied in detail, using a wide range of methods. 2. Results and discussion 2.1. Synthesis and characterization Seven platinum(II) sac complexes with the tertiary phosphines were prepared using the different synthetic strategies shown in Scheme 1. The precursors [PtCl(sac)(COD)] and [Pt(sac)2(COD)], originally synthesized by our research group [32], were used in the preparation of trans-[Pt(sac)2(PPh3)2] (1), cis-[PtCl(sac)(PPh2Cy)2] (2), trans-[Pt(sac)2(PPh2Cy)2] (3), trans-[PtCl(sac)(PPhCy2)2] (4), trans-[Pt(H)(sac)(PphCy2)2] (5), trans-[PtCl(sac)(Pcy3)2] (6) and trans-[Pt(H)(sac)(Pcy3)2] (7) through the replacement of COD (1,5cyclooctadiene) by the phosphines. Complexes 1e7 were isolated in moderate to good yields (from 30 to 90%) as bright yellow solids that were air and moisture stable. The complexes are highly soluble in MeOH, MeCN, CH2Cl2, CHCl3, DMSO and DMF, but exhibit very poor solubility in water.

Conductivity measurements in MeOH indicate the nonelectrolyte behavior of the complexes [33]. The complexes were characterized by elemental analysis, spectroscopic methods. The electronic spectra of the complexes display the typical intraligand absorptions due to p/p* and n/p* transitions centered at ca. 225, 265 and 272 nm, respectively. The IR spectra of 1e7 show the sharp bands of the carbonyl group of sac in the range of 1683e1663 cm
1, while the asymmetric and symmetric vibrations of the sulphonyl group of sac appear at ca. 1245 and 1150 cm
1, respectively. The phosphine ligands are characterized by the presence of the medium-to-strong absorption bands centered at ca. 1000 and 517e502 cm
1 assigned to the symmetric and asymmetric vibrations of the PeC group, respectively, in addition to the n(CH) vibrations around 2900 cm
1. In the IR spectra of 5 and 7, the medium absorption bands at 2239 and 2217 cm
1 were attributed to the n(PteH) stretchings [34], correlating with the electron-withdrawing properties of PPhCy2 and Pcy3 ligands at the cis position [35]. The NMR spectral data obtained in DMSO‑d6 are presented in the experimental section with the corresponding assignments (see also Fig. S1 for the spectra). In the 1H NMR spectra of the complexes, the signals corresponding to the phenyl protons of both sac and phosphine ligands appear in the region of ca. 6.94e8.17 ppm, while the protons of Cy in the phosphines are observed in the range of 2.40e0.44 ppm as multiplets. The presence of the PteH bonds in complexes 5 and 7 is evidenced by the triplets at
17.81 and
18.18 ppm, respectively, being consistent with the 1H NMR spectra of the Pt (II) complexes of various phosphines with the same bond [34,36,37]. As shown in Scheme 1, the hydrido ligand is trans to the anionic N atom of the sac ligand. The observed substantial negative chemical shifts in the spectra of 5 and 7 clearly indicate that the sac ligand has a relatively weak trans influence effect [36]. In the 31P{H} NMR spectra of the complexes, the signals of phosphine ligands experience strong deshielding upon coordination to Pt (II). The 31P
195Pt coupling is visible in the spectra of the Pt (II) complexes with 1JPteP values in the range of 2138e2754 Hz. The coupling constants are in accordance with those trans-Pt (II) complexes of phosphines [38], since the cis[PtCl2(PR3)2] complexes present much higher 1JPteP values around 3500 Hz [39]. The ESI-MS spectra of 1
7 show peaks corresponding to the species [M þ Na]þ, [M
sac]þ and [M
Cl]þ (Fig. S2). 2.2. X-ray structures Single crystals of 2 and 5e7 suitable for X-ray crystallography were obtained by the slow evaporation of their concentrated solutions. Details of the structure analysis are given in Table S1. The

O S

O O

Cl

PPh2Cy PPh2Cy

S

S

(2) [Pt(sac)2(COD)]

O

L

Cl

N

O

O

O

L = PPh 3 (1) PPh2Cy (3)

L CH 3CN, MeOH 65 oC

Pt S

L

O

L CH3CN, MeOH 65 oC

Pt N

O

O

[PtCl(sac)(COD)]

O

Pt N

N

L

L O

O

L

Scheme 1. Synthetic route of complexes 1
7.

L

N S

L = PPhCy 2 (4) PCy 3 (6)

H Pt

O

O

L = PPhCy 2 (5) PCy 3 (7)

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molecular structures of the complexes are shown in Fig. 1, while selected bond distances and angles are listed in Table S2. As shown in Fig. 1, the platinum (II) cations show a distorted square planar geometry. The sac ligand in the complexes is N-coordinated via the negatively charged N atom. In addition, the chloride ligand occupies the cis position to the sac ligand in 2, but the trans position in 6. Interestingly, 5 and 7 contain a hydrido ligand trans to sac. Highly disordered DMSO in 5, and two DMF molecules in 6 were eliminated. The complexes containing two monophosphine ligands usually exhibit a trans configuration, but 2 has two phosphine ligands at the cis position. The Pt
N bond distances range from 2.048 to 2.157 Å, similar to those reported for the Pt complexes with sac [18e25,29e31], while the Pt
P and Pt
Cl bond lengths are in good agreement with values found in cis-[PtCl (sac) (PPh3)2] [31]. The Pt
H bond distances in 5 and 7 are 1.25 (8) and 1.20 (4) Å, respectively, and fall in the range of the similar bonds in the Pt complexes deposited in the Cambridge Structural Database (CSD) [40]. The packing of molecules in the solid state is dominated by a number of weak C
H$$$O (sulfonyl) hydrogen bonds. In addition, the chlorides in 2 and 6 interact weakly with neighboring complexes, forming several C
H$$$Cl interactions. 2.3. Cytotoxic activity The antiproliferative activity of complexes 1
7 was tested against different human cancer cell lines such as breast (MCF-7), lung (A549), prostate (DU145), colon (HCT116) and human bronchial epithelial cells (BEAS-2B). 20 mM of each complex was tested initially for the toxicity screening using Sulforhodamine B (SRB) assay. In addition, the previously reported Pt (II) complex, cis[PtCl(sac)(PPh3)2] [31], was also included in the toxicity test. The

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SRB assay showed that only three platinum (II) complexes (1, 3 and 5) demonstrated the most efficient antigrowth effect, being promisingly cytotoxic against all the cell lines tested (see Fig. S3 for viability screening). These complexes were subjected to further testing using the most sensitive cytotoxicity assay ATP at different concentrations ranging from 0 to 40 mM to determine the dose of the complexes which causes death in 50% of cells (IC50) at 48 h (Table 1 and Fig. S4). The anticancer activity of the complexes was also compared with that of cisplatin. Overall, the three potent platinum(II) complexes have remarkable growth inhibitory effects. In all the cell lines, complex 3 presents higher cytotoxic activity than cisplatin and the rest of the complexes. In addition, 1 shows better inhibition effects than cisplatin in MCF-7 and HCT116 cancer cells, while complex 5 displays moderate cytotoxicity in the tested cell lines, being more active against MCF-7 cells. In addition, it should be noted that all complexes display poor selectivity for the cancer cells over the noncancerous BEAS-2B cells. The complexes containing a chloride ligand (2, 4 and 6) are expected to exhibit notable cytotoxicity due to the replacement of this ligand inside the cells similar to cisplatin. The chloride ligand in these complexes is usually positioned trans to sac and the weak trans effect of the sac ligand substantially decreases the lability of the chloride ligand, thereby resulting in a low toxicity of these complexes. As discussed above, the structures of the three potent Pt(II) complexes reveal a trans configuration and the mechanism underlying the biological activity of these complexes is most likely different from that of cisplatin. The potential factors involved in the mechanism underlying the cytotoxic effects of these trans-configured complexes were therefore investigated using different methodologies.

Fig. 1. Structures of 2 and 5e7. Highly disordered DMSO in 5, and two DMF molecules in 6 were removed by squeezing.

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Table 1 Cytotoxicity of 1, 3, 5 and cisplatin against four human cancer cell lines. IC50 (mM)a 1 8.4 ± 0.1 10.1 ± 1.7 14.5 ± 0.5 13.0 ± 0.7 8.1 ± 0.3

MCF-7 A549 DU145 HCT116 BEAS-2Bb

3 3.8 ± 0.4 4.0 ± 0.1 4.6 ± 0.1 4.8 ± 0.1 2.4 ± 0.1

5 17.0 ± 3.7 30.1 ± 1.0 37.7 ± 0.1 36.7 ± 0.2 27.7 ± 1.2

cisplatin 24.0 ± 4.0 2.5 ± 0.9 9.8 ± 4.5 15.5 ± 2.3 4.6 ± 0.2

a The IC50 values were determined by the ATP assay after 48 h of exposure to the platinum complexes. Data represent the mean ± SD of at least three independent experiments. b Noncancerous cells.

2.4. Stability in solution A potential anticancer drug is expected to be stable in solution to produce accurate activity. MeOH and DMSO were used to prepare the solutions of the complexes for binding affinity and cytotoxic activity studies, respectively. Thus, the stability of 1, 3 and 5 was determined in MeOH, DMSO and saline (an aqueous solution of 0.9% NaCl, approximating physiological conditions). Owing to the insolubility of all complexes in water, they were first dissolved in a small amount of MeOH and then mixed with saline. The UV absorption spectra of the MeOH and DMSO solutions recorded at different time intervals (0, 24 and 48 h) remain unchanged with increasing time, indicating the stability of these complexes in these solutions (Fig. S5). However, a considerable decrease of the intensity of the UV bands is observed in the case of saline after 24 and 48 h, and associated with the slight precipitation of the Pt complexes in the solution. The precipitates obtained from the saline solutions after 48 h were characterized by UV, 1H and 31P spectroscopy. As shown in Fig. S6, the spectra of the solids are essentially identical to those of the corresponding complexes, indicating that the complexes maintain their structure without substantial degradation in the saline environment. 2.5. Lipophilicity and cellular uptake To quantify the lipophilicity of the platinum complexes, wateroctanol partition coefficients (P) were measured using the shakeflask method. The resulting log P values are reported in Table 2. More positive log P values correspond to higher lipophilicity. The measured log P of cisplatin is consistent with literature values [41]. The overall order of lipophilicity follows the sequence 3 > 1 > 5 > cisplatin. The sac ligand contains both polar and hydrophobic groups. In addition, the cyclohexane is more lipophilic than benzene [42], Thus, the introduction of Cy groups in the tertiary phosphines tunes the lipophilicity of the present complexes in agreement with the Ag(I) complexes with the same monophosphines [43]. It is most likely that the presence of the hydrido ligand significantly reduces the lipophilicity of 5. The cellular accumulation of the platinum complexes (1, 3 and 5) and cisplatin in MCF-7 cells was investigated for possible

Table 2 Lipophilicity and cellular uptake of the potent complexes and cisplatin by MCF7 cells (ng Pt/106 cells).

membrane cytosol nucleus cytoskeleton Total log P

1

3

5

cisplatin

34.6 ± 0.6 53.4 ± 1.3 20.9 ± 0.9 98.2 ± 1.6 207.1 ± 2.3 1.01 ± 0.04

44.9 ± 2.6 191.2 ± 4.8 23.9 ± 0.8 60.9 ± 1.7 320.9 ± 5.8 1.28 ± 0.03

3.0 ± 0.5 5.4 ± 0.3 1.0 ± 0.1 54.2 ± 2.5 63.6 ± 2.6 0.92 ± 0.06

4.0 ± 0.3 3.9 ± 0.1 3.2 ± 0.1 2.6 ± 0.2 13.7 ± 0.4
2.28 ± 0.07

relationship between the cellular uptake, lipophilicity and cytotoxicity of these complexes. After 4 h treatment with the complexes (25 mM), MCF-7 cells were divided into cytosolic, membrane, nuclear and cytoskeletal fractions. Then, the Pt content of each fraction was determined by differential pulse stripping voltammetry. The three platinum complexes were taken up by cells more effectively than cisplatin (Table 2). Most of the platinum is found in the cytoplasm (cytosol þ cytoskeleton). It seems that the cellular uptake is a function of lipophilicity of the platinum complexes, reflecting higher diffusion ability of the hydrophobic complexes through the cell membrane. Moreover, higher cellular uptake of the complexes correlates with their stronger cytotoxic activity.

2.6. DNA binding studies The platinum-based anticancer drugs are known to target the nuclear DNA. For this reason, various biochemical and biophysical methods were employed to quantify and dissect the binding affinity and binding modes of the potent platinum complexes towards fish sperm (FS) DNA. The changes in the structure of DNA due to interaction with drugs can be assigned either by hypochromism or by hyperchromism in the UV spectra. Significant increases in the absorption intensity of FS-DNA at lmax ¼ 260 nm are recorded upon addition of increasing amount of the platinum(II) complexes (1, 3 and 5), suggesting non-covalent groove binding between DNA and the complexes (Fig. 2 and Fig. S7). The hyperchromicity is accompanied by a few nm blue shifting. The binding constants (Kb) of 1, 3 and 5 towards FS-DNA evaluated by monitoring the changes in absorbance of the UV spectra show that the binding affinity of the complexes increases in the order 5 > 3 > 1 (Table 3), being much lower than those of [Pd2((C,N)L)2(m-sac)2] [44] (Kb ¼ 1.05  105 M
1) and [Pd(sac)(terpy)](sac)$4H2O (Kb ¼ 1.0 (±0.05)  105 M
1 [45]. Competitive binding between ethidium bromide (EB) and the platinum(II) complexes was carried out to further clarify the DNA binding mode of the complexes. It is well known that EB emits intense fluorescence because of its strong intercalation between the adjacent base pairs of DNA and the strong fluorescence may be quenched by adding other competitive molecules. Since EB intercalates DNA in the minor groove, its displacement by a compound is suggestive of an intercalative or minor groove binding. The effect of the complexes on the FS-DNAeEB system was followed by changes in the fluorescence intensity of EB. As shown in Fig. 2 and Fig. S8, the fluorescence intensity of the DNAeEB system is gradually reduced with increasing amounts of the complexes, due to the displacement some EB molecules from the DNAeEB complex. The binding constants such as KSV, Kapp and KF obtained from the analysis of the quenching data (Table 3) indicate that the complexes successfully compete with EB with a binding affinity of 1 > 3 > 5, supporting intercalative or groove binding mode of the

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Fig. 2. (A) UV spectra of FS-DNA solutions (50 mM) with increasing amounts of 3 (0e15 mM) of in Tris-HCl buffer. Inset: plot of 1/[complex] vs. 1/(AeA0). (b) Fluorescence emission spectra of EB-bound DNA solutions in the absence and presence of increasing concentrations of 3 (0e50 mM) in Tris-HCl. The Inset shows the SternVolmer plot of the fluorescence data.

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complexes. In addition, the value of n is about 1, suggesting the formation of a stoichiometric adduct between the complexes and DNA. The binding of the free phosphine compounds to FS-DNA was also studied by UV and fluorescence spectroscopy (Figs. S6 and S7). The binding parameters listed in Table 3 indicate that the binding propensity of the ligands varies in the following order: PPh3 > PPh2Cy > PPhCy2 > Pcy3, depending of the number of the Ph groups in the ligands, and the free ligands show remarkably low affinity towards DNA compared to their platinum complexes. Corresponding thermodynamic parameters of DNA binding are listed in Table S3. The quenching constant values decrease with increasing temperature, suggesting a static quenching mechanism. Moreover, the negative signs of both DH and DS are associated with van der Waals forces and hydrogen bonding interactions between the complexes and DNA [46]. Intercalation and groove binding modes can be distinguished by employing hydrodynamic methods. The measurement of viscosity of DNA solutions was accepted as the most sensitive method to determine the binding mode of drugs, since a classical intercalator causes an increase in the viscosity of DNA solutions, while a groove binder does not have an apparent change in the viscosity [47]. As shown in Fig. S9, the viscosity of DNA solutions do not change practically in the presence of increasing amounts of 1, 3 and 5, affirming that the complexes interact with FS-DNA through groove or non-intercalative mode of binding, similar to Hoechst, a DNA groove binder. Heating of a DNA solution induces conformational changes, especially separation of the double strands, called DNA denaturation (or DNA melting). Owing to the low transition probability of the p electrons of the DNA bases, the UV absorbance of a doublestranded DNA is less than that for a single stranded DNA and the melting point is measured at 260 nm as a function of temperature of the DNA solutions. Complexes 1, 3 and 5 at a complex/DNA ratio of 0.25 cause significant increases in the melting point of DNA by 3.5, 14.5 and 12.5, respectively (Fig. S10). The relatively large DT values are clearly indicative of the increased stability of the DNA double helix in the presence of the platinum(II) complexes, due to their strong interactions with DNA. Computational docking is extremely useful tool to understand interactions between drugs and biological targets. Among the most cytotoxic complexes, the crystal data of 5 was obtained. Therefore, only this complex was subjected to molecular docking studies to

Table 3 Values of binding constants for the interaction of the potent complexes with DNA and HSA.

DNA binding 1 3 5 PPh3 PPh2Cy PPhCy2 PCy3 HSA binding 1 3 5 PPh3 PPh2Cy PPhCy2 PCy3

Kb  10
4 (M
1)

KSV  10
4 (M
1)

Kapp  10
6 (M
1)

KF  10
5 (M
1)

n

0.8 ± 0.1 2.1 ± 0.2 7.6 ± 0.3 0.9 ± 0.2 0.8 ± 0.1 0.7 ± 0.2 0.4 ± 0.1

6.5 ± 0.2 5.4 ± 0.1 3.4 ± 0.1 1.1 ± 0.1 0.8 ± 0.1 0.7 ± 0.2 0.4 ± 0.1

1.7 1.3 1.1 e e e e

13.3 ± 0.1 10.8 ± 0.1 1.6 ± 0.1 0.11 ± 0.01 0.07 ± 0.02 0.07 ± 0.02 0.05 ± 0.01

1.2 1.2 1.1 0.9 1.0 1.1 1.2

2.7 ± 0.2 2.2 ± 0.2 2.1 ± 0.1 1.2 ± 0.3 1.0 ± 0.2 0.5 ± 0.2 0.4 ± 0.1

6.9 ± 0.2 5.8 ± 0.2 2.1 ± 0.2 0.6 ± 0.2 0.6 ± 0.2 0.5 ± 0.2 0.4 ± 0.1

e e e e e e e

8.4 ± 0.1 5.6 ± 0.1 2.8 ± 0.1 0.04 ± 0.002 0.05 ± 0.002 0.04 ± 0.00 0.05 ± 0.002

1.2 1.1 1.2 0.9 1.0 1.0 1.1

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identify the potential binding mode and energy as well as intermolecular interactions between the complex and two B-DNA structures, GC-rich 1QC1 (CCGCCGGCGG) and AT-rich 1DN9 (CGCATATATGCG). Molecular docking studies reveal that in both cases, the docked complex fits into the major groove of DNA comfortably, involving hydrogen bonding and relatively long-range hydrophobic interactions with the DNA bases (Fig. S11). One of the sulfonyl oxygens of sac interacts with the NH2 groups of cytosine in 1QC1 and adenine in 1DN9, forming strong hydrogen bonds with the bond lengths of 2.13 and 2.36 Å, respectively. The relative binding energies of the docked structures are the similar and calculated as
26.36 and
26.78 kJ mol
1, correlating well with the binding energy (
27.91 kJ mol
1) obtained experimentally from the fluorescence studies. The DNA docking models indicate nonselectivity of the platinum (II) complex between the GC- and ATrich sequences.

2.7. Gel electrophoresis To assess DNA binding affinity of 1, 3 and 5, a gel electrophoresis assay of pBR322 plasmid DNA incubated with increasing concentrations of the complexes for 4 h at 37  C was performed by monitoring the presence of different DNA forms. pBR322 plasmid DNA seems to be a mixture of mainly of supercoiled form I and a small amount of open circular form II. The interaction of the Pt(II) complexes with DNA is shown to induce conformational changes in plasmid DNA in a concentration-dependent manner (Fig. 3a). At 50 mM, the complexes do not exhibit any nuclease activity, but at 100 and 250 mM, DNA cleavage is apparent giving rise to the conversion of form I to form II and form III (linear form) with a similar pattern for all complexes. About 50% of the supercoiled DNA is nicked to produce form II, when incubated with complex 1, while a large proportion (ca. 61 and 87%) of the DNA is linearized in the case of the 250 mM concentration of 3 and 5, respectively. The present results clearly show that complexes 1, 3 and 5 effectively cleave pBR322 plasmid DNA in the absence of added oxidizing or

Fig. 3. (A) Cleavage of pBR322 plasmid DNA (100 ng) by the cytotoxic complexes 1, 3 and 5 after incubation for 4 h at 37  C. (b) Effect of groove binders DAPI and MG (100 mM) on the cleavage of plasmid DNA with and without of 1, 3 and 5 (100 mM). (c) Effect of 1, 3 and 5 (50 mM) on the cleavage of the plasmid DNA digested with BamHI and HindIII enzymes.

reducing agent and this efficient DNA cleavage is very promising for candidate anticancer agents. To evaluate the groove binding preferences of the complexes towards DNA, the nuclease activity of the Pt(II) complexes was studied in the presence of a minor groove binder, 4,6- diamidino-2phenylindole (DAPI), and a major groove binder, methyl green (MG). The groove binders reveal different behaviour on the nuclease activity of the plasmid DNA incubated with 1, 3 and 5 (Fig. 3b). The addition of DAPI has no effect on the inhibition of DNA cleavage activity of the complexes, whereas the nuclease activity of the complexes is completely inhibited by the presence of MG, suggesting that these Pt(II) complexes act as DNA major groove binders, in agreement with the results obtained in the DNA binding studies. The specificity of 1, 3 and 5 towards the base sequence of DNA was assessed by the enzyme inhibition assay using two restriction endonucleases with different recognition sites, namely BamHI (GYGATCC) and HindIII (AYAGCTT). As shown in Fig. 3c, approximately 90% of the supercoiled DNA is linearized by both enzymes. The endonuclease action of HindIII is considerably inhibited by 1 and 5, and partially inhibited by 3. All three complexes seem to be ineffective to inhibit the activity of BamHI. These results obviously suggest the binding affinity of the complexes towards the AT-rich site at the major grooves. 2.8. HSA binding studies Human serum albumin (HSA) is the most abundant plasma protein, capable of delivering drugs to the target via binding. Therefore, a study of the plasma protein was carried out for the determination of binding properties of the new platinum complexes, using UV absorption and fluorescence emission methods. As illustrated in Fig. S12, HSA produces a medium UV absorption band at 278 nm, assigned to the p e p* transitions of the aromatic amino acids. The incremental addition of the potent complexes results in gradual increase in the absorbance of the HSA solutions, indicating binding of the complexes with a static quenching mechanism. The binding constants (Kb) remain on the same order of magnitude for all of the complexes, being approximately 2.4 (±0.3)  104 M
1 (Table 3). The complexes seem to have moderate binding values compared to those of warfarin, and phenylbutazone drugs known to selectively bind to HSA with high affinity (Kb ¼ 3.4  105 and 7  105 M
1, respectively) [48]. On the other hand, it was emphasized that a high binding affinity towards HSA limits the transport of the drug and decreases the in vivo anticancer activity compared to the in vitro activity [49], suggesting that low HSA binding appears to be advantageous. Thus, HSA may be considered as a good carrier for transfer of these Pt(II) complexes. Intrinsic protein fluorescence is mainly attributed to the tryptophan (Trp), tyrosine (Tyr) and phenylalanine (Phe) residues in HSA. As shown in Fig. 4 and Fig. S13, the fluorescence emission of HSA is quenched regularly with the increasing concentration of these complexes. All of the quenching (KSV) and fluorescence constant (KF) values given in Table 3 follow the order: 1 > 3 > 5. Contrary to the complexes, the free phosphine ligands displayed very low affinity for HSA as deduced from the binding data given in Table 3, Figs. S11 and S12. Negative DH and DS values obtained from temperature-dependent emission measurements correspond to both hydrogen bonding and van der Waals interactions between the complexes and HSA (Table S4). The synchronous fluorescence spectra in (Fig. S14) indicate that the microenvironment polarities of the Trp and Tyr residues are not influenced by the binding of the platinum complexes. In addition, 3D fluorescence spectra of HSA in the presence of the platinum complexes clearly show that peaks 2 and 3 corresponding to the

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cells were treated with IC90 concentrations of complexes 1, 3, 5 and cisplatin for 12 and 24 h. 2.9.1. M30 assay Keratin 18 (K18) is a cytoskeleton protein of epithelial cells and is cleaved by caspases, forming a fragment termed caspase-cleaved cytokeratin 18 (ccK18 or M30 antigen) during the early phases of apoptosis. The ccK18 is recognized by a specific monoclonal antibody (M30). So, the detection of ccK18 fragments is used to measure the cell death response of cells against drugs [50,51]. Apoptosis caused by complexes 1, 3 and 5 was evaluated by measuring the level of ccK18 in the cells studied. Compared to the control cells, dramatic increases of the total ccK18 are observed in all the cancer cell lines after exposure of the platinum complexes (Fig. 5. The effect of the complexes on the formation of ccK18 seems to be similar for all the cell lines. In MCF-7 and DU145 cells, the complexes cause approximately two-fold increase in the ccK18 levels with respect to cisplatin. However, the ccK18 content of A549 cells treated with cisplatin is twice of those of the platinum complexes. The present results clearly indicate the induction of apoptosis by the present complexes.

Fig. 4. Emission and 3D fluorescence spectra of HSA in the absence and presence of 3. Inset: Stern-Volmer plot of the fluorescence data.

two characteristic fluorescence emissions of HSA are dramatically quenched by the addition of the three complexes, suggesting that binding of the complexes causes a significant change in the conformation of HSA, inducing some unfolding in the polypeptide chain (Fig. 4 and Fig. S15). To analyze binding specificity and intermolecular interactions, complex 5 was docked into the crystal structure of HSA. The docking results reveals that the complex is located within IIA subdomain of HSA, involving in the interactions with the amino acid residues Glu153, Lys195, Trp214, Arg218, Arg222, His242, Arg257 and Ala291 within a distance range of 4 Å (Fig. S16). The complex forms two hydrogen bonds with the Arg218 and Arg222 residues (2.24 and 2.54 Å, respectively). In addition, hydrophobic interactions also present between the complex, and Lys195, Trp214, His242, and Ala291 residues.

2.9.2. Detection of apoptosis via Annexin V and Caspase 3/7 activity assays Viable, early apoptotic, late apoptotic and necrotic cells can be differentiated by Annexin V assay through detection of phosphatidylserine [52], while caspases are most often associated with apoptosis [53,54]. Therefore, in addition to the M30 assay, the apoptosis inducing ability of 1, 3 and 5 in MCF-7 and HCT116 cells was studied by flow cytometry through Annexin V and Caspase 3/7 assays, compared to control, non-treated cells and to the reference drug cisplatin (Fig. 6, S17 and S18). These cells treated with 1, 3, 5 and cisplatin for 12 and 24 h show a reduction in the proportion of living cells over time, with a concomitant increase in cells undergoing apoptosis in early and late stages. In most cases, the percentage of death cells are found higher than those treated with cisplatin. Notably, all complexes are very effective against HCT116 cells, compared to MCF-7 cells, owing to a significant increase in the population of cells in the early apoptosis phase at 12 h. Annexin V positivity and Caspase 3/7 activity results at 24 h showed that all cells exposed to the complexes undergo apoptotic cell death with the apoptosis inducing ability of 1 and 3 being much higher than 5 and cisplatin. 2.9.3. Generation of reactive oxygen species Another important factor in apoptosis is the excessive generation of ROS, which causes oxidative stress resulting in

2.9. Mechanism of action The mechanism of action of the most cytotoxic complexes (1, 3 and 5) was clarified by the means of M30 Elisa and flow cytometry assays, including Annexin V, Caspase-3/7 activity, mitochondrial membrane potential, oxidative stress and DNA damage assays. The

Fig. 5. CcK18 levels obtained after 48 h treatment of the cells with the IC90 values of the platinum complexes.

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Fig. 6. Flow cytometric quantification of Annexin V/7-AAD positivity (a) and caspase3/7 activity (b) in MCF-7 cells treated with the IC90 values of 3 and cisplatin for 12 and 24 h.

macromolecular damage and eventually leads to cell death [55]. Intracellular ROS levels in MCF-7 and HCT116 cells incubated with 1, 3, 5 and cisplatin for 12 and 24 h were determined by flow cytometry (Fig. 7a and Fig. S19). Exposure of the complexes

obviously promotes the production of ROS in MCF-7 and HCT116 cell lines with time. The percentage of stressed cells (M2) increases, due to a remarkable increase in intracellular ROS levels. In comparison with cisplatin, all complexes induce the higher

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membrane permeability and damages the respiratory chain resulting in increased ROS production [56,57]. The disruption in the mitochondrial membrane may trigger the release of cytochrome c from the mitochondria, initiating events leading to apoptosis. It has been proved that mitochondria are potential targets for platinum chemotherapeutics toxic to cancer cells [58,59] and therefore, targeting them becomes a novel strategy for cancer therapy [60]. As illustrated in Fig. 7b and Fig. S20, in response to the treatment with complexes 1, 3 and 5, both MCF-7 and HCT116 cells are found to demonstrate time dependent increases in the mitochondrial depolarization, reflected by increases in percentage of depolarized cells (depolarized-live þ depolarized-dead cells). The loss of mitochondrial membrane potential in both cell lines are considerably higher in the case of the platinum complexes compared to cisplatin. These findings provide the direct evidence that the new complexes specifically target mitochondria by collapsing of mitochondrial membrane potential that leads to apoptosis. The mitochondrial dysfunction in the cells correlates well with the highly increased ROS production induced by the platinum complexes. 2.9.5. Damage to double-strand DNA The binding studies and the plasmid assay provide preliminary information on how the potent complexes (1, 3 and 5) interact with DNA. To evaluate formation of double-strand DNA breaks (DSBs) in living cells, both MCF-7 and HCT116 cells were treated with the IC90 concentrations of complexes 1, 3, 5 and cisplatin for 12 and 24 h, and DNA damage was determined by measuring the levels of phosphorylated H2AX (ɣH2AX), a well-established DNA damage marker [61]. All complexes activated H2AX phosphorylation in both cell lines, indicative of DNA damage (Figs. 7c and Figs. S21). The percentage of ɣH2AX expression in MCF-7 cells treated with 3 for 12 h is similar to those exposed to cisplatin, while much higher ɣH2AX levels were recorded in HCT116 cells incubated with the complexes. These results clearly demonstrate that the Pt (II) complexes act as double-strand DNA damaging agents. 3. Conclusions

Fig. 7. Intracellular ROS levels (a), changes in mitochondrial membrane potential (b), and double-strand DNA damage (c) in MCF-7 cells incubated with the IC90 values of 3 and cisplatin for 12 and 24 h.

elevation of ROS in HCT116 cells at 12 and 24 h, while 3 is more prominent for the production of ROS in MCF-7 cells.

2.9.4. Depolarization of mitochondrial membrane potential The mitochondria are a source of ROS, and also a target of excessive generation of ROS and therefore, play a crucial role in cell growth and apoptosis. Excess ROS increases the mitochondrial

Novel platinum(II) saccharin complexes containing the tertiary phosphine ligands following progressive phenyl replacement by cyclohexyl were prepared using different synthetic strategies and characterized by spectroscopic techniques and X-ray crystallography. Most of the complexes showed trans configuration. In vitro anticancer activity screening via SRB viability assay showed that among the platinum complexes, only trans-configured 1, 3 and 5 displayed promising biological activity compared to cisplatin. The complexes are mainly localized in cytoplasm and the highest cytotoxicity of 3 can be correlated with its lipophilicity and cellular uptake. All complexes exhibit DNA cleavage activity towards plasmid DNA. Binding measurements as well as computational docking data suggest that the potent complexes interact with DNA, presumably by groove-binding mechanism, and HSA through hydrophobic IIA subdomain. The apoptotic response of MCF-7 and HCT116 cells to the new platinum complexes and cisplatin was examined by M30 assay and quantified by the activity of Annexin V and caspases 3/7. Furthermore, generation of ROS, depolarization of mitochondrial membrane potential and DNA damage induced by the complexes were analyzed by flow cytometry. In summary, these results suggest that the antiproliferative mechanisms of action of the platinum complexes involve activation of apoptotic pathways through externalization of phosphatidylserine, activation of caspases, excessive production of ROS, mitochondrial dysfunction and genomic DNA damage. Since mitochondria- and DNA-targeted complexes are promising agents for chemotherapy, the trans-configured platinum

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complexes (1, 3 and 5) can be considered as potential anticancer agents acting especially though the mitochondrial and DNA damage. These results will further be helpful for design and synthesis of new Pt(II) complexes as potent anticancer drugs. 4. Experimental 4.1. Materials and measurements All commercially available chemicals were used without further purification. Microanalyses were performed using a Costech elemental analyzer. UV-vis spectra were measured on a Perkin Elmer Lambda 35 spectrophotometer. IR spectra were recorded on a Perkin Elmer Spectrum Two FT-IR spectrophotometer. 1H, 13C and 31 P NMR spectra were recorded on a Bruker spectrometer at 400, 100 and 162 MHz, respectively in DMSO‑d6 at room temperature (rt). Me4Si was used as an internal reference for 1H and 13C NMR, while OP(Oph)3 in acetone-d6 (0.0485 M) was used as a reference for 31P NMR. Fluorescence spectra were recorded at room temperature with a Varian Cary Eclipse spectrophotometer equipped with a Xe pulse lamp of 75 kW. For all fluorescence measurements, the slits were maintained at 5 nm. The ESI mass spectra were recorded using a Bruker Daltonics Microtof II-ESI-TOF mass spectrometer. The electrical conductivity measurements of the complexes in MeOH were carried out with HANNA HI 5521 at rt. Melting points are measured using a BUCHI 560 instrument. 4.2. Synthesis of platinum(II) complexes Following procedures were used for the synthesis of complexes 1e7. Each of PPh2Cy, PphCy2 and Pcy3 (0.5 mmol) dissolved in MeOH or MeCN (10 mL) was added to a solution of [PtCl (sac) (COD)] (0.25 mmol) in MeCN (10 mL). After a day of reflux, the solvents were removed to yield complexes 2, 4 and 6. Similarly, complexes 1, 3, 5 and 7 were obtained in the same way replacing [PtCl (sac) (COD)] with [Pt (sac)2(COD)]. Crystallization from MeOH, MeCN, DMF and DMSO yielded the single crystals of 2, 5, 6 and 7. trans-[Pt(sac)2(PPh3)2] (1). Yield: 209 mg (77%). Mp: 211e218  C (decomp.). Anal. Calc. for C50H38N2O6P2PtS2 (%): C, 55.4; H, 3.5; N, 2.6. Found C, 55.2; H, 3.5; N, 2.8. IR (n/cm
1): 3058w (CH), 1678s (CO-sac), 1592w, 1481 m, 1435s, 1310s, 1285 m, 1247s nas (SO2), 1171 m, 1156s ns (SO2), 1090s, 999w (PeCsym), 952s nas (CNS), 748vs, 694vs, 676vs, 590s, 549vs, 527vs, 512vs (PeCasym), 497vs. 1H-NMR (400 MHz, DMSO‑d6, d, ppm): 7.96e6.94 (m, 30H, Ph protons and 8H-sac). 13C NMR (100 MHz, DMSO‑d6, d, ppm): 165.3 (C7-sac), 142.4 (C1-sac), 134.4 (t, Cipso-Ph, JPeC ¼ 7 and 7 Hz), 132.5 (C3-sac), 131.7 (C6-sac), 131.3 (Cortho-Ph), 129.2 (C4-sac), 128.8 (t, Cpara-Ph JPeC ¼ 5 and 5 Hz), 128.4 (d, Cmetha-Ph, JPeC ¼ 11 Hz), 122.9 (C5-sac), 119.7 (C2-sac). 31P NMR (DMSO‑d6, 162 MHz): d 15.17 (s, 1 JPteP ¼ 2138 Hz, 2JPP ¼ 22.7 Hz). UV-vis (MeOH) lmax/nm (3max/ dm3mol
1cm
1) 275 (34 197). Molar conductivity, LM (MeOH, 298 K, 10
3 M) 8 S cm2 mole1 (nonelectrolyte). ESIeMS (m/z, MeOH): 1107.6 (17%, calc. 1107.0) [M þ Na]þ, 830.0 (44%, calc. 830.8) [Pt (PPh3)2 (MeOH)7/2 e H]þ, 718.9 (100%, calc. 718.6) [Pt (PPh3)2 e H]þ. cis-[PtCl(sac)(PPh2Cy)2] (2). Yield: 202 mg (85%). Mp: 170e172  C. Anal. Calc. for C43H46ClNO3P2PtS (%): C, 54.4; H, 4.9; N, 1.5. Found C, 54.6; H, 4.7; N, 1.7. IR (n/cm
1): 3063w, 2922 m, 2851 m (CH), 1673s (CO-sac), 1598w, 1484w, 1434 m, 1321w ns (CNS), 1306s, 1296 m, 1243 m nas (SO2), 1154s, 1145 m ns (SO2), 1120 m, 1100 m, 1002 m (PeCsym), 946s nas (CNS), 791 m, 749s, 691vs, 677s, 594vs, 562 m, 548s, 532vs, 517vs (PeCasym), 486s. 1HNMR (400 MHz, DMSO‑d6, d, ppm): 7.99e6.96 (m, 20H, Ph protons and 4H-sac), 2.25e0.48 (m, 22H, Cy protons). 13C NMR (100 MHz, DMSO‑d6, d, ppm): 162.9 (C7-sac), 143.0 (C1-sac), 134.1 (m, Cipso-Ph),

133.6 (C3-sac), 132.0 (C6-sac), 131.6 (d, Cortho-Ph, JPeC ¼ 7 Hz), 131.4 (C4-sac), 129.2 (d, Cpara-Ph, JPeC ¼ 8 Hz), 128.7 (d, Cmetha-Ph, JPeC ¼ 9 Hz), 123.6 (C5-sac), 120.4 (C2-sac), 30.3 (Cipso-Cy), 29.5 (Cortho-Cy), 26.9 (d, Cmetha-Cy, JPeC ¼ 7 Hz), 26.1 (Cpara-Cy). 31P NMR (DMSO‑d6, 162 MHz): d 12.68 (s, 1JPteP ¼ 3535 Hz; 2JPP ¼ 14.6 and 14.6 Hz). UV-vis (MeOH) lmax/nm (3max/dm3mol
1cm
1) 275 (22 777). Molar conductivity, LM (MeOH, 298 K, 10
3 M) 4 S cm2 mole1 (nonelectrolyte). ESIeMS (m/z, MeOH): 972.0 (21%, calc. 972.4) [M þ Na]þ, 913.4 (40%, calc. 913.9) [M
Cl]þ, 767.4 (100%, calc. 767.2) [M
sac]þ, 731.3 (70%, calc. 730.8) [Pt (PPh2Cy)2 e H]þ. trans-[Pt(sac)2(PPh2Cy)2] (3). Yield: 181 mg (66%). Mp: 160e162  C. Anal. Calc. for C50H50N2O6P2PtS2 (%): C, 54.8; H, 4.6; N, 2.6. Found C, 55.0; H, 4.8; N, 2.4. IR (n/cm
1): 3054w, 2927 m, 2852 m (CH), 1664 m (CO-sac), 1591w, 1484w, 1435s, 1334w ns (CNS), 1291s, 1247 m nas (SO2), 1169 m, 1152s ns (SO2), 1100 m, 1000w (PeCsym), 963 m nas (CNS), 848w, 747s, 695vs, 678s, 595s, 523vs, 513vs (PeCasym). 1H-NMR (400 MHz, DMSO‑d6, d, ppm): 8.06e6.97 (m, 20H, Ph protons and 8H-sac), 1.88e0.61 (m, 22H, Cy protons). 13C NMR (100 MHz, DMSO‑d6, d, ppm): 165.5 (C7-sac), 142.9 (C1-sac), 134.0 (t, Cipso-Ph, JPeC ¼ 6 Hz), 133.4 (C3-sac), 132.7 (d, Cortho-Ph, JPeC ¼ 13 Hz), 132.3 (d, Cpara-Ph, JPeC ¼ 9 Hz), 131.9 (C6sac), 130.5 (C4-sac), 128.4 (t, Cmetha-Ph, JPeC ¼ 5 and 5 Hz), 122.9 (C5sac), 119.7 (C2-sac), 36.3 (d, Cipso-Cy, JPeC ¼ 15 Hz), 36.1 (Cortho-Cy), 29.3 (Cpara-Cy), 26.4 (t, Cmetha-Cy, JPeC ¼ 16 and 16 Hz). 31P NMR (DMSO‑d6, 162 MHz): d 34.07 (s, 1JPteP ¼ 2409 Hz). UV-vis (MeOH) lmax/nm (3max/dm3mol
1cm
1) 245 (63 400), 274 (32 906). Molar conductivity, LM (MeOH, 298 K, 10
3 M) 9 S cm2 mol-1 (nonelectrolyte). ESIeMS (m/z, MeOH): 870.7 (100%, calc. 869.8) [Pt (sac)2(PPh2Cy) (MeCN) þ H]þ, 731.5 (76%, calc. 730.8) [Pt (PPh2Cy)2 e H]þ. trans-[PtCl(sac)(PPhCy2)2] (4). Yield: 82 mg (34%). Mp: 160e164  C. Anal. Calc. for C43H58ClNO3P2PtS (%): C, 53.7; H, 6.1; N, 1.5. Found C, 53.4; H, 6.4; N, 1.7. IR (n/cm
1): 3067w, 3049w, 2925vs, 2850s (CH), 1663 m (CO-sac), 1591w, 1482w, 1448 m, 1435s, 1334w ns (CNS), 1293s, 1247 m nas (SO2), 1169s, 1153s ns (SO2), 1119 m, 1100 m, 1003 m (PeCsym), 959 m nas (CNS), 851 m, 746vs, 696vs, 679 m, 596vs, 555 m, 535vs, 518vs, 502vs (PeCasym), 477s. 1H-NMR (400 MHz, DMSO‑d6, d, ppm): 8.17e7.04 (m, 10H, Ph protons and 4H-sac), 1.85e0.64 (m, 44H, Cy protons). 13C NMR (100 MHz, DMSO‑d6, d, ppm): 166.5 (C7-sac), 143.4 (C1-sac), 135.2 (t, Cipso-Ph, JPeC ¼ 6 and 6 Hz), 134.7 (d, Cortho-Ph, JPeC ¼ 3 Hz), 133.5 (C3-sac), 133.3 (C6-sac), 132.9 (Cpara-Ph), 130.8 (C4-sac), 128.1 (Cmetha-Ph), 123.1 (C5-sac), 121.7 (C2-sac), 31.8(t, Cipso-Cy, JPeC ¼ 15 and 15 Hz), 28.6 (Cortho-Cy), 28.1 (Cpara-Cy), 26.5 (t, Cmetha-Cy, JPeC ¼ 16 Hz). 31P NMR (DMSO‑d6, 162 MHz): d 44.15 (s, 1JPteP ¼ 2174 Hz, 2 JPP ¼ 97.2 Hz). UV-vis (MeOH) lmax/nm (3max/dm3mol
1cm
1) 264 (21 489), 272 (16 823). Molar conductivity, LM (MeOH, 298 K, 10
3 M) 3 S cm2 mole1 (nonelectrolyte). ESIeMS (m/z, MeOH): 743.0 (100%, calc. 742.8) [Pt (PPhCy2)2 e H]þ. trans-[Pt(H)(sac)(PPhCy2)2] (5). Yield: 209 mg (90%). Mp: 160e163  C. Anal. Calc. for C43H59NO3P2PtS (%): C, 55.7; H, 6.4; N, 1.5. Found C, 55.4; H, 6.6; N, 1.8. IR (n/cm
1): 3058w, 2928vs, 2854s (C-H), 2234 m (PteH), 1663vs (CO-sac), 1594 m, 1447s, 1436vs, 1325 m ns (CNS), 1285vs, 1250s nas (SO2), 1151vs ns (SO2), 1119s, 1107, 1058vs, 1004s (PeCsym), 960vs nas (CNS), 851 m, 747vs, 695vs, 680s, 596vs, 561vs, 535vs, 518vs, 506vs (PeCasym), 488s. 1H-NMR (400 MHz, DMSO‑d6, d, ppm):
17.81 (dt, H, 2JHeP ¼ 12 and 12 Hz), 8.02e7.37 (m, 10H, Ph protons and 4H-sac), 2.16e0.44 (m, 44H, Cy protons). 13C NMR (100 MHz, DMSO‑d6, d, ppm): 162.8 (C7-sac), 141.8 (C1-sac), 135.1 (C3-sac), 134.6 (Cipso-Ph), 131.8 (d, Cortho-Ph, JPeC ¼ 7 Hz), 131.6 (C6-sac), 130.8 (m, Cpara-Ph), 128.7 (d, Cmetha-Ph, JPeC ¼ 10 Hz), 128.2 (C4-sac), 124.8 (C5-sac), 121.2 (C2-sac), 26.5 (m, Cipso-Cy), 26.2 (d, Cortho-Cy, JPeC ¼ 11 Hz), 25.8 (Cmetha-Cy), 24.8 (Cpara-Cy). 31P NMR (DMSO‑d6, 162 MHz): d 44.18 (s,

V.T. Yilmaz et al. / European Journal of Medicinal Chemistry 155 (2018) 609e622 1 JPteP ¼ 2430 Hz, 2JPP ¼ 92.3 Hz). UV-vis (MeOH) lmax/nm (3max/ dm3mol
1cm
1) 265 (18 409), 272 (15 521). Molar conductivity, LM (MeOH, 298 K, 10
3 M) 5 S cm2 mole1 (nonelectrolyte). ESIeMS (m/ z, MeOH): 744.9 (100%, calc. 744.8) [M
sac]þ, 313.6 (23%, calc. 313.5) [PPhCy2 þ K]þ. trans-[PtCl(sac)(PCy3)2] (6). Yield: 73 mg (30%). Mp: 210e212  C (decomp.). Anal. Calc. for C43H70ClNO3P2PtS (%): C, 52.9; H, 7.4; N, 1.4. Found C, 52.9; H, 7.5; N, 1.5. IR (n/cm
1): 2918vs, 2850s (CH), 1683s (CO-sac), 1596w, 1444 m, 1308s, 1296s, 1246s nas (SO2), 1172vs, 1154s ns (SO2), 1142 m, 1127 m, 1003 m (PeCsym), 950 m nas (CNS), 888 m, 847 m, 752s, 735 m, 678s, 595vs, 561s, 536s, 512vs (PeCasym), 491s. 1H-NMR (400 MHz, DMSO‑d6, d, ppm): 7.94e7.74 (m, 4H-sac), 1.85e1.57 (m, 40H, Cy protons), 1.31e1.15 (m, 26H, Cy protons). 13C NMR (100 MHz, DMSO‑d6, d, ppm): 160.4 (C7-sac), 141.5 (C1-sac), 133.8 (C3-sac), 131.3 (C6-sac), 129.1 (C4-sac), 123.8 (C5-sac), 120.7 (C2-sac), 35.3 (Cipso-Cy), 34.7 (Cortho-Cy), 26.8 (d, Cmetha-Cy, JPeC ¼ 12 Hz), 26.3 (d, Cpara-Cy, JPeC ¼ 9 Hz). 31P NMR (DMSO‑d6, 162 MHz): d 48.54 (s, 1JPteP ¼ 2430 Hz). UV-vis (MeOH) lmax/nm (3max/dm3mol
1cm
1) 222 (55 640), 265 (18 731). Molar conductivity, LM (MeOH, 298 K, 10
3 M) 4 S cm2 mole1 (nonelectrolyte). ESIeMS (m/z, MeOH): 974.8 (15%, calc. 974.6) [M þ H]þ, 809.0 (100%, calc. 809.4) [PtCl(PCy3)2 (MeOH)1/2]þ, 754.9 (94%, calc. 754.9) [Pt (PCy3)2 e H]þ, 515.5 (72%, calc. 515.5) [Pt (PCy3) (MeCN) e H]þ. Trans-[Pt(H)(sac)(PCy3)2] (7). Yield: 197 mg (84%). Mp: 200e201  C (decomp.). Anal. Calc. for C43H71NO3P2PtS (%): C, 54.9; H, 7.8; N, 1.5. Found C, 54.8; H, 7.7; N, 1.6. IR (n/cm
1): 2922vs, 2849s (CH), 2217 m (PteH), 1662s (CO-sac), 1596w, 1446 m, 1334w, 1321w ns (CNS), 1298vs, 1283 m, 1246 m nas (SO2), 1171s, 1154vs, 1142 m ns (SO2), 1005 m (PeCsym), 955 m nas (CNS), 848 m, 783 m, 752s, 679 m, 594vs, 553 m, 540s, 521s, 511vs (PeCasym), 491 m. 1H-NMR (400 MHz, DMSO‑d6, d, ppm):
18.18 (t, H, 2JHeP ¼ 12 and 12 Hz), 7.80e7.73 (m, 4H-sac), 2.40e0.78 (m, 66H, Cy protons). 13C NMR (100 MHz, DMSO‑d6, d, ppm): 166.2 (C7-sac), 143.5 (C1-sac), 133.6 (C3-sac), 132.8 (C6-sac), 132.6 (C4-sac), 123.5 (C5-sac), 120.1 (C2-sac), 35.3 (s, Cipso-Cy), 34.7 (s, Cortho-Cy), 26.8 (d, Cmetha-Cy, JPeC ¼ 11 Hz), 26.4 (d, Cpara-Cy, JPeC ¼ 9 Hz). 31P NMR (DMSO‑d6, 162 MHz): d 48.54 (s, 1JPteP ¼ 2754 Hz). UV-vis (MeOH) lmax/nm (3max/ dm3mol
1cm
1) 228 (67 460), 269 (16 721). Molar conductivity, LM (MeOH, 298 K, 10
3 M) 10 S cm2 mole1 (nonelectrolyte). ESIeMS (m/z, MeOH): 757.0 (47%, calc. 756.9) [M
sac]þ, 614.8 (100%, calc. 614.5) [Pt (sac)2 (MeOH) þ Na]þ, 582.8 (44%, calc. 582.4) [Pt (sac)2 þ Na]þ.

4.3. X-ray structure determination Single crystal X-ray diffraction data were collected on a Rigaku Xcalibur X-ray diffractometer with EOS CCD detector using Mo-Ka radiation (0.71073 Å) with u-scan mode. The data collection, cell refinement and data reduction were performed using CrysAlispro [62]. Using Olex2 [63], the structures were solved with the ShelXT structure solution program [64], using Intrinsic Phasing and refined with the ShelXL refinement package using Least Squares minimization [65]. All non-hydrogen atoms were refined anisotropically, while all hydrogen atoms were located at calculated positions and refined by using riding model. Crystal structures of 5 and 6 contain solvent accessible void volumes due to a disordered DMSO molecule in 5, and two disordered DMF molecules in 6. The solvent mask procedure employed by Olex2 were used to remove the electron densities. A number of DFIX/ISOR/SADI/RIGU restrains and equal Uij constraints (EADP) were applied to refine some moieties of the structures. Details of the data collection and structure refinement are given in Table S1. Crystallographic data and refinement parameters of the reported structures have been deposited at the Cambridge

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Crystallographic Data Centre with CCDC numbers 1828365
1828368. These data can be obtained free of charge via www.ccdc.cam.ac.uk/data_request/cif, or by emailing data_ [email protected], or by contacting The Cambridge Crystallographic Data Centre, 12 Union Road, Cambridge CB2 1EZ, UK; fax: þ44 1223 336033. 4.4. Partition coefficients Partition coefficients of 1, 3, 5, and cisplatin in n-octanol/water were determined by the classical shake flask method using the standard protocol [66]. Two parallel experiments were performed for each sample and the amount of the complexes in the n-octanol phase was analyzed by UV-vis spectrophotometry. Partition coefficient (log P) defined as the logarithmic ratio of the equilibrium concentrations of the dissolved complex between the organic and aqueous phases (log CO/CW) was estimated. 4.5. Cellular uptake MCF-7 cells were exposed to 25 mM of 1, 3, 5, and cisplatin at 37  C for 4 h. Then, the incubation medium was removed and the cells were washed with cold phosphate-buffered saline twice, tyripsinized and the cell suspension was counted. The FractionPREP kit (BioVision) was used to isolate subcellular fractions (cytosol, nucleus, membrane/particulate and cytoskeletal) according to the supplier's instructions. 2.5 mL of ultrapure HNO3 (69%) were added to the fractions and the solutions were heated to dryness. Then, the solids were dissolved in 1 mL of HCl (37%) and heated to dryness to remove traces of HNO3. The Pt content in each fraction was determined by the differential pulse stripping voltammetry, according to the method reported in the literature [67], using an Epsilon electrochemical workstation with a conventional three electrode system with hanging mercury drop working electrode, a glassy carbon auxiliary electrode and a Ag/AgCl reference electrode. Each measurement was repeated three times and the results were expressed as ng Pt/106 cells. 4.6. Biological activity tests 4.6.1. Cell lines Breast cancer (MCF-7), lung cancer (A549), prostate cancer (DU145), colon cancer (HCT116) and human normal bronchial epithelial cells (BEAS-2B) were cultured with RPMI 1640 culture media supplemented with penicillin G (100 U/mL), streptomycin (100 mg/mL), L-glutamine, and 10% fetal bovine serum at 37  C in a humidified atmosphere containing 5% CO2. For all compounds, 40 mM stock solutions were prepared by dissolving the compounds in DMSO. Further dilutions were prepared in cell culture medium. Final concentration of DMSO was maximum 0.1% in order to avoid its toxic effect on cells. 4.6.2. SRB cell cytotoxicity assay SRB cytotoxicity assay was preferred for viability screening of compounds as suggested by National Cancer Institute of America (NCI) [68]. In brief, cells were seeded in 96 well plate at the density of 5  103 cells/well and treated with 20 mM dose of all complexes for 48 h. At the end of the treatment period, cells were in situ fixed with 50 mL 50% (w/v) trichloroacetic acid (TCA) at 4  C for 1 h. After fixation step, wells were washed 5 times with deionized water and allowed air dry at room temperature. 0.4% SRB solution added 50 mL into each well and incubated at room temperature for 30 min. Then, plates were washed wit 1% acetic acid to eliminate non-specific bindings and unbounded dye. The bounded SRB dye was dissolved by adding 150 mL 10 mM Trisbase per each well.

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Measurement executed at 564 nm optical density (Lumistar Omega microplate reader, Germany). Cisplatin was used as a positive control. Experiments were performed two dependent experiments in triplicate. 4.6.3. ATP viability assay The ATP viability method, which based on the measurement of intracellular ATP content in cells, is highly sensitive and susceptible as it is based on luminescence. For this reason, it is more reliable than other colorimetric tests (such as MTT, MTS and XTT) used to determine cell viability. It also shows that this method has a good correlation between the number of viable cells and the relative light unit (RLU) readings even in low cell numbers (up to 20 cells), which is more suitable for cancer cell studies. In order to determine cell viability, MCF-7, A549, DU145, HCT116 and BEAS-2B cells were seeded 5  103 cells/well and treated for 48 h with complexes 1, 3, 5, which were found to be effective according to the SRB cytotoxicity test. Cisplatin was used as a positive control. The ATP content in the treated cells and control cells was measured using a luminometer (Lumistar Omega microplate reader, Germany) according to the principle of luciferin-luciferase bioluminescence reaction. Measuring time was 1 s. Results were obtained as Relative Light Unit (RLU) and the % viability of the samples was calculated according to the RLU values obtained from the control cells. Viability was calculated by using the formula below: Viability (%) ¼ [100  (Sample RLU) / (Control RLU)]

4.6.4. M30 (caspase-cleaved cytokeratin 18) Elisa method During cell death such as apoptosis and necrosis, cytokeratin 18 (CK18), an important protein of the cell skeleton, is extracellularly released. The M30 monoclonal antibody recognizes particularly the Asp396 fragment of CK18 (M30), which makes use of CKs as an apoptotic marker. Thus, caspase-cleaved CK18 (also known as M30), an apoptosis-specific marker, can be detected by ELISA. MCF7, A549, DU145, HCT116, and BEAS-2B cell lines were seeded 5  103 cells/well in 96-well cell culture plates in order to demonstrate the possible presence of apoptosis in the cells. Then treated with IC90 doses of complexes 1, 3, 5 and cisplatin which were calculated according to the ATP viability assay results. After 48 h of treatment, the cells were lysed using 10% NP-40. The supernatant was used after centrifugation at 2000 rpm for 10 s. The obtained samples were screened for the presence of apoptosis using the M30 CytoDeath ™ ELISA kit (Peviva, Sweden) according to manufacturer's instructions. 4.6.5. Cytofluorimetric analysis MCF-7 and HCT116 cell lines were plated at 3  105 cells per well (in dublicate) into 6-well plates and treated with IC90 concentrations of complexes 1, 3, 5 and cisplatin for 12 and 24 h. At the end of the treatment, cells were collected and analyzed for the detection of early/late apoptosis and cell death mode using Annexin V/Dead Cell kit (MCH100105, Millipore) and caspase 3/7 kit (MCH100108, Millipore) respectively, according to the manufacturer's instructions. The live, dead, early and late apoptotic cells were counted with the Muse Cell Analyzer (Millipore). To carry out the MitoPotential assay, MCF-7 and HCT116 cells exposed to IC90 concentrations of 1, 3, 5 and cisplatin for 12 and 24 h were washed with PBS, and collected by trypsinization and then were incubated with the Muse MitoPotential working solution (MCH100110, Millipore) for 20 min at 37  C. At the end of the incubation 7-AAD was added as a dead cell marker for an additional 5 min. Cells were analyzed by using the Muse Cell Analyzer

(Millipore). To evaluate the effect of IC90 concentration of complexes 1, 3, 5 and cisplatin on ROS generation in MCF-7 and HCT116 cells at 12 and 24 h, the assay was conducted on the Muse Cell Analyzer (Millipore) using the ROS kit (MCH100111-2, Millipore). Staining procedure was followed as per the manufacturer's protocol. To assess the double-strand DNA breaks, MCF-7 and HCT116 cells exposed to 1, 3, 5 and cisplatin at the IC90 concentrations for 12 h and 24 h were centrifuged at 300  g for 5 min, washed once with PBS and fixed with the Muse Fixation Buffer, which is a component of the Muse ɣH2AX Activation Dual Detection (kit MCH200101, Millipore) for 5 min on ice. After fixation, cells were permeabilized by ice-cold Muse Permeabilization Buffer and incubated on ice for 5 min. The cells were centrifuged (300  g, 5 min), resuspended in 45 mL 1  Assay Buffer, and incubated with the mixture of 2.5 mL of antiphospho-Histone H2AX and 2.5 mL of anti-Histone H2AX, PECy5 (30 min, dark, at rt). At the end of the incubation, cells were resuspended in 100 mL of 1  Assay Buffer, centrifuged (300  g, 5 min), and resuspended in 150 mL of 1  Assay Buffer. The data were acquired on the Muse Cell Analyzer (Millipore). 4.7. DNA binding DNA binding experiments were carried out in the Tris-HCl buffer (20 mM Tris-HCl/NaCl, pH ¼ 7.0). Absorption titration experiments of FS-DNA (50 mM) in Tris-HCl were performed by increasing the concentration of the most cytotoxic complexes 1, 3 and 5 (0e15 mM) dissolved in MeOH. The intrinsic binding constants (Kb) were was calculated according to the Benesi-Hildebrand equation [69]. 1/(AeA0) ¼ 1/{Kb(AmaxeA0)[Q]} þ 1/[AmaxeA0] where A0 is the absorption intensity of DNA, A is the absorption intensity of DNA interacted with a metal complex, Amax is the saturated absorption intensity of the DNAemetal complex adduct and [Q] is the concentration of the metal complex. The binding constant (Kb) was graphically evaluated by plotting 1/[AeA0] versus 1/[Q]. In fluorescence titration experiments, 1, 3 and 5 (0e50 mM) were added to FS-DNA solutions pre-treated with ethidium bromide (EB) (NP/EB ¼ 10) in Tris-HCl. The emission spectra of these solutions were recorded upon excitation at lex ¼ 295 nm, and 293, 297 and 300 K. The quenching constants (KSV) were determined using the Stern-Volmer equation [70]. F0/F ¼ 1 þ KSV[Q] where F0 and F are the fluorescence intensities in the absence and presence of the complexes, respectively. [Q] is the total concentration of the quencher (metal complex). The apparent binding constant (Kapp) was estimated from the following equation [71]. KEB[EB] ¼ Kapp[Q] in which [Q] is the concentration of the quencher causing a 50% reduction in the fluorescence intensity of EB-bound DNA, KEB ¼ 1.0  107 M
1. The binding constant KF is determined from the Scatchard equation [72]. Log(F0eF)/F ¼ logKF þ nlog[Q] The plot of log (F0eF)/F versus log [Q] is drawn and fitted linearly

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and the number of binding site (n) per nucleotide was obtained from the slope. In viscosity measurements, the FS-DNA solutions (0.8 mM bp) were mixed with increasing amounts of 1, 3 and 5, and the relative viscosity of these solutions were measured using an Ubbelohde viscometer at 20  C. Viscosity values were calculated from the observed flow time of DNA-containing solutions (t) corrected for the flow time of buffer alone (t0), h ¼ t e t0. DNA melting was performed using the UV/vis spectrophotometer equipped with a Peltier temperature programmer. The temperature of the FS-DNA solutions (100 mM) containing 1, 3 and 5 (25 mM) was monitored from 25 to 95  C. The melting temperature (Tm) were determined from the plot of relative absorbance (A/A25) at 260 nm versus temperature. 4.8. Gel electrophoresis assay The cleavage of pBR322 plasmid DNA by 1, 3 and 5 was performed agarose gel electrophoresis after incubation of the samples containing 100 ng plasmid DNA and MeOH solutions of the complexes (0e250 mM) in 50 mM Tris-HCl/NaCl buffer (pH 7.2) at 37  C for 4 h. In a separate experiment, the plasmid DNA was incubated with MG and DAPI (100 mM) for 1 h and then to the solution, 1, 3 and 5 (100 mM) were added. The final solutions were incubated another 3 h. The samples were electrophoresed for 45 min at 120 V on 1.0% agarose gel using 1X TBE buffer (pH 8.0). In restriction enzyme inhibition studies, the plasmid DNA was incubated with 50 mM of the complexes at 37  C in the buffer (pH 7.2) for 1 h. These solutions were subsequently incubated separately with HindIII and BamHI (2 units) for 15 min. The results of incubation were obtained from 1.5% (w/v) agarose gel electrophoresis in 1X TBE buffer. The gels were then stained using 1 mg cm
3 EB and photographed under UV light. 4.9. HSA binding The stock solution of HSA was prepared in Tris-HCl buffer (5 mM Tris-HCl, 10 mM NaCl, pH ¼ 7.4). Absorption titration experiments of the HSA solutions (10 mM) containing 1, 3 and 5 (0e10 mM) in MeOH were performed measuring the absorbance changes at 280 nm. The binding constant (Kb) was calculated using the BenesiHildebrand equation.70 Fluorescence titration experiments were carried out at a constant concentration of HSA (5 mM) and varying concentrations of 1, 3 and 5 (0e10 mM) upon excitation at 280 nm, and 293, 297 and 300 K. Synchronous fluorescence spectra were measured at two different Dl values (difference between the lex and lem), setting Dl ¼ 15 nm and Dl ¼ 60 nm for Tyr and Trp, respectively. The three-dimensional excitation and the emission spectra of HSA (5 mM) in the absence and presence of 1, 3 and 5 (10 mM) were recorded in the range of 200e400 nm and 200e500 nm, respectively, with a 5 nm interval. 4.10. Molecular docking Molecular docking studies were carried out using Autodock/ Vina [73]. The crystal structures of 1QC1 (CCGCCGGCGG), 1DN9 (CGCATATATGCG) and 1H9Z (HSA) were taken from the Protein Data Bank. The binding site was centered on the DNA and a grid box was created with 60  60  60 points and a 0.375 Å grid spacing in which almost the entire macromolecules were involved. For each docking calculation, 10 different poses were required within the energy range of 2 kcal mol
1. All other parameters were kept at their default values. The docked molecules were visualized by Discovery Studio 3.5 software.

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