Ferrocene-Conjugated Copper(II) Complexes of - American Chemical

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Ferrocene-Conjugated Copper(II) Complexes of L-Methionine and Phenanthroline Bases: Synthesis, Structure, and Photocytotoxic Activity Tridib K. Goswami,† Sudarshan Gadadhar,‡ Mithun Roy,† Munirathinam Nethaji,† Anjali A. Karande,*,‡ and Akhil R. Chakravarty*,† †

Department of Inorganic and Physical Chemistry, Indian Institute of Science, Bangalore 560012, India Department of Biochemistry, Indian Institute of Science, Bangalore 560012, India

‡

S Supporting Information *

ABSTRACT: Ferrocene-conjugated reduced Schiff base (Fc-metH) copper(II) complexes of L-methionine and phenanthroline bases, namely, [Cu(Fc-met)(B)](NO3), where B is 1,10-phenanthroline (phen in 1), dipyrido[3,2-d:2′,3′-f]quinoxaline (dpq in 2), dipyrido[3,2-a:2′,3′-c]phenazine (dppz in 3), and 2-(naphthalen-1-yl)1H-imidazo[4,5-f ][1,10]phenanthroline (nip in 4), were prepared and characterized and their photocytotoxicity studied (Fc = ferrocenyl moiety). Complexes [Cu(Ph-met)(B)](NO3) of the reduced Schiff base from benzaldehyde and L-methionine (PhmetH) and B (phen in 5, dppz in 6) were prepared and used as control species. Complexes 1 and 5 were structurally characterized by X-ray crystallography. Complex 1 as a discrete monomer has a CuN3OS core with the thiomethyl group as the axial ligand. Complex 5 has a polymeric structure with a CuN3O2 core in the solid state. Complexes 5 and 6 are formulated as [Cu(Phmet)(B)(H2O)](NO3) in an aqueous phase based on the mass spectral data. Complexes 1−4 showed the Cu(II)−Cu(I) and Fc+−Fc redox couples at ∼0.0 and ∼0.5 V vs SCE, respectively, in DMF−0.1 M [Bun4N](ClO4). A Cu(II)-based weak d−d band near 600 nm and a relatively strong ferrocenyl band at ∼450 nm were observed in DMF−Tris-HCl buffer (1:4 v/v). The complexes bind to calf thymus DNA, exhibit moderate chemical nuclease activity forming •OH radical species, and are efficient photocleavers of pUC19 DNA in visible light of 454, 568, and 647 nm, forming •OH radical as the reactive oxygen species. They are cytotoxic in HeLa (human cervical cancer) and MCF-7 (human breast cancer) cells, showing an enhancement of cytotoxicity upon visible light irradiation. Significant change in the nuclear morphology of the HeLa cells was observed with 3 in visible light compared to the nonirradiated sample. Confocal imaging using 4 showed its nuclear localization within the HeLa cells.

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hepatotoxicity, limiting its use.26,27 In addition, the active species generated by Photofrin is singlet oxygen formed in a type II energy transfer process involving the triplet state of the photosensitizer and molecular oxygen (3O2).28−30 The generation of triplet oxygen is very important for the efficacy of an organic PDT agent. Metal-based PDT agents having a redoxactive metal ion provide alternate pathways to form hydroxyl radicals as the reactive species. The chemistry of metal-based photocytotoxic complexes is relatively unexplored compared with organic PDT agents.13 Metal complexes of rhodium(II), ruthenium(II), and platinum(IV) are known to show novel photocytotoxic activity.10−12,31−35 The present work stems from our interest to design and develop new ferrocene conjugates of amino acids that are suitable to bind copper(II) ligated to phenanthroline base to generate mixed-metal heterodinuclear species having two redoxactive metal centers. We have designed and synthesized the complexes to study their photocytotoxic activity over a wide

INTRODUCTION The current impetus in the development of transition metalbased anticancer agents is from the successful use of cisplatin and its analogues, such as carboplatin and oxaliplatin, as anticancer drugs.1−5 The platinum-based chemotherapeutic agents suffer from dark toxicity, drug selectivity, and drug resistivity. This has necessitated further research to find cisplatin alternatives. Lippard and co-workers reported platinum(IV) prodrugs that generate cisplatin on cellular reduction in the presence of thiols.6−9 The platinum(IV) complexes having potentially labile ligands constitute a new generation of platinum-based anticancer agents. Sadler and co-workers showed that six-coordinate platinum(IV) complexes having photolabile ligands are generally nontoxic in the dark but becomes potentially cytotoxic on irradiation with light, generating four-coordinate platinum(II) species.10−12 The metal-based photochemotherapeutic agents are promising alternatives to photofrin, its analogues, and phthalocyanine bases that are currently used for photodynamic therapy (PDT) as anticancer agents using red light of 620− 800 nm wavelengths.10−20 PDT and its various aspects have been extensively reviewed.21−25 Photofrin causes skin sensitivity and © 2012 American Chemical Society

Received: November 9, 2011 Published: March 22, 2012 3010

dx.doi.org/10.1021/om201102k | Organometallics 2012, 31, 3010−3021

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spectral range of visible light. In addition, bioorganometallic chemistry is in its infancy, with the potential to generate pharmaceuticals for medicinal applications of primary and metastatic secondary forms of tumors and for their use as antimalarial, antifungal, and anti-HIV agents, antibiotics, and radiopharmaceuticals.36−46 Our interest in designing ferrocene-based photocytotoxic agents is primarily due to the stability of the ferrocenyl moiety in a biological medium besides its lipophilicity and redox activity.41 The utility of ferrocene is earlier evidenced from the enhanced activity of ferrocifen, the ferrocene-appended anticancer drug tamoxifen, against both hormone-independent (ER−) and hormone-dependent (ER+) cancer with respect to tamoxifen.42 Similarly, ferroquine, the ferrocene-attached version of the antimalarial drug chloroquine, is more active than chloroquine.43 Ruthenium-arene and ruthenium-cyclopentadienyl half-sandwich complexes are known as organometallic antitumor agents, with the water-soluble ruthenium(II)arene-pta (RAPTA) complexes showing promising results as antimetastatic agents.36−38 Mohler and co-workers reported tungsten and iron cyclopentadienyl complexes showing DNA cleavage upon photoirradiaton with UV light.44,45 Barring a few reports of organometallic agents showing DNA cleavage activity in UV light, complexes showing similar activity within the PDT spectral window are virtually limited to the ones from our laboratory.47−51 We reported copper(II) complexes of ferroceneappended dipicolylamine and terpyridine moieties and phenanthroline bases, and these complexes photocleave DNA in red light via formation of hydroxyl radicals.47−49 The presence of a redox-active ferrocenyl moiety is found to facilitate generation of reactive hydroxyl radicals compared with the singlet oxygen species. Herein, we present the chemistry of ferrocene-appended copper(II) complexes as multifunctional model nucleases capable of showing efficient DNA cleavage activity in red light. The incorporation of a ferrocenyl moiety enables us to explore the DNA cleavage activity of the complexes over a much larger visible spectral window. The synthesis, crystal structure, DNA binding, photoinduced DNA cleavage activity, and photocytotoxic properties of the ferrocene-appended reduced Schiff base (Fc-metH) copper(II) complexes of L-methionine and phenanthroline bases, viz., [Cu(Fc-met)(B)](NO3), where B is 1,10-phenanthroline (phen in 1), dipyrido[3,2-d:2′,3′-f ] quinoxaline (dpq in 2), dipyrido[3,2-a:2′,3′-c]phenazine (dppz in 3), and 2-(naphthalen-1-yl)-1H-imidazo[4,5-f ][1,10]phenanthroline (nip in 4), are reported here (Chart 1). Complexes [Cu(Ph-met)(B)](NO3), where B is phen (5) or dppz (6) and Ph-metH is the reduced Schiff base derived from benzaldehyde and L-methionine, are prepared as controls. Complexes 5 and 6 are formulated as [Cu(Ph-met)(B)(H2O)](NO3) in an aqueous solution phase. Significant enhancement in the DNA photocleavage and photocytotoxicity is observed for the ferrocene-appended complexes compared to the control species. Complex 4, having a naphthyl fluorophore, is used for cellular imaging, and the results show significant localization of the complex within the nucleus of the HeLa cells. A preliminary communication on the DNA photocleavage activity of the phen (1) and dppz (3) complexes has been published.51 We have now explored the photodynamic potential of the complexes in HeLa and MCF-7 cells and their utility in imaging cancer cells to study the nuclear uptake of the complexes.

Chart 1. Schematic Drawings of the Complexes [Cu(Fc-met)(B)](NO3) (B = phen, 1; dpq, 2; dppz, 3; nip, 4) and [Cu(Ph-met)(B)](NO3) (B = phen, 5; dppz, 6) and the Heterocyclic Bases Useda

a

X is carboxylate oxygen from another molecule in the solid-state structure or H2O in an aqueous medium.

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RESULTS AND DISCUSSION Synthesis and General Aspects. Ferrocene-conjugated ternary copper(II) complexes [Cu(Fc-met)(B)](NO3) (1−4) having heterocyclic bases (B = phen, dpq, dppz, nip) were synthesized in good yield (∼80%) from a reaction of the reduced Schiff base Fc-metH with copper(II) nitrate trihydrate and the respective phenanthroline base in methanol (Chart 1). To study the effect of the ferrocenyl moiety on the DNA photocleavage and anticancer activity, complexes [Cu(Ph-Met)(B)](NO3) (B = phen in 5 and dppz in 6) as controls were prepared by reacting the reduced Schiff base Ph-metH with copper(II) nitrate trihydrate and the phenanthroline bases. The complexes were characterized by various spectroscopic and analytical techniques. Selected physicochemical data are given in Table 1. The stability of the complexes 1−4 in a solution phase was evidenced from their ESI-MS spectra, showing essentially the molecular ion peak as [M − (NO3)]+ in aqueous methanol (Figures S1−S6, Supporting Information). The mass spectral peaks of complexes 5 and 6 correspond to [Cu(Ph-met)(B)(H2O)]+. The IR spectra of the complexes displayed characteristic stretching bands at ∼1630 cm−1 due to an asymmetric COO stretch and at ∼1385 cm−1 for NO3−. The complexes are 1:1 electrolytic with molar conductance values of ∼85 S m2 M−1 in DMF at 25 °C. Magnetic moment values of ∼1.8 μB at 25 °C suggest the presence of a one-electron paramagnetic 3d9-Cu(II) center in the complexes. The UV− visible spectra of 1−6, recorded in DMF−Tris-HCl buffer (1:4 v/v), display a broad and weak copper-centered d−d band in the range 595−690 nm (Figure 1). An intense ferrocenecentered band assignable to the 1A1g → 1E1g transition is observed near 440 nm in 1−4.52 The ligand-centered electronic transitions are observed in the UV region. Complex 4 exhibits an emission spectral band at 415 nm on excitation at 300 nm in DMF−Tris-HCl buffer (1:4 v/v) at 25 °C with a quantum yield (φ) value of 0.16 in DMF (Figure 1). The emissive property of this complex is found to be suitable for a cellular imaging study. The complexes are redox active in DMF−0.1 M TBAP (Table 1). Complexes 1−4 show a quasi-reversible response near 0.5 V vs SCE assignable to the Fc+−Fc couple (Fc, ferrocenyl moiety). There is a significant positive shift of ∼100 mV of the 3011


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Table 1. Selected Physicochemical Data for the Complexes [Cu(Fc-met)(B)](NO3) (B = phen, 1; dpq, 2; dppz, 3; nip, 4) and [Cu(Ph-met)(B)](NO3) (B = phen, 5; dppz, 6) Ef/V (ΔEp/mV)b complex 1 2 3 4 5 6

IR/cm

−1

[ν

(NO3−)]

1385 1385 1385 1385 1384 1384

−1

λmax/nm (ε/M 437 438 436 470 622 601

(255), (240), (325), (620), (118) (116)

597 605 595 690

−1 a

cm )

Fc −Fc

(144) (120) (135) (140)

0.47 (90) 0.54 (105) 0.53(150) 0.49 (140)

+

Cu(II)−Cu(I) −0.06 −0.05 −0.03 −0.12 −0.14 −0.13

(120) (445) (400) (420) (158) (260)

ΛMc/S m2 M−1

μeffd/μB

90 86 80 82 82 88

1.83 1.80 1.81 1.77 1.79 1.81

a In DMF−Tris-HCl buffer (1:4 v/v). The bands at ∼450 and 600 nm are ferrocene-based and Cu(II)-based, respectively. bFc+−Fc and Cu(II)− Cu(I) couple in DMF−0.1 M TBAP, Ef = 0.5(Epa + Epc), ΔEp = (Epa − Epc), where Epa and Epc are the anodic and cathodic peak potentials, respectively. The potentials are vs SCE. Scan rate = 50 mV s−1. cMolar conductivity in DMF. dMagnetic moment at 298 K using solid powdered samples of the complexes.

significant difference in the Cu(II)/Cu(I) redox values of two different types of complexes could be due to their structural differences, as evidenced from their crystal structures. The Cu(II) redox state is better stabilized in 5 and 6 in the presence of an axial aqua ligand. Axial binding of SMe to Cu(II) in 1−4 stabilizes the reduced state. The redox data suggest the presence of different coordination environments in these two types of complexes. The data also suggest the stability of the fivecoordinate geometries and lack of any fluxional behavior of the donor atoms. Crystal Structure. Complex 1 as its hexafluorophosphate salt (1a) and complex 5 as its perchlorate salt (5a) were structurally characterized by the single-crystal X-ray diffraction method. Complex 1a crystallized in the P1 space group of the triclinic crystal system with two molecules in the unit cell (Figure S9, Supporting Information). Complex 5a crystallized in the orthorhombic P212121 space group with four molecules in the unit cell. The ORTEP view of complex 5a is shown in Figure 3 (Figure S10, Supporting Information). Selected bond

Figure 1. Electronic spectra of [Cu(Fc-met)(dppz)](NO3) (3, ) and [Cu(Ph-met)(dppz)(H2O)](NO3) (6, ---) in DMF−Tris-HCl buffer (1:1 v/v). The arrows indicate the wavelengths of light used for the photoinduced DNA cleavage studies. The inset shows the emission spectrum of [Cu(Fc-met)(nip)](NO3) (4, ...) in DMF.

Fe(III)−Fe(II) potential in 1−4 compared to that of ferrocene (0.43 V). The complexes also show a quasi-reversible cyclic voltammetric response near 0.0 V, which is assignable to the Cu(II)−Cu(I) couple (Figure 2, Figure S7, Supporting

Figure 3. ORTEP view of the polymeric complex in [Cu(Ph-met)(phen)](ClO4) (5a) showing 50% probability thermal ellipsoids and the atom-numbering scheme for the metal and heteroatoms. The hydrogen atoms are omitted for clarity.

distance and angle data for 5a are given in Table S1 (see Supporting Information). The structure of 1, which was earlier published by us, consists of a discrete binuclear heterobimetallic complex having Cu(II) and Fe(II) centers.51 The crystal structure shows an axially elongated square-pyramidal (4 + 1) geometry of the Cu(II) center with a CuN3OS coordination geometry (τ = 0.001 and 0.134 for molecules A and B, respectively).53 The copper(II) site in 1 has structural similarity with [Cu(L)(bpy)](ClO4), which models the CuB center of dopamine β-hydroxylase (DβH) with a {CuII(L-his)3(L-met)(H2O)} core (HL = (2-(methylthio)phenyl)salicylaldimine).54

Figure 2. Cyclic voltammograms of 1 () and 5 (---) showing the redox processes in DMF−0.1 M TBAP (scan rate: 50 mV s−1).

Information). Complexes 5 and 6 have the Cu(II)−Cu(I) couple near −0.14 V vs SCE (Figure S8, Supporting Information). Ligand reductions are observed near −1.1 and −1.7 V. The 3012


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Table 2. DNA Binding Data for Complexes [Cu(Fc-met)(B)](NO3) (B = phen, 1; dpq, 2; dppz, 3; nip, 4) and [Cu(Phmet)(B)](NO3) (B = phen, 5; dppz, 6) 1 Kba/M−1 [s] ΔTmb/°C a

(7.5 ± 0.3) × 104 [0.07] 2.1

2 (1.7 ± 0.8) × 105 [0.10] 3.6

3

4

(3.7 ± 0.5) × 105 [0.13] 4.2

5

(1.8 ± 0.4) × 105 [0.09] 3.7

(4.8 ± 0.5) × 104 [0.03] 1.5

6 (3.6 ± 0.2) × 105 [0.11] 4.0

Intrinsic equilibrium DNA binding constant from the UV−visible experiment. bChange in the calf thymus DNA melting temperature.

Figure 4. (a) Derivative plots showing DNA melting temperatures using calf thymus DNA (70 μM NP) in the absence and presence of 20 μM ethidium bromide (EB) and complexes 1−6 in 5 mM phosphate buffer (pH = 6.8). (b) Effect of increasing concentration of EB (▲), Hoechst dye (◀), 1 (■), 2 (●), 3 (▼), and 4 (▶) on the relative viscosity of calf thymus DNA at 37.0(±0.1) °C in 5 mM Tris-HCl buffer (pH = 7.2, [ct-DNA] = 150 μM).

hypochromism of the spectral bands.55 The Kb values of the complexes from the McGhee and von Hippel method range from (4.8 ± 0.5) × 104 to (3.7 ± 0.5) × 105 M−1, giving the order of DNA binding strength as 3 ∼ 6 > 4 ∼ 2 > 1 > 5 (Table 2). The extended aromatic phenazine ring of the dppz complex seems to facilitate partial intercalation of this base through the DNA groove, resulting in higher binding strength of the dppz complex 3 than its dpq or phen analogue. The phen complexes 1 and 5 display similar binding strength to calf thymus DNA. The DNA melting experiments were carried out to investigate the effect of DNA duplex stability on binding the complexes. The duplex DNA at its melting temperature unwinds to give single-strand DNA, thus increasing the absorbance at 260 nm as the bases separate from each other. An intercalator generally increases the stability of the double helix, making thermal denaturation more difficult. A classical intercalator, namely, ethidium bromide (EB), stabilizes the duplex DNA to a significant extent, causing the DNA to melt at a significantly higher temperature.56 The dppz complex 3 stabilizes calf thymus DNA, giving a ΔTm value of 4.2 °C. Complexes 1−6 have comparatively lower ΔTm values, ranging from 1.5 to 4.2 °C, than EB, indicating a primarily DNA groove binding propensity of the complexes (Figure 4a, Table 2). Viscometric titration experiments were done to elucidate the relative specific viscosity of calf thymus DNA in the presence of the complexes. Unwinding and elongation of the DNA double helix occur upon intercalation, thus resulting in a change in the relative viscosity. EB as a classical intercalator shows a significant increase in the relative viscosity of the DNA solution due to an increase in the overall DNA contour length on binding to DNA.57 In contrast, groove binding or partially intercalating molecules cause little or no effect on the relative viscosity of the DNA solution. The plots of relative viscosity (η/η0)1/3, where η and η0 are the respective specific viscosity of DNA in the presence and absence of the complex, vs [complex]/[DNA] ratio for 1−6 and their comparisons with the classical intercator EB and the groove

The structure of complex 5a consists of a one-dimensional zigzag chain (Figure S11, Supporting Information). Two adjacent [Cu(Ph-met)(phen)] units are bridged by a carboxylate oxygen atom forming the chain in which the copper(II) exhibits a distorted square-pyramidal CuN3O2 geometry (τ = 0.273). The copper is coordinated to two nitrogen atoms of phen and one oxygen atom of the carboxylate and one nitrogen atom of Ph-met. The fifth coordination site is occupied by one carboxylate oxygen from another [Cu(Ph-met)(phen)] unit. The complex also shows an intramolecular π−π stacking interaction involving the phenyl ring of Ph-met and the planar aromatic ring of phen, giving an interplanar distance of 3.637 Å and an angle of 11.82°. The distance between the copper(II) centers bridged by the carboxylate oxygen of Ph-met is 5.943 Å. The Cu−N bond distances are in the range 1.994(5)−2.039(4) Å. The equatorial and axial Cu−O bond distances are 1.934(3) and 2.220(4) Å, respectively. The structural study shows that the ternary (phen)Cu(II) complexes of Fc-met and Ph-met ligands differ remarkably in their molecular structures. The structural difference is expected to influence their DNA cleavage activity in a significant manner. The polymeric structure of complex 5 seems to undergo conversion in an aqueous phase to form discrete monomeric species of formulation [Cu(Phmet)(phen)(H2O)]+, as evidenced from the mass spectral study. DNA Binding Property. The binding propensity of the complexes to calf thymus DNA was studied using spectral, DNA melting, and viscometric methods. Selected DNA binding data are given in Table 2. The intrinsic DNA binding constant (Kb) of the complexes was determined by UV−visible absorption titration by monitoring the change in the absorption intensity of the ligand-centered band of the complexes 1−6 at ∼270 nm. Observed significant hypochromicity with minor bathochromic shift suggests primarily the groove binding nature of the complexes to DNA in Tris-HCl buffer medium (Figure S12, Supporting Information). Small molecules that π-stack between two DNA base pairs are typical DNA intercalators, causing much larger bathochromic shift and 3013


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binder Hoechst dye show only minor changes in the relative specific viscosity of the complexes, indicating probable surface aggregation and/or a groove binding nature of the complexes to DNA (Figure 4b). Chemical Nuclease Activity. The DNA cleavage activity of the complexes 1−6 (10 μM) was studied in the presence of glutathione (GSH, 1 mM) as a reducing agent and hydrogen peroxide (H2O2, 200 μM) as an oxidizing agent using supercoiled (SC) pUC19 DNA (0.2 μg, 30 μM) in 50 mM Tris−HCl/50 mM NaCl buffer (pH 7.2) (Figure 5, Figure S13,

nuclease activity only in the presence of methyl green, while no significant inhibition was observed in the presence distamycin-A, suggesting the major groove binding nature of the dppz complex 3 (Figure S16, Supporting Information). DNA Photocleavage Activity. The photoinduced DNA cleavage activity of the complexes was studied using supercoiled pUC19 DNA (30 μM, 0.2 μg) in Tris-HCl/NaCl (50 mM, pH, 7.2) buffer. Gel electrophoresis was carried out to estimate the extent of nicked circular (NC) DNA formation from SC DNA (Figures 6, 7, Figures S17−S19, Supporting Information).

Figure 5. Bar diagram showing the chemical nuclease activity of complexes 1−6 (10 μM) using SC pUC19 DNA (0.2 μg, 30 μM b.p.) in the presence of 1.0 mM glutathione (GSH) as a reducing (gray) and 200 μM H2O2 as an oxidizing agent (black).

Figure 6. Bar diagram showing visible light-induced DNA cleavage activity of complexes 1−6 using SC pUC19 DNA (0.2 μg, 30 μM b.p.) for an exposure time of 2 h and laser power of 50 mW. [Complex] = 20 μM for 454 nm and 25 μM for other wavelengths. Color code: blue, 454 nm; green, 568 nm; red, 647 nm; black, unexposed to light.

Supporting Information). The bar diagram is shown in Figure 5. H2O2 and GSH are chosen due to the presence of the Fe(III)−Fe(II) and Cu(II)−Cu(I) redox couples in these ferrocenyl conjugates. Complexes 1−4 showed chemical nuclease activity in the presence of H2O2 or GSH. Complex 1 showed moderately higher chemical nuclease activity in the presence of H2O2 than complex 5 possibly due to the involvement of the ferrocenyl moiety. Control experiments with the precursor species and the phenanthroline bases, H2O2, or GSH alone did not show any cleavage of DNA under similar experimental conditions (Figure S14, Supporting Information). The chemical nuclease activity follows the order 3 (Fc-dppz) ∼ 6 (Ph-dppz) > 2 (Fc-dpq) ∼ 4 (Fc-nip) > 1 (Fc-phen) ∼ 5 (Ph-phen). The order of nuclease activity follows the duplex DNA binding strength of the complexes. To investigate the mechanistic aspects of the chemical nuclease activity, several additives, namely, hydroxyl radical scavengers (catalase, DMSO, KI) or singlet oxygen quenchers (NaN3, TEMP), were used as inhibitors (Figure S15, Supporting Information). Hydroxyl radical scavengers were found to inhibit the DNA cleavage activity of the complexes, while the singlet oxygen quenchers did not show any apparent inhibitory effect in the presence of GSH or H2O2. The mechanistic data indicate the involvement of reactive hydroxyl radicals in the DNA cleavage reactions. The DNA groove binding preference of the complexes was studied using the DNA major groove binder methyl green and the DNA minor groove binder distamycin in the DNA cleavage reactions. The chemical nuclease activity of 1 and 2 was significantly inhibited in the presence of distamycin (100 μM). No apparent inhibitory effect was observed in the presence of methyl green (100 μM), indicating a minor groove binding preference of 1 and 2. Complex 3 inhibited the chemical

Figure 7. Gel electrophoresis diagram showing the mechanistic data for the visible light-induced DNA cleavage activity of [Cu(Fcmet)(dppz)](NO3) (3) at 454 nm (20 μM, lanes 1−9) and 647 nm (25 μM, lanes 10−17) using SC pUC19 DNA (0.2 μg, 30 μM b.p.) for an exposure time of 2 h: lane 1, DNA control (454 nm); lane 2, DNA + 3; lane 3, DNA + NaN3 + 3; lane 4, DNA + TEMP + 3; lane 5, DNA + L-His + 3; lane 6, DNA + SOD + 3; lane 7, DNA + DMSO + 3; lane 8, DNA + KI + 3; lane 9, DNA + catalase + 3; lane 10, DNA + 3; lane 11, DNA + NaN3 + 3; lane 12, DNA + TEMP + 3; lane 13, DNA + L-His + 3; lane 14, DNA + SOD + 3; lane 15, DNA + DMSO + 3; lane 16, DNA + KI + 3; lane 17, DNA + catalase + 3. Additive concentrations: KI, NaN3, TEMP, L-His, 1.0 mM; DMSO, 4 μL; catalase, 4 units; SOD, 4 units.

Monochromatic visible light of 454 nm (50 mW), 568 nm (50 mW), and 647 nm (50 mW) wavelengths was used from a tunable continuous-wave (CW) Ar−Kr mixed-gas ion laser. The wavelengths chosen are based on the presence of the Fc-centered and copper(II)-centered electronic spectral bands of the complexes near 450 and 600 nm, respectively (Figure 1). Control experiments using DNA alone showed no apparent photocleavage of DNA in visible light. The phen complex 1 (20 μM) showed significant DNA cleavage activity in blue light of 454 nm, giving ∼60% cleavage of SC DNA to its NC form. A 20 μM solution of the dppz complex 3 completely nicked SC DNA, while complexes 2 (B = dpq) and 4 (B = nip) cleaved ∼80% of SC DNA in this blue light. Experiments using the Ph-met complexes 5 and 6 showed much lower DNA photocleavage activity than their ferrocenyl analogues, giving an order 3014


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of 3 (Fc-met-Cu-dppz) > 4 (Fc-met-Cu-nip) ∼ 2 (Fc-met-Cudpq) > 6 (Ph-met-Cu-dppz) > 1 (Fc-met-Cu-phen) > 5 (Phmet-Cu-phen) (Figure 6, Figure S17, Supporting Information). The complexes also showed DNA photocleavage activity in green light of 568 nm and red light of 647 nm. Complexes 2−4 (25 μM) essentially completely cleaved SC DNA in green light. The phen complex (1, 25 μM) has lower DNA cleavage activity presumably due to its lower binding strength to DNA and photoinactive nature of phen. The complexes are significant DNA photocleavers in 647 nm light, which falls within the PDT spectral window of 620−800 nm. The role of the ferrocenyl moiety in the DNA photocleavage activity is apparent from the reduced activity of the phenyl analogues 5 and 6 in blue light of 454 nm. Complexes 5 and 6 also show lower DNA cleavage activity in green and red light. The cleavage activity at 568 and 647 nm wavelengths follows the order as observed in blue light (Table 3). Control experiments using copper(II) nitrate,

photocleavage reactions were studied in visible light of 454 and 647 nm wavelengths using the dppz complex 3 and different additives (Figure 7). Singlet oxygen quenchers, namely, NaN3, TEMP, or L-His, did not show any significant inhibition in the cleavage activity. This eliminates the possibility of a type II pathway forming singlet oxygen (1O2) species. The hydroxyl radical scavengers, namely, DMSO, KI, and catalase, inhibited the DNA cleavage, suggesting the formation of hydroxyl radical via a photoredox pathway.20 Superoxide dismutase (SOD) showed partial inhibition of the DNA photocleavage activity. The involvement of superoxide radical anion is thus a possibility. The partial inhibition in the presence of SOD could be due to its spontaneous dismutation generating •OH radicals.58 Although the nature of the photochemical processes involved in the DNA photocleavage reactions is not known, the results suggest a photoredox pathway to be operative involving the visible bands of the redox-active ferrocenyl moiety and the copper(II) center since the ligands alone are inactive under similar experimental conditions.59 Cytotoxicity Study. The cytotoxicity of the complexes 1−6 in the dark and visible light in HeLa (human cervical carcinoma) and MCF-7 (human breast adenocarcinoma) cancer cells was evaluated using the MTT (3-(4,5-dimethylthiazol-2-yl)-2,5diphenyltetrazolium bromide) assay (Figures 8 and 9,

Table 3. Selected DNA (SC pUC19, 0.5 μg) Cleavage Data for [Cu(Fc-met)(B)](NO3) (B = phen, 1; dpq, 2; dppz, 3; nip, 4) and [Cu(Ph-met)(B)](NO3) (B = phen, 5; dppz, 6) in Visible Light reaction conditionsa DNA DNA DNA DNA DNA DNA DNA

control +1 +2 +3 +4 +5 +6

%NC form (λ = 454 nm)



2 62 85 95 82 33 63

3 55 78 92 77 39 75

3 58 82 97 79 38 73

a In Tris-buffer medium (pH = 7.2). λ, laser wavelength. Photoexposure time (t) = 2 h. Concentration of complexes 1−6 was 20 μM for the 454 nm and 25 μM for the 568 and 647 nm experiments.

Fc-metH, Ph-metH, phen, dpq, dppz, or nip alone did not show any apparent DNA photocleavage activity under similar reaction conditions (Figure S18, Supporting Information). The ferrocenyl moiety and the copper(II) center are not electronically coupled in the presence of a reduced Schiff base linker. The presence of the ferrocenyl moiety significantly augments the DNA photocleavage activity of the copper(II) species by providing a broader spectral range than the copper(II) species alone. The DNA photocleavage activity of 5 and 6 differs from those of 1−4 due to their major structural differences. Complexes 5 and 6 cleave DNA hydrolytically in the dark, with complex 6 showing ∼18% nicking of DNA in the dark, while its ferrocenyl analogue shows only ∼6% cleavage. This could be attributed to the presence of a labile axial ligand in the Ph-met complexes, which is occupied by a water molecule in the buffer medium. In contrast, the Fc-met structure is rigid due to sulfur coordination at the axial site, making the phosphodiester bond activation energetically unfavorable. Complexes 5 and 6 cleave DNA in the dark in the presence of radical scavengers, thus suggesting noninvolvement of any radical species in the cleavage reaction. The results indicate the hydrolysis of the phosphate backbone (Figure S19, Supporting Information). All the complexes do not show any significant DNA photocleavage activity under argon, suggesting oxygen-dependent mechanistic pathways. The reactive oxygen species could form following the type I and/or type II pathway. The mechanistic aspects of the DNA

Figure 8. Cell viability plots for the cytotoxic effect of the dppz complexes 3 (■) and 6 (●) in HeLa cells in the dark (black symbols) and in the presence of visible light (red symbols, 400−700 nm, 10 J cm−2).

Figure 9. Cell viability plots for the cytotoxic effect of the dppz complexes 3 (■) and 6 (●) in MCF-7 cells in the dark (black symbols) and in the presence of visible light (red symbols, 400− 700 nm, 10 J cm−2).

Figures S20−S23, Supporting Information). A dose-dependent antiproliferative activity of the complexes was observed in 3015


Organometallics

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Table 4. The IC50 Values of Photofrin, Cisplatin, [Cu(Fc-met)(B)](NO3) (B = phen, 1; dpq, 2; dppz, 3; nip, 4), and [Cu(Phmet)(B)](NO3) (B = phen, 5; dppz, 6) in HeLa and MCF-7 Cells HeLa

MCF-7

IC50 (μM)

IC50 (μM)

IC50 (μM)

IC50 (μM)

compound

darka

visible lightb

darka

visible lightb

1 2 3 4 5 6 Photofrinc cisplatind

5.66 (±0.35) 8.24 (±0.41) 2.61 (±0.16) 3.81 (±0.20) 12.1 (±0.6) 9.47 (±0.53) >41 71.3(±2.9)

1.85 (±0.05) 2.83 (±0.07) 0.70 (±0.04) 1.54 (±0.13) 5.78 (±0.25) 3.44 (±0.23) 4.3 (±0.2) 68.7(±3.4)

5.54 (±0.21) 13.74 (±0.62) 4.08 (±0.39) 3.87 (±0.29) 6.73 (±0.31) 4.75 (±0.33)

1.15 (±0.08) 6.32 (±0.15) 0.26 (±0.02) 1.37 (±0.21) 2.03 (±0.20) 0.47 (±0.03)

a

The IC50 values for complexes 1−6 correspond to 24 h incubation in the dark. bThe IC50 values correspond to 4 h incubation in the dark followed by photoexposure to visible light (400−700 nm, 10 J cm−2). cThe Photofrin IC50 values (633 nm excitation; fluence rate: 5 J cm−2) are taken from ref 60 (converted to μM using the approximate molecular weight of Photofrin, 600 g M−1). dThe IC50 values are taken from ref 20 for 4 h incubation. The IC50 value is 7.2 μM on 24 h incubation in the dark.

both HeLa and MCF-7 cells. Photoirradiation with visible light (400−700 nm) resulted in an enhanced cytotoxicity of the complexes. The complexes showed ∼2−4-fold increase in the cytotoxicity in HeLa cells in visible light when compared with the nonirradiated samples. The dppz complex 3 gave an IC50 value of 0.7 μM when exposed to visible light. The IC50 values of the complexes, Photofrin and cisplatin are listed in Table 4.20,60 The observed photocytotoxicity is comparable to that of Photofrin. Cisplatin, which lacks any photoactive moiety, showed an IC50 value of ∼70 μM in both dark and light under similar assay conditions. Photoexposure of the cells in the absence of the complex did not reduce cell viability. Complexes displayed antiproliferative activity on MCF-7 cell lines in the dark, and photoenhanced cytotoxicity was observed on exposure to visible light. A significantly better cytotoxicity of the ferroceneconjugated dppz complex (3) was observed in MCF-7 cells compared to the HeLa cells. The ligands and the metal salt were nontoxic to the cancer cells both in the dark and in visible light (Table S2, Supporting Information). The dpq complex 2 was found to be relatively nontoxic compared to the other complexes, and this could be due to its reduced uptake or quick efflux from the cancer cells. Acridine Orange/Ethidium Bromide Dual Staining (ref 61). To gain insight into the mechanistic aspects of cell death, acridine orange/ethidium bromide (AO/EB) dual staining of the HeLa cells treated with the photocytotoxic dppz complex 3 was done to ascertain changes in the nuclear morphology upon PDT (Figure 10). In comparison to the cells cultured in the dark, a significantly higher percentage of cells treated with complex 3 (5 μM) and irradiated with light showed apoptotic nuclei wherein the nuclei were condensed and stained intensely with EB, and the AO staining was low (Figure 10, panels c and d). Cells cultured in the dark and cells treated with DMSO showed negligible nuclear condensation and no EB staining (Figure 10, panels a and b). This indicates that the photoirradiated cells are in the late stage of apoptosis, which enables EB to enter the cells and stain the nucleus. FACScan Analysis for Apoptosis. Complex 3 showed significant activity with respect to its photocytotoxic property in light. To confirm whether the cytotoxic effect is by induction of apoptosis, flow cytometric analysis of the HeLa cells treated with complex 3 was done at three different complex concentrations.61,62 Cells treated with the complex in the dark were exposed to visible light and then stained with propidium iodide

Figure 10. Acridine orange (white arrow)/ethidium bromide (red arrow) (AO/EB) dual staining of HeLa cells treated with complex 3 (5 μM) to study the nuclear morphology. Cells treated with only DMSO in (a) the dark and (b) light. (c) Cells treated with complex 3 in the dark. (d) Cells treated with complex 3 and irradiated with visible light (400−700 nm, 10 J cm−2). The scale bar in panel a corresponds to 20 μm.

(PI) to observe any fragmented DNA, which is indicated by the increase in the sub-G1/G0 population. It was observed that a concentration of 1.25 μM of the complex induced apoptosis in the cells by 24 h, and the percentage of apoptotic population increased on increasing the complex concentration (Figure 11, Figure S24, Supporting Information). Significant induction of apoptosis by the complex was observed in the dark only at the maximum concentration used for the experiment, but still this was appreciably lower than the cellular damage observed when the cells were exposed to light. Confocal Imaging. Uptake and localization of the anticancer agents in specific organelles of the cell is a key step for their activity.63 Complex 4, bearing a fluorescent naphthyl moiety, is designed and synthesized for cellular imaging using confocal microscopy to study its uptake and localization in the cancer cells. HeLa cells, on treatment with complex 4 for different time intervals, showed nuclear localization with its predominant presence in the nucleus as early as by 1 h, as seen from the blue fluorescence in panel a of Figure 12. The complex 3016


Organometallics

Article

Schiff base, are prepared and studied. The molecular structures of the Fc-met and Ph-met complexes show significant structural differences. The Fc-met complex binds to Cu(II) in a tridentate mode with an elongated Cu−S axial bond. The −SMe moiety remains nonbonded in the Ph-met complex. The Ph-met complex shows hydrolytic cleavage of DNA, while its Fc-met analogue is inactive under similar reaction conditions. The complexes exhibit efficient groove-binding propensity to ctDNA, with partial intercalative binding of the dppz complexes. The complexes show moderate chemical nuclease activity in the presence of both oxidizing (H2O2) and reducing (GSH) agents possibly involving both the Fe(II) and Cu(II) centers. The Fc-met complexes display significant photoinduced DNA cleavage activity in blue, green, and red light. The phen and dppz complexes are significantly photocytotoxic in HeLa and MCF-7 cancer cells. The dppz complex 3 is found to induce morphological changes in HeLa cells upon photoirradiation compared to the nonirradiated samples. The nip complex 4, having a naphthyl fluorophore, on confocal imaging, shows significant localization of the complex within the nucleus of the HeLa cells. Being able to localize in the nucleus, DNA could be a major target of these compounds. The ferrocenyl copper(II) conjugates form a class of 3d-metal-based organometallic complexes that are efficient synthetic photonucleases and photocytotoxic agents. The ferrocenyl moiety enhances the PDT effect of the copper(II) complex by participating in the photoredox pathway in the presence of the redox-active Fe(II) center. This work constitutes a new development in PDT, showing the utility of organometallic complexes as nucleus-targeting photocytotoxic agents that are effective in the PDT spectral window.

Figure 11. Bar diagram showing percent apoptosis of HeLa cells induced by complex 3 at different concentrations in the absence or presence of visible light (400−700 nm) obtained from FACScan analysis.

■

EXPERIMENTAL SECTION

Materials and Measurements. All reagents and chemicals were obtained from commercial sources (s.d. Fine Chemicals, India; SigmaAldrich, USA) and used as such. Solvents used were purified by reported procedures.64 Supercoiled pUC19 DNA (cesium chloride purified) was purchased from Bangalore Genie (India). Tris(hydroxymethyl)aminomethane-HCl (Tris-HCl) buffer solution was prepared using deionized and sonicated triple-distilled water. Calf thymus (ct) DNA, agarose (molecular biology grade), distamycin, methyl green, KI, catalase, NaN3, L-histidine, superoxide dismutase, 2,2,6,6-tetramethyl-4-piperidone (TEMP), acridine orange, ethidium bromide, bromophenol blue, xylene cyanol, Dulbecco’s modified Eagle’s medium (DMEM), propidium iodide, MTT, and bovine serum albumin (BSA) were purchased from Sigma (USA). Dipyrido[3,2-d:2′,3′-f ]quinoxaline and dipyrido[3,2-a:2′,3′-c]phenazine were prepared by reported literature procedures using 1,10-phenanthroline5,6-dione as a precursor reacted with ethylenediamine and 1,2phenylenediamine, respectively.65,66 2-(Naphthalen-1-yl)-1H-imidazo[4,5-f ][1,10]phenanthroline was prepared following a literature procedure using 1,10-phenanthroline-5,6-dione and 1-naphthaldehyde as precursors.67 The ligand Fc-MetH was prepared according to the literature procedure.68 The elemental analysis was carried out using a Thermo Finnigan Flash EA 1112 CHN analyzer. The infrared, UV−visible, and emission spectra were recorded on a PerkinElmer Lambda 35, PerkinElmer Spectrum One 55, and PerkinElmer LS 55 spectrophotometer, respectively. Molar conductivity measurements were performed using a Control Dynamics (India) conductivity meter. Room-temperature magnetic susceptibility data were obtained from a George Associates Inc. Lewis-coil force magnetometer using Hg[Co(NCS)4] as a standard. Experimental susceptibility data were corrected for diamagnetic contributions.69 Cyclic voltammetric measurements were carried out at 25 °C on a EG&G PAR model 253 VersaStat potentiostat/galvanostat using a three-electrode setup with a glassy carbon working, platinum wire auxiliary, and saturated calomel

Figure 12. Time-course collection of confocal microscopic images of HeLa cells treated with complex 4 (5 μM) and propidium iodide (PI). (a, d, and g) Blue emission of complex 4 upon excitation with 405 nm laser; the respective images were taken after 1, 2, and 4 h. (b, e, and h) Red emission of PI. (c, f, and i) Merged images of the first two panels. The scale bar in panel a corresponds to 20 μm.

remained in the nucleus even after 4 h (Figure 12, panel g). The cells were stained for the nucleus with PI, which stains the nucleic acids in the presence of RNase, which degrades the cellular RNA (Figure 12, panels b, e, and h). The pink color in panels c, f, and i results from merging of the blue and red colors, indicating nuclear localization of complex 4.

■

CONCLUSION Four ferrocene-appended L-methionine reduced Schiff base copper(II) complexes containing phenanthroline bases are synthesized and characterized, and their DNA cleavage activity and cytotoxicity studied. The design of the complexes is based on the idea of incorporating the ferrocenyl moiety in a heterobinuclear unit having a Cu(II) center that is bound to a phenanthroline base that acts as photosensitizer-cum-DNA binder. To explore the effect of the ferrocenyl moiety on the cellular activity, complexes 5 and 6, having Ph-met as a reduced 3017


Organometallics

Article

(7.43, m, 5H), Ha (3.81, d, 1H, 2JHH = 12.8 Hz), Ha’ (3.62, d, 1H, 2JHH = 12.4 Hz), Hb (3.20, m, 1H), Hc (1.89, m, 2H), Hd (2.54, t, 2H, 3JHH = 7.6 Hz), He (2.11, s, 3H) (Figure S26, Supporting Information). Synthesis of [Cu(Ph-met)(B)](NO3) (B = phen, 5; dppz, 6). Complexes 5 and 6 were prepared following a similar procedure to that described for 1−4 (1.0 mmol scale). The stirring time followed by the addition of the respective phenanthroline base was 90 min, which resulted in the formation of a dark blue solution. A dark blue solid was obtained on rotary evaporation of the solvent. The solid was washed with cold methanol and water and finally dried in vacuo over P2O5 (yield: 0.48 g, ∼85% for 5, 0.52 g, ∼78% for 6) (Scheme S3, Supporting Information). Anal. Calcd for C24H24N4O5SCu (5): C, 52.98; H, 4.45; N, 10.30. Found: C, 52.79; H, 4.50; N, 10.55. ESI-MS in MeOH: m/z 499 [(M + H2O) − (NO3−)]+. UV−visible in DMF−Tris-HCl buffer (1:4 v/v) [λmax/nm (ε/M−1 cm−1)]: 622 (118), 274 (29 500). μeff = 1.79 μB at 298 K. ΛM = 82 S m2 M−1 in DMF at 25 °C. Selected IR data (KBr phase, cm−1): 3420br, 3190w, 3060w, 2915w, 2425w, 1630vs (COOasym), 1520m, 1385vs (NO3−),1105w, 805m, 750w, 725m, 700m, 650w, 550w. Anal. Calcd for C30H26N6O5SCu (6): C, 55.76; H, 4.06; N, 13.01. Found: C, 55.41; H, 4.30; N, 12.84. ESI-MS in MeOH: m/z 601 [(M + H2O) − (NO3−)]+. UV−visible in DMF−Tris-HCl buffer (1:4 v/v) [λmax/nm (ε/M−1 cm−1)]: 601(116), 378 (8660), 360 (8740), 276 (38 000). μeff = 1.81 μB at 298 K. ΛM = 88 S m2 M−1 in DMF at 25 °C. Selected IR data (KBr phase, cm−1): 3462br, 1915w, 2426w, 1633vs (COOasym), 1498 m, 1384vs (NO3), 1232w, 1137w, 1049w, 896w, 826m, 756w, 732w, 701m, 618w, 555w, 486w, 426w. Solubility and Stability. The complexes were soluble in MeOH, DMF, DMSO, MeCN, and an aqueous mixture of the solvents, less soluble in CHCl3 and CH2Cl2, and insoluble in hydrocarbon solvents. They were stable in both solid and solution phases. The stability of the complexes in the biological medium was assessed by monitoring the electronic absorption and mass spectra of the complexes in the presence of serum protein (Figures S27−S29, Supporting Information). BSA was used as a model for human serum albumin. No significant change in the electronic spectra was observed in the presence of BSA, indicating the stability of the complexes (Figure S27, Supporting Information). Mass spectral study was done to assess the stability of the complexes (Figure S28, Supporting Information). Complex 1 (200 μM) was treated with a BSA (10 μM) solution, and the sample was recorded initially and after 8 h by ESI-MS. The data revealed that the complex was stable in the presence of BSA, as no other peaks appeared in the spectra. To confirm the presence of the intact protein in solution, the same solution was recorded for MALDI-TOF MS at the same time points. In addition, differential pulse voltammetry was performed to see any probable degradation of the complex in the presence of BSA by recording the voltammetric responses of the Fc-metH ligand and complex 1 in the presence of BSA. Voltammograms of the complex remained unchanged during the study period of 8 h, inferring that the Fc-metH ligand did not leach out as a result of degradation of the complex (Figure S29, Supporting Information). X-ray Crystallographic Procedure. The crystal structures of [Cu(Fc-met)(phen)](PF6) (1a) and [Cu(Ph-met)(phen)](ClO4) (5a) were obtained by the single-crystal X-ray diffraction method. Crystals of 1a were obtained from the MeOH solution of the complex in the presence of NH4PF6 on slow evaporation of the solvent.51 Crystals of 5a were obtained from diffusion of diethyl ether into a methanol solution of 5 in the presence of NaClO4. Crystal mounting was done on a glass fiber with epoxy cement. All geometric and intensity data were collected at room temperature using an automated Bruker SMART APEX CCD diffractometer equipped with a fine-focus 1.75 kW sealed tube Mo Kα X-ray source (λ = 0.71073 Å) with increasing ω (width of 0.3° per frame) at a scan speed of 5 s per frame. Intensity data, collected using an ω−2θ scan mode, were corrected for Lorentz−polarization effects and for absorption.70 The structure solution was done by the combination of Patterson and Fourier techniques and refined by full-matrix least-squares method using the SHELX system of programs.71,72 All hydrogen atoms belonging to the complex were in their calculated positions and refined using a riding model. All non-hydrogen atoms were refined anisotropically.

reference (SCE) electrode. Tetrabutylammonium perchlorate (TBAP) (0.1 M) was used as a supporting electrolyte in DMF. The electrochemical data were uncorrected for junction potentials. 1H NMR spectra were recorded at room temperature on a Bruker 400 MHz NMR spectrometer. Electrospray ionization mass spectral measurements were done using an Esquire 3000 plus ESI (Bruker Daltonics) spectrometer. Fluorescence microscopic investigations were carried out on a Leica DM IL microscope with an integrated Leica DFC400 camera and IL50 image software. Confocal microscopy was done using a confocal scanning electron microscope (Leica, TCS SP5 DM6000). MALDI TOF mass spectra were recorded using a Bruker Daltonics Ultraflex MALDI TOF/TOF mass spectrometer. Synthesis of [Cu(Fc-met)(B)](NO3) (B = phen, 1; dpq, 2; dppz, 3; nip, 4). Complexes 1−4 were prepared by following a general synthetic procedure in which a methanol solution of LiOH·H2O was treated with Fc-metH (347 mg, 1.0 mmol) followed by its addition to a solution of Cu(NO3)2·3H2O (240 mg, 1.0 mmol) in methanol (10 mL). The mixture was stirred for 1.0 h, and a methanol solution of the respective heterocyclic base was added with continuous stirring (B: phen, 0.19 g; dpq, 0.23 g; dppz, 0.29 g; nip, 0.35 g; 1.0 mmol). The color of the solution turned dark green after 45 min. A dark green solid obtained on rotary evaporation was isolated, washed with cold methanol and water, and finally dried under vacuum over P4O10 (yield: 0.54 g, 83% for 1; 0.56 g, 80% for 2; 0.61 g, 81% for 3; 0.63 g, 77% for 4) (Scheme S1, Supporting Information). Anal. Calcd for C28H28N4O5SFeCu (1): C, 51.58; H, 4.33; N, 8.59. Found: C, 51.39; H, 4.51; N, 8.36. Selected IR data (KBr phase, cm−1): 3435br, 2920w, 1635vs (COOasym), 1385vs (NO3−), 1105m, 1040w, 825m, 730m, 480m (vs, very strong; m, medium; w, weak; br, broad). ESI-MS in MeOH: m/z 589 [M − (NO3−)]+. UV−visible in DMF−Tris-HCl buffer (1:4 v/v) [λmax/nm (ε/M−1 cm−1)]: 597 (140), 437 (255), 272 (24 360). μeff = 1.83 μB at 298 K. ΛM = 80 S m2 M−1 in DMF at 25 °C. Anal. Calcd for C30H28N6O5SFeCu (2): C, 51.18; H, 4.01; N, 11.94. Found: C, 51.01; H, 3.95; N, 11.68. Selected IR data (KBr phase, cm−1): 3420br, 3085w, 2920w, 1635vs (COOasym), 1385vs (NO3−), 1185m, 1045w, 815m, 735m, 480m. ESI-MS in MeOH: m/z 641 [M − (NO3−)]+. UV−visible in DMF−Tris-HCl buffer (1:4 v/v) [λmax/nm (ε/M−1 cm−1)]: 605 (120), 438 (240), 337 (4450), 325 (6155), 258 (44 760). μeff = 1.80 μB at 298 K. ΛM = 86 S m2 M−1 in DMF at 25 °C. Anal. Calcd for C34H30N6O5SFeCu (3): C, 54.15; H, 4.01; N, 11.14. Found: C, 53.89; H, 4.18; N, 10.88. Selected IR data (KBr phase, cm−1): 3425br, 2920w, 1630vs (COOasym), 1385vs (NO3−), 1105m, 1040w, 825m, 730m, 485m. ESI-MS in MeOH: m/z 691 [M − (NO3−)]+. UV−visible in DMF−Tris-HCl buffer (1:4 v/v) [λmax/nm (ε/M−1 cm−1)]: 594 (135), 436 (325), 376 (6915), 358 (6770), 273 (36 590). μeff = 1.81 μB at 298 K. ΛM = 90 S m2 M−1 in DMF at 25 °C. Anal. Calcd for C39H34N6O5SFeCu (4): C, 57.25; H, 4.19; N, 10.27. Found: C, 56. 98; H, 4.08; N, 10.48. Selected IR data (KBr phase, cm−1): 3414br, 3053w, 2912w, 2827w, 1562vs (COOasym), 1385vs (NO3−), 1361s, 1312m, 1259w, 1139w, 1087m, 1052w, 1026w, 809s, 778s, 727m, 660w, 648w, 472w, 430w (s, strong). ESI-MS in MeOH: m/z 755 [M − (NO3−)]+. UV−visible in DMF−Tris-HCl buffer (1:4 v/v) [λmax/nm (ε/M−1 cm−1)]: 690 (140), 470 (620), 297 (28 400), 260 (33 600). μeff = 1.77 μB at 298 K. ΛM = 82 S m2 M−1 in DMF at 25 °C. Synthesis of Ph-metH. L-Methionine (0.3 g, 2.0 mmol) was dissolved in dry methanol (20 mL) on addition of NaOH (0.08 g, 2.0 mmol) with continuous stirring. Benzaldehyde (0.2 mL, 2.0 mmol) was added dropwise to the above solution. The mixture was refluxed for 1.0 h, cooled, and then treated with an excess of solid NaBH4 with constant stirring. The solvent was removed after stirring for ∼15 min to get a sticky mass, which was dissolved in water and treated with dilute HCl to maintain a pH of ∼5−6. A white solid that precipitated out was isolated, thoroughly washed with water and cold methanol, and finally dried under vacuum over P2O5 (Scheme S2, Supporting Information). Yield: 0.35 g (∼73%). Anal. Calcd for C12H17NO2S: C, 60.22; H, 7.16; N, 5.85. Found: C, 59.95; H, 7.10; N, 5.78. ESI-MS in MeOH: m/z 262 [M + 23] (Figure S25, Supporting Information). Selected IR (KBr phase, cm−1): 1570vs (COOasym), 1375vs (COO)sym. 1H NMR (D2O, ppm): δ HPh 3018


Organometallics

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The perspective views of the molecules were obtained by ORTEP.73 Selected crystallographic data are given in Table S3. DNA Binding Methods. Calf thymus DNA binding experiments were performed in Tris-HCl and phosphate buffers at an ambient temperature. The DNA was found to be free of protein impurity, as evidenced from the ratio of the absorbance values of the DNA at 260 and 280 nm in Tris-HCl buffer as 1.9:1. Concentration of the DNA (in base pairs) was determined by absorption spectroscopy using its molar absorption coefficient value of 6600 M−1 cm−1 at 260 nm. In the UV−visible absorption titration experiments, the complex solution (50 μM) in 5 mM Tris-HCl buffer (pH 7.2) containing 20% DMF was titrated with the 210 μM DNA, and the intensity of the band at ∼260 nm was monitored for the complexes. Correction was made for the absorption of DNA itself. The spectra were recorded after equilibration for 5 min, allowing the complexes to bind to the DNA. The intrinsic equilibrium binding constant (Kb) and the fitting parameter (s) of the complexes to DNA were obtained by the McGhee−von Hippel (MvH) method using the expression of Bard and co-workers by measuring the change of the absorption intensity of the spectral bands with increasing concentration of DNA by regression analysis using eq 1:

the addition of the complex. Superoxide dismutase was used as a scavenger of O2•− radicals. The samples after incubation in a dark chamber were added to the loading buffer containing 0.25% bromophenol blue, 0.25% xylene cyanol, and 30% glycerol (3 μL), and the solution was finally loaded on 1% agarose gel containing 1.0 μg/mL ethidium bromide. Electrophoresis was carried out in a dark chamber for 2.0 h at 60 V in TAE (Tris-acetate EDTA) buffer. Bands were visualized by UV light and photographed. The extent of DNA cleavage was calculated from the intensities of the bands using a UVITEC Gel Documentation System. Corrections were done for the low level of NC form present in the original supercoiled DNA sample and for the low affinity of EB binding to the SC compared to the NC form of DNA.77 The concentrations of the complexes and additives corresponded to that in the 20 μL final volume of the sample using Tris buffer. The estimated error in measuring the band intensities was 3−5%. Cell Viability Assay. HeLa (human cervical carcinoma) and MCF-7 (human breast adenocarcinoma) cells were maintained in DMEM, supplemented with 10% fetal bovine serum (FBS), 100 IU/mL of penicillin, 100 μg/mL of streptomycin, and 2 mM Glutamax at 37 °C in a humidified 5% CO2 incubator. The adherent cultures were grown as a monolayer and were passaged once in 4−5 days by treating with 0.25% Trypsin-EDTA. The photocytotoxicity of the complexes was assessed using the MTT assay based on the ability of mitochondrial dehydrogenases in the viable cells to cleave the tetrazolium rings of MTT and form dark blue membrane-impermeable crystals of formazan that were measured at 540 nm, giving an estimate of the number of viable cells.78 Approximately, 15 × 103 HeLa cells or 2 × 104 MCF-7 cells were plated in the wells of a 96-well culture plate in DMEM supplemented with 10% fetal bovine serum and cultured overnight. Different concentrations of the complexes were added to the cells, and incubation was continued for 4 h in the dark. After incubation, the medium was replaced with 50 mM phosphate buffer (pH 7.4) containing 150 mM NaCl (PBS), and photoirradiation was done for 1 h in visible light of 400−700 nm using a Luzchem photoreactor (model LZC-1, Ontario, Canada; light fluence rate 2.4 mW cm−2; light dose 10 J cm−2). PBS was replaced with 10% DMEM after irradiation. Incubation was continued for a further period of 20 h in the dark followed by addition of 25 μL of 4 mg mL−1 of MTT to each well and incubated for an additional 3 h. The culture medium was discarded, and a 200 μL volume of DMSO was added to dissolve the formazan crystals. The absorbance at 540 nm was determined using an ELISA microplate reader (BioRad, Hercules, CA, USA). The cytotoxicity of the test compounds was measured as the percentage ratio of the absorbance of the treated cells over the untreated controls. The IC50 values were determined by nonlinear regression analysis (GraphPad Prism). Nuclear Staining Experiment. HeLa cells cultured on coverslips were photoirradiated with visible light of 400−700 nm (light fluence rate 2.4 mW cm−2; light dose 10 J cm−2) following 4 h of incubation in the dark in the presence of 5 μM complex 3. The cells were then allowed to recover for 2 h, washed three times with PBS, stained with an acridine orange/ethidium bromide mixture (1:1, 10 μM) for 15 min, and observed at 20× magnification with a fluorescence microscope.79 FACScan Analysis (ref 80). HeLa cells (0.5 × 106 cells) plated overnight were treated with different concentrations of complex 3 in DMEM for 4 h, followed by exposure to visible light (light fluence rate 2.4 mW cm−2; light dose 10 J cm−2) for 1 h. The cells were then cultured overnight, harvested, and fixed using chilled 70% ethanol. The cells were then treated with 50 μg/mL RNase A overnight at 37 °C and stained with propidium iodide staining solution (20 μg/mL PI in PBS) for 2 h at 4 °C. The cells were analyzed for apoptosis using flow cytometry (FACSCanto, Beckton Dickenson). The percentage of cells undergoing apoptosis was determined against a suitable control. The results depicted are the mean of three different experiments, taking duplicates for each treatment. Confocal Microscopy. HeLa cells (4 × 104 cells/mm2), plated on coverslips, were incubated with 5 μM of the nip complex 4 for different time intervals from 1 to 4 h in the dark, fixed with 4% paraformaldehyde for 10 min at room temperature, and washed with PBS.81 This was followed by incubation with PI staining solution (50 μg/mL RNase A,

(εa − ε f )/(ε b − ε f ) = (b − (b2 − 2Kb2C t[DNA]t /s)1/2 )/2KbC t...

(1)

where b = 1 + KbCt + Kb[DNA]t/2s, εa is the extinction coefficient observed for the charge transfer absorption band at a given DNA concentration, εf is the extinction coefficient of the complex free in solution, εb is the extinction coefficient of the complex when fully bound to DNA, Kb is the equilibrium binding constant, Ct is the total metal complex concentration, [DNA]t is the DNA concentration in nucleotides, and s as the fitting parameter gives an estimate of the binding site size in base pairs.74,75 The nonlinear least-squares analysis was done using Origin Lab, version 6.1. DNA thermal denaturation studies were carried out by monitoring the absorption intensity of the ct-DNA (70 μM) at 260 nm varying the temperature from 40 to 90 °C at a rate of 0.5 °C per minute, both in the absence and in the presence of the complexes 1−6 (20 μM) in 5 mM phosphate buffer (pH 6.8). The experiments were done using a complex to DNA molar ratio of 2:7, and the measurements were made using a Cary 300 Bio UV−visible spectrometer with a Cary temperature controller on increasing the temperature of the solution by 0.5 °C min−1. Viscometric titration experiments were performed using a Schott Gerate AVS310 automated viscometer that was thermostated at 37(±0.1) °C in a constant-temperature bath. The concentration of ct-DNA was 150 μM. The flow time was measured with an automated timer. The data are presented by plotting relative specific viscosity of DNA [(η/η0)1/3] vs [complex]/[DNA], where η is the viscosity of DNA in the presence of the complex and η0 is the viscosity of DNA alone in 5 mM Tris-HCl buffer medium. The viscosity values were calculated from the observed flow time of DNAcontaining solutions (t) corrected for that of the buffer alone (t0); η = (t − t0)/t0.76 DNA Cleavage Experiments. The cleavage of supercoiled pUC19 DNA (30 μM, 0.2 μg, 2686 base-pairs) was studied by agarose gel electrophoresis using the metal complexes in a 50 mM tris(hydroxymethyl)methane-HCl (Tris-HCl) buffer (pH 7.2) containing 50 mM NaCl. For photoinduced DNA cleavage studies, the reactions were carried out in visible light of 454, 568, and 647 nm wavelengths using a Spectra Physics Stabilite 2018-RM water-cooled mixed-gas ion laser (CW beam diameter at 1/e2 = 1.8 mm ± 10% and beam divergence with full angle = 0.7 mrad ± 10%). The laser beam power at the sample position (5 cm from the aperture with a solution path length of 5 mm) was 50 mW, measured using a Spectra Physics CW laser power meter (model 407A). After light exposure, each sample was incubated for 1.0 h at 37 °C and analyzed for the photocleaved products using agarose gel electrophoresis. The mechanistic studies were carried out using different additives for quenching singlet oxygen (NaN3, 1 mM; TEMP, 1 mM; L-histidine, 1 mM) and scavenging hydroxyl radicals (DMSO, 4 μL; KI, 1 mM; catalase, 4 units) prior to 3019


Organometallics

Article

20 μg/mL PI in PBS) for 1 h at 42 °C. The cells were washed free of excess PI, and the coverslips were mounted using 90% glycerol solution containing Mowiol, an antifade, and sealed. Images were acquired using a confocal scanning microscope (Leica, TCS SP5 DM6000) and analyzed using the LAS AF Lite software.

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ASSOCIATED CONTENT

S Supporting Information *

CIF files giving crystallographic data for complexes 1a and 5a, selected bond distances and bond angles of 5a (Table S1), cytotoxicity data (Table S2), selected crystallographic data for 5a (Table S3), synthetic schemes (S1−S3) and figures showing mass spectra of the complexes (S1−S6), cyclic voltammograms (S7, S8), ORTEP and packing diagrams (S9−S11), DNA binding plots (S12), gel electrophoresis and bar diagrams (S13−S19), cellular plots (S20−S24), ligand mass and 1H NMR spectra (S25, S26), and figures of stability measurements (S27−S29). This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected],. Fax: 91-80-23600814 (A.A.K.). E-mail: [email protected]. Fax: 91-80-23600683 (A.R.C.). Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank the Department of Science and Technology (DST), Government of India, for financial support (SR/S5/MBD-02/ 2007). We are thankful to DST for a CCD difractometer facility, and Alexander von Humboldt Foundation, Germany, for donation of an electroanalytical system. We thank Dr. Omana Joy, Puja Pai, and Kavya A. for the FACS data using the facility funded by the Department of Biotechnology, Government of India, and IRIS, IISc, and P. Janardhan for confocal microscopy. The authors are thankful to B. V. S. K. Chakravarthi for his help toward carrying out initial cellular studies. A.R.C. thanks DST for a J. C. Bose national fellowship. T.K.G. and S.G. thank CSIR, New Delhi, for research fellowships.

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REFERENCES

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