basu et al-2016-european journal of inorganic chemistry

8/17/2019 Basu Et Al-2016-European Journal of Inorganic Chemistry

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DOI: 10.1002/ejic.201501105 Full Paper

Photoactive Anticancer Compounds

Mitochondria-Targeting Iron(III) Catecholates for Photoactivated

Anticancer Activity under Red LightUttara Basu,[a] Ila Pant,[b] Paturu Kondaiah,*[b] and Akhil R. Chakravarty*[a]

Abstract: Iron(III) catecholates [Fe(R′-bpa)(R-dopa)Cl] (1, 2)with a triphenylphosphonium (TPP) moiety, where R′-bpais 2-(TPP-N ,N -bis((pyridin-2-yl)methyl)ethanamine) chloride(TPPbpa) and R-dopa is 4-{2-[(anthracen-9-yl)methylamino]-ethyl}benzene-1,2-diol (andopa, 1) or 4-{2-[(pyren-1-yl)-methylamino]ethyl}benzene-1,2-diol (pydopa, 2), were synthe-sized and their photocytotoxicity studied. Complexes 3 and 4with [phenyl-N ,N -bis(pyridin-2-yl)methyl]methanamine (phbpa)were used as controls. The catecholate complexes showed anabsorption band near 720 nm. The 5e– paramagnetic com-plexes showed a FeIII/FeII irreversible response near –0.45 V and

Introduction

Mitochondria being the cellular organelle responsible for oxida-tive damage, calcium metabolism, and cell death are activelyinvolved in several human diseases resulting from its dysfunc-tion or mutation of genes.[1–3] Mitochondrial disruption affects

the metabolic pathway from glucose oxidation to glycolysis(Warburg effect), which is an indication of malignant cells.[4]

The alterations in cancer cells that result in evasion from apop-tosis can be targeted to inhibit proliferation. Thus, it is impera-tive to target and deliver bio-active molecules to the mitochon-dria to address mitochondrial dysfunctions associated with sev-eral human diseases including cancer. Lipophilic cations can ac-cumulate in the mitochondria of cells owing to the higher (neg-ative) mitochondrial membrane potential.[5–8] Smith et al. devel-oped a strategy to target bio-active molecules to the mitochon-dria by attaching a lipophilic triphenylphosphonium cation(TPP, PPh3

+) with an alkyl linker.[9,10] These molecules wereshown to rapidly permeate the lipid bilayers because of the

large negative mitochondrial membrane potential (180–200 mV) and accumulate inside the mitochondria in culturedcells. Chen and co-workers showed the disruption of cell mor-

[a] Inorganic and Physical Chemistry Department, Indian Institute of Science,Bangalore 560012, Karnataka, India

E-mail: [email protected]

http://ipc.iisc.ernet.in/arc.html

[b] Department of Molecular Reproduction Development and Genetics, IndianInstitute of Science,

Bangalore 560012, Karnataka, India

E-mail: [email protected]

www.mrdg.iisc.ernet.in/PK.htm

Supporting information for this article is available on the WWW under

http://dx.doi.org/10.1002/ejic.201501105.

Eur. J. Inorg. Chem. 2016, 1002–1012 © 2016 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim1002

a quasi-reversible catechol/semiquinone couple near 0.5 V ver-sus saturated calomel electrode (SCE) in DMF/0.1 M tetrabutyl-ammonium perchlorate. They showed photocytotoxicity in red/visible light in HeLa, HaCaT, MCF-7, and A549 cells. Complexes1 and 2 displayed mitochondrial localization, reactive oxygenspecies (ROS) generation under red light, and apoptotic celldeath. Control complexes 3 and 4 exhibited uniform distribu-tion throughout the cell. The complexes showed DNA photocle-avage under red light (785 nm), forming hydroxyl radicals asthe ROS.

phology selectively against cancer cells both in vitro and in vivoby many lipophilic cations.[11] In this regard, Neamati and co-workers developed TPP-appended chlorambucil molecules formitochondria-targeted cytotoxicity in breast and pancreaticcancer cells.[12] A similar strategy was adopted by Dhar et al. tosynthesize a mitochondria-targeting cisplatin analog, Platin-M,

and a nanoparticle-based zinc(II) phthalocyanine photosensi-tizer that was shown to disrupt the mitochondrial genome in-stead of the nuclear DNA.[13,14] Guo and co-workers developeda cation-appended copper complex for mitochondria targetingthat was cytotoxic against cisplatin-resistant tumor cellsthrough multiple mechanisms of action.[15,16]

Photodynamic therapy (PDT) is a relatively new mode of can-cer treatment, which depends on the retention of the photo-sensitizers in the tumor cells followed by their selective activa-tion under red light in the presence of molecularoxygen.[17–20] Photosensitizers are light-sensitive compoundsthat cause localized oxidative damage within the target cellsupon irradiation. Several organic molecules, including porphy-

rins, texaphyrins, and phthalocyanines, have been reported asefficient PDT agents. However, their potential usage in cancertreatment is limited as a result of skin sensitivity and acutehepatotoxicity on prolonged use. Metal complexes with theirvaried structural, photophysical, and photochemical propertiesare thought to be ideal alternatives to the organic chromo-phores as they could circumvent the shortcomings of the or-ganic drugs.[21] Redox-active metal complexes may provide aphotoredox pathway for the generation of different reactiveoxygen species (ROS), leading to tumor damage as an alterna-tive to the generation of singlet oxygen. Research in this direc-tion has resulted several 3d–5d metal complexes that show in

vitro photocytotoxicity in visible or UV-A light.[22–25]

However,


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the examples of 3d transition-metal complexes that can bephoto-excited within the PDT window of 620–800 nm for bettertissue penetration are rare in the literature.[26,27]

We have recently reported a class of iron(III) catecholatesthat showed red light induced cytotoxicity with the complexesprimarily localizing in the nucleus of HeLa and HaCaT cells.[28,29]

Considering the drug resistance associated with nuclear target-ing drugs such as cisplatin or its analogs as a result of theextensive repair of the drug–DNA adducts by the nucleotideexcision repair (NER) pathway,[30,31] we have attempted tochange the cellular localization mode of these iron(III) com-plexes from the nucleus to the mitochondria of the cells. Toachieve this objective, we have designed and synthesized newiron(III) catecholates with a cationic TPP moiety as a pendant in[Fe(R′-bpa)(R-dopa)Cl] (1, 2), where R′-bpa is 2-{TPP-N ,N -bis[(pyridin-2-yl)methyl]ethanamine} chloride (TPPbpa) and R-dopa is functionalized dopamine, that is, 4-{2-[(anthracen-9-yl)methylamino]ethyl}benzene-1,2-diol (andopa, 1) or 4-{2-[(pyren-1-yl)methylamino]ethyl}benzene-1,2-diol (pydopa, 2),

and studied their altered cellular activity (Figure 1). The posi-tively charged lipophilic TPP cation was incorporated to drivethe complexes to the mitochondria. To retain the photocyto-toxic behavior of the complexes, the dopamine unit was func-tionalized with planar, aromatic, and photoactive anthracenyland pyrenyl groups. Moreover, to establish the role of the TPPunit, two similar complexes in which TPPbpa was substituted by[phenyl-N ,N -bis(pyridin-2-yl)methyl]methanamine (phbpa) (3, 4)were synthesized and used as controls. Herein, we report thephotocytotoxicity under red light and cellular localization of thecomplexes 1–4.

Figure 1. Schematic drawing of the structures of the iron(III) complexes 1–4.

Table 1. Selected physicochemical and calf thymus (ct)-DNA binding data for the complexes 1–4.

Parameters Complex 1 Complex 2 Complex 3 Complex 4

ESI-MS [m/ z ][a] 442.12 454.32 686.19 710.24 λ [nm] (ε [M–1 cm–1])[b] 748 (1030) 685 (3850) 745 (885) 675 (4850)

λem [nm] (Φ)[c] 425 (0.03) 395 (0.04) 420 (0.04) 400 (0.04)

Λ [S m2 M–1][d] 147 153 67 73µeff [µB]

[e] 5.81 5.83 5.82 5.85K b [M

–1][f] 1.8 ± 0.3 × 105 2.4 ± 0.5 × 105 1.3 ± 0.4 × 105 2.0 ± 0.6 × 105

[a] In methanol. [b] In DMF. [c] In DMSO. Excitation wavelengths are 390 nm for complexes 1 and 3 and 340 nm for 2 and 4. Quantum yield calculated byusing anthracene in ethanol and quinine sulfate in sulfuric acid as standards. [d] Molar conductance in DMF. [e] Magnetic moment values at 298 K. [f] Intrinsic

binding constants of the complexes to ct-DNA in 5 % DMF-Tris-HCl buffer medium.

Eur. J. Inorg. Chem. 2016, 1002–1012 www.eurjic.org © 2016 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim1003

Results and Discussion

Synthesis and General Aspects

Ligand TPPbpa was synthesized in three steps. Pyridine-2-car-baldehyde and 2-aminoethanol were stirred for 3 d in the pres-ence of sodium tris-acetoxyborohydride and glacial acetic acid

to give 2-{bis[(pyridin-2-yl)methyl]amino}ethanol. This was fol-lowed by chlorination with thionyl chloride and subsequentheating with triphenylphosphine at 100 °C in the presence of atrace amount of potassium iodide in n-butanol for 4 d to isolatethe ligand as a brown precipitate. The product was washed withchloroform and diethyl ether, dried in vacuo and characterizedby 1H NMR spectroscopy (Scheme S1, Figure S1 in the Support-ing Information). Ligand phbpa was synthesized by followingreported literature procedures.[32] The catecholate ligands weresynthesized in three steps starting from the Schiff bases by re-action with 2-(3,4-dimethoxyphenyl)ethanamine and anthra-cene-9-carbaldehyde or pyrene-1-carbaldehyde in ethanol. Thiswas followed by the reduction of the Schiff bases with sodium

borohydride in methanol. The ether linkage was finally depro-tected with boron tribromide (1 M solution in CH2Cl2) to affordthe required ligand, andopa or pydopa, as a yellow precipitate(Scheme S2). Both the ligands were characterized by 1H NMRspectroscopy (Figure S2 and Figure S3). The complexes weresynthesized by a general reaction starting from one equivalentof anhydrous ferric chloride and one equivalent of dipicolyl-amine ligand in methanol, which was stirred for an hour. Amethanolic solution of one equivalent of the catecholate li-gands previously deprotonated with two equivalents of triethyl-amine was added to the reaction mixture. Stirring for 3 hafforded violet precipitates of the complexes 1–4 (Figure 1).

The complexes were characterized by physicochemical andanalytical data. Selected data are given in Table 1. The ESI-MSof the complexes recorded in methanol showed a single molec-ular ion peak for 1 and 2, corresponding to the species [M –2Cl]2+ whereas the peaks for complexes 3 and 4 correspondedto the species [M – Cl]+ (Figures S4–S7). The IR spectra are char-acteristic of complexes showing C–O stretches of the catechol-ate within the range 1270–1260 cm–1 (Figure S8). The UV/Visiblespectra of the complexes in DMF showed a broad and intenseabsorption band around 700–750 nm that originated from thecharge transfer from the catecholate π orbital to the iron(III)dπ* orbital (Figure 2, a and Figure S9).[33] The hump near450 nm was also due to charge transfer from catechol to


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iron(III). The molar conductivity values of complexes 1 and 2 inDMF were approximately 150 S m2 M–1, in accord with their 1:2electrolytic behavior. Complexes 3 and 4 behaved as 1:1 elec-trolytes under identical conditions, giving values around70 S m2 M–1. This reaffirms our previous observations in whichthe chloride ion coordinated to the metal center is displaced

by solvent molecules.[18]

Anthracene-based emission was ob-served for complexes 1 and 3 near 420 nm, whereas 2 and 4emitted at 400 nm owing to the presence of the pyrene group(Figure 2, b and Figure S9). Magnetic moment values of approxi-mately 5.8 µB at room temperature are in agreement with theirfive-electron paramagnetic nature. The redox active complexesdisplayed an irreversible metal-based redox response near–0.45 V versus saturated calomel electrode (SCE) in DMF/0.1 Mtetrabutylammonium perchlorate and a quasi-reversible redoxresponse near 0.5 V, which is assignable to the catechol/semi-quinone couple (Figures S10 and S11).[29,34]

Figure 2. (a) The UV/Vis spectrum of complex 2 in DMF. (b) The excitation(dotted line) and emission (solid line) spectra of complex 2 (10 µM) in DMSO.

Theoretical Studies

The molecular structures of the complexes 1 and 2 were opti-mized by using density functional theory (DFT) (Figure 3, a andFigure S12).[35–37] The initial coordinates were obtained fromthe crystal structure of a previously reported complex[Fe(andpa)(cat)(NO3)] and were used for further optimiza-tion.[28,29] Time-dependent density functional theory ( TD-DFT)

Figure 3. (a) The energy-optimized structure of complex 2, showing the label-ing of the metal and the heteroatoms. (b) The frontier molecular orbitals of

complex 2.


calculations were conducted to assign the electronic transitionsof the complexes, which tallied reasonably well with the experi-mental results (Table S1). The visible absorption bands near750 nm and 420 nm were assigned to the catecholate (π) toiron(III) (π*) charge-transfer transitions.[29] The electronic transi-tions below 400 nm were assigned to intra-ligand charge-trans-

fer (ILCT) transitions involving the anthracenyl or the pyrenylgroups.[38] The orbitals involved in the LMCT transitions thatresult in the rich photophysical properties of the complexes areshown in Figure 3 (b, and Figure S12).

Stability Studies

The stability of the complexes 1–4 under physiological condi-tions was assessed by using UV/Visible spectral scans at differ-ent time intervals for 24 h. The spectra were recorded withsolutions of the complexes in 1 % DMSO/Dulbecco's phosphatebuffered saline (DPBS) and remained unaltered with respect tothe peak positions and their intensities (Figure S13). Hence, thecomplexes showed stability in the buffer medium and weredeemed suitable for biological assays.

Cytotoxicity

The anti-proliferative activities of the complexes 1–4 to inhibitcellular growth and induce cell death upon excitation with redlight (600–720 nm, 50 J cm–2) and broad band visible light(400–700 nm, 10 J cm–2) in various mammalian cell lines, thatis, HeLa (human cervical cancer), MCF-7 (human breast cancer),A549 (human lung cancer), and HaCaT (human skin keratino-cyte), were investigated by using MTT assay (Figure 4). The com-

plexes were found to be photocytotoxic with IC50 values be-tween 8–25 µM in broad band visible light (400–700 nm) and16–34 µM in red light (600–720 nm). Complexes 1 and 2 withthe triphenylphosphonium cation were found to be more cyto-toxic than complexes 3 and 4. The anthracene or the pyrene

Figure 4. The cell viability plots of 1–4 in HeLa (a), MCF-7 (b), A549 (c), andHaCaT (d) cells upon exposure to visible light (400–700 nm, 10 J cm –2, greenbars) for 1 h and red light (600–720 nm, 50 J cm –2, red bars) for 30 min or

kept in the dark (black bars).


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Table 2. Cytotoxicity values (IC50 [µM]) of the complexes 1–4 in different cell lines.

Cell lines Complex 1 Complex 2 Complex 3 Complex 4Light[a] Dark [b] Light[a] Dark [b] Light[a] Dark [b] Light[a] Dark [b]

HeLa 10.8 [25.0] 66.3 8.0 [16.6] 66.4 15.5 [32.3] 86.0 15.3 [27.4] 85.7MCF-7 16.4 [26.4] 63.2 13.9 [17.0] 68.8 24.9 [30.7] 83.9 22.0 [30.2] 85.9A549 13.4 [18.9] 63.8 10.7 [14.4] 70.9 14.3 [27.8] 86.1 13.5 [27.5] 82.6HaCaT 10.2 [19.8] 61.4 10.8 [17.4] 61.8 14.5 [27.2] 81.5 11.8 [23.3] 86.0

[a] IC50 values in µM corresponding to the cells treated with the respective complex and exposed to broad band visible light (400–700 nm, 10 J cm–2) for 1 h.

The values in parenthesis correspond to the IC50 values in µM under red light (600–720 nm, 50 J cm–2) for an exposure time of 30 min. [b] IC50 values in µM

corresponding to the cells treated with the complexes and kept in the dark. The error in the values varied between ±0.5 µ M in light to ± 2.7 µM in the dark.

photoactive moieties in complexes 1 and 2 (or their respectivecontrols; 3 and 4) behaved similarly and no significant differ-ence in their activities was observed. The dark toxicities werenear 60 µM for the TPP-appended complexes 1 and 2 and>80 µM for 3 and 4 (Table 2, Figures S14–S17). The dark toxicityof 1 and 2 might be due to the inherent toxicity of the TPPcation as reported previously.[39] The free ligand TPPbpa gavean IC50 value of approximately 50 µM under both light and dark

conditions.

Generation of Reactive Oxygen Species

Photocytotoxic iron(III) complexes are known for their ability togenerate reactive oxygen species (ROS), especially radicals suchas hydroxyl and superoxide, when exposed to light irradia-tion.[40] These are powerful oxidizing agents that can oxidizenon-fluorescent species, that is, 2′,7′-dichlorofluorescein diacet-ate (DCFDA) to a green fluorescent entity 2′,7′-dichlorofluores-cein (DCF).[41] The cell permeable fluorogenic probe undergoeshydrolysis of the ester bonds, eventually getting oxidized to

DCF in presence of ROS. Thus, we performed a DCFDA assay inHeLa cells for complexes 1 and 2 by using flow cytometry. Cellsalone did not show any signal for DCF fluorescence. The cells

Figure 5. Flow cytometric analysis (FACS) for ROS generation by complex 2using DCFDA dye in (a) HeLa and (b) HaCaT cells. The ROS generation isevident from the shift in the cell population showing a positive signal forDCF when treated with the complex and irradiated with red light (600–720 nm, 50 J cm–2) for 30 min compared with the cells alone and on treat-ment with 2 but kept in the dark [D, dark; L, light]. (c) Formation of the DNAladder isolated from the HeLa cells on treating with complex 2 for 4 h fol-lowed by exposure to red light (600–720 nm, 50 J cm–2, L) for 30 min or kept

in the dark (D). M denotes a marker of 100 base pairs.


treated with the dye showed a minor shift in the cell populationcorresponding to the basal cellular ROS. A similar shift in cellpopulation was also noted when the cells treated with the com-plexes were incubated in the dark. However, when the experi-ment was repeated in presence of red light (600–720 nm,50 J cm–2), a major shift in the cell population was noted, corre-sponding to a positive signal for DCF; see Figure 5 (a and b).This establishes the role of the ROS in cell death, species that

are produced from the complexes only with photo-irradiationbut not in the dark.

Cell Death Pathway

Cell death generally proceeds through apoptosis or necrosis. [42]

Whereas apoptosis is a programmed cell death pathway, necro-sis occurs as a result of gross injury to the cell and is oftenaccompanied by inflammation. To assess the cell death path-way, we performed a DNA fragmentation assay, which relies onthe activation of endonucleases with the onset of apoptosisand subsequent cleavage of chromatin DNA into internucleos-

omal fragments of roughly 180 base pairs (bp) and its multiples.Thus, when the DNA is run on a gel, it gives a ladder-like pat-tern. HeLa cells were treated with the TPP complex 2 for 4 hand exposed to red light irradiation (600–720 nm, 50 J cm–2). Anidentical experiment was also performed in the dark to serveas a control. The DNA was extracted and run on agarose gel.The results are shown in Figure 5 (c). There was no visible lad-der-like pattern for the sample kept in the dark. However, thelight irradiated sample showed a distinct ladder formation asseen by comparing with the DNA marker (M). This affirms thatcomplex 2 induces apoptotic cell death in HeLa cells.

Apoptosis in cells can be quantified by using an annexin-V-FITC/PI assay.[43] Annexins are a group of homologous proteinsthat bind to phosphatidylserine (a phospholipid) in a calcium-dependent manner. Annexins tagged with the fluorophore fluo-rescein isothiocyanate (FITC) give a quantitative measure of thecell population undergoing apoptosis. As membrane flipping isone of the early phenomena in the apoptotic pathway, this as-say can be used to distinguish between early and late apoptoticcells. Propidium iodide stains the DNA of the cell populationfor which the membrane has been totally compromised. Thus,a flow cytometric study was performed by using the complexes1–4 in HeLa cells under different experimental conditions. Inone set of samples, the cells were treated with the complexesfor 4 h and exposed to red light (600–720 nm, 50 J cm–2). A

second set was prepared similarly except that the samples were


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kept in the dark throughout. The results are tabulated inTable S2. There were almost three to five fold increases in thepercentage of apoptotic cell population upon irradiation com-pared with the dark controls. The results of this experimentsuggest an apoptotic pathway for cell death.

Cell Cycle Analysis

The effect of the complexes 1–4 on the progression of the cellcycle under different experimental conditions was studied todetermine the impact of light on the toxicity and estimate theapoptotic cell population. The study was carried out in HeLaand HaCaT cells and the results were consistent (Figure 6). Thepremise of several DNA binding dyes, such as propidium iodide(PI), is that the process occurs stoichiometrically. Thus, cells thatare in the S phase will have more DNA than cells in the G1phase and will proportionally take up more dye and will fluo-resce more brightly until they have doubled their DNA content.The cells in the G2 phase will be approximately twice as brightas cells in G1 as DNA replication occurs in the S phase. Whenthe cells were incubated with the complexes and kept in thedark for 4 h, there was no significant change in the populationof the various phases of the cell cycle compared with the con-trol set where cells alone were used. Again, when similarlytreated cells were exposed to red light (600–720 nm,50 J cm–2), a major population of the cells was observed belowthe G1 phase. The presence of this sub-G1 phase in the cellcycle indicates that the complexes are rather harmless in thedark but become cytotoxic in the presence of light (Figures S18and S19). The percentage of the sub-G1 population was againhigher in the case of cells treated with the TPP complexes 1 or

2 compared with those treated with 3 or 4. This again is inagreement with the higher anti-proliferative activities of thetwo complexes as discussed in the preceding section.

Figure 6. The percentage of total cell population in the sub-G1 phase of the

cell cycle in HeLa (a) or HaCaT (b) cells alone or when treated with complexes1–4 and exposed to red light (600–720 nm, 50 J cm–2, grey bars) for 30 minor kept in the dark (black bars). The control sample corresponding to cellsalone in the x axis is shown by 0.

Cellular Internalization Studies

The passage of the complexes inside HeLa and HaCaT cells wasobserved by using a flow cytometer. The fluorescence of thependant anthracenyl or the pyrenyl group was used to estimatethe cell population, which showed a positive signal for the com-pound uptake. Though not quantitative, this study enabled us

to ensure that the complexes are accumulated inside the cells


and the cytotoxicity is not a consequence of a remnant amountpresent in the media. The cytometry results showed that thecomplexes were internalized by a significant percentage of thecell population (ca. 60–80 %) in 4 h (Figure 7, Figures S20 andS21). This study prompted us to explore the localization patternof the complexes inside the HeLa and HaCaT cells, which is

discussed in the subsequent sections.

Figure 7. The percentage of total cell population showing a positive signalfor the fluorescence of the complexes 1–4 after incubation for 4 h in dark inHeLa (a) or HaCaT cells (b). The control sample corresponding to cells alonein the x axis is shown by 0.

Cellular and Sub-Cellular Localization

The cellular uptake profile of a complex is indicative of its bio-availability and stability under physiological conditions. To in-vestigate the precise pattern of localization of the complexes1–4 inside HeLa or HaCaT cells, we performed confocal micros-copy experiments using nucleus staining dye PI, which has redemission ( λem: 620 nm). The complexes are blue fluorescent( λem = 410–420 nm), which allowed us to perform a dual stain-ing assay. The merged images of the blue and the red fluores-cence for complexes 1 and 2 showed that they did not really

overlap with each other. There was rather a distinct blue colorbeyond the periphery of the red region, which denotes thenucleus. Thus, it was concluded that both 1 and 2 predomi-nantly accumulate in the cytoplasm (Figure S22). In contrast,complexes 3 and 4 showed both cytoplasmic as well as nuclearlocalization (Figure S23).

The TPP moiety was appended in the design of the com-plexes 1 and 2 to direct them to the mitochondria of the cells.Hence, we probed the mitochondria-targeting properties of thefour complexes by using Mitotracker® Deep Red (MTR). Cellswere treated similarly and were visualized under a confocalscanning microscope. The merged images for the complexes 1and 2 showed a distinct pink color arising from the exact over-lap of the red and blue fluorescence of MTR and the complexes,respectively, as can be seen in the panels i(d) and iii(d) of Fig-ure 8 (Figure S24). Complexes 3 and 4 did not show such pre-cise overlap of their fluorescence signals with MTR, as shown inthe panels ii(d) and iv(d) of Figure 8 (Figure S24). This studysubstantiates the role of the TPP moiety, which drives the com-plexes solely to the mitochondria owing to the fine balance of its lipophilicity and the positive charge.

To further verify our observation, another co-localizationstudy was done with an endoplasmic reticulum (ER) marker,that is, ER Tracker Green® (ERTG, λem: 510 nm), and complex 2in HeLa and HaCaT cells. It was observed from the merged im-

ages, as shown in the panels (d) and (h) in Figure S25, that the


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Figure 8. Confocal microscopy images of complexes 2 and 4 in HaCaT (i, ii)and HeLa (iii, iv) cells showing their sub-cellular localization pattern after

incubation for 4 h in the dark. Panels (a) correspond to the bright field imagesof the cells, panels (b) denote the emission of Mitotracker® Deep Red, panels(c) are the fluorescence of the complexes, and panels (d) correspond to themerged images of panels (b) and (c). The scale bar corresponds to 10 µm.

green and the blue fluorescence signals of the ER marker andthe complex did not coalesce accurately. This points to the factthat the TPP-appended complex does not accumulate signifi-cantly in the ER of the cells, but inside the mitochondria.

Detection of ROS in Mitochondria

The generation of ROS inside the cells was also evidenced by

using a confocal microscope. Our aim was to observe the localeof ROS generation inside the cell. HeLa or HaCaT cells weretreated with the complexes 1 and 2 for 4 h and then subjectedto light irradiation (600–720 nm, 50 J cm–2). The cells were then

Figure 9. Confocal images of complex 2 in HaCaT cells [(a)–(h)] and HeLa cells[(i)–(p)] after 4 h incubation followed by exposure to red light (600–720 nm)for 30 min or kept in the dark for 1 h. Panels (a), (e), (i), and (m) show thefluorescence of complex 2. Panels (b), (f), (j), and (n) are the fluorescenceimages of the Mitotracker® Deep Red (MTR). Panels (c), (g), (k), and (o) showfluorescence of DCF. Panels (d), (h), (l), and (p) are the merged images of the

complex, MTR, and DCF. Scale bar: 10 µm.


treated with MTR or DCFDA, then mounted on slides. Themerged images showed that there was exact overlap of thefluorescence of MTR, or DCF and the complexes, giving a bluishwhite color, as can be seen in the panels (h) and (p) of Figure 9(Figure S26). This suggests that the DCF was generated insidethe mitochondria of the cells as a consequence of ROS genera-

tion. Cell death occurs as a result of oxidative stress occurringinside the mitochondria. No ROS were detected in the absenceof light as evident from the panels (d) and (l) of Figure 9 (Fig-ure S26).

DNA Binding Studies

The mitochondria-targeting TPP complexes (1, 2) and their con-trols (3, 4) were tested for their ability to bind to calf thymus(ct)-DNA. DNA is one of the potent targets inside the cells thatmight be affected by the complexes. Thus, UV/Visible titrationswere carried out with a fixed concentration of the complexes

by varying the concentration of DNA. The hypochromic effectobserved with increasing DNA concentrations was fitted by us-ing the McGhee–von Hippel (MvH) equation and the intrinsicDNA binding constant was found to be in the order of 105 M–1.[44,45] The binding constants of the pyrene-appendedcomplexes 2 and 4 were marginally higher than those of 1 and3 with an anthracenyl moiety. This observation is a conse-quence of the higher planarity of the pyrene group, which inter-acts well with the DNA base pairs.[46] Again, the TPP-appendedcomplexes showed greater DNA binding affinity than the con-trol complexes. The DNA binding order is: 2 > 4 > 1 > 3 (Fig-ure 10 and Figure S27). Measuring the change in the viscosityof DNA before and after addition of the complexes offered an-

other method to study the interaction of DNA with the bindingmolecule. As the complexes bind between the DNA base pairs,there is an increase in the contour length, which leads to anincrease in its viscosity. Thus, the viscosities of DNA alone, inpresence of the complexes 1–4, in presence of the classical DNAintercalator ethidium bromide (EB), or Hoechst dye as a DNAgroove binder were measured. The plot of relative viscosity(η/η0)

1/3 versus [complex]/[DNA] is shown in Figure 10. The DNAbinding ability order followed the trend 2 > 4 > 1 > 3 , whichis consistent with the DNA binding data.

Figure 10. (a) Spectral traces of complex 2 showing the effect of addition of ct-DNA (250 µM) in 5 % DMF/Tris-HCl buffer (pH 7.2). (b) Plots showing theeffect of addition of an increasing quantity of the complexes 1–4, ethidiumbromide (EB as the DNA intercalator), and Hoechst dye (as the DNA groove

binder) on the relative viscosity of ct-DNA at 37.0 °C.


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DNA Cleavage Studies

The cleavage of supercoiled (SC) pUC19 DNA to its nicked circu-lar (NC) or linear form in presence of the complexes 1–4 wasstudied under different conditions in the light or dark and inpresence of various additives. A solution of complex 2 whenadded to DNA and incubated in the dark resulted in partial

cleavage (ca. 25 %) to its NC form. This hydrolytic cleavage is aconsequence of the labile coordination site occupied by thechloride ion in the solid state. A similar experiment conductedafter irradiation with a continuous-wave (CW) diode laser lightat 785 nm for 2 h showed approximately 80 % cleavage of SCDNA to its NC form (Figure 11, a and Figure S28). The othercomplexes also showed similar DNA photocleavage potential.There was no significant photocleavage of DNA under an argonatmosphere, which led us to believe that oxygen species wereinvolved in the DNA photocleavage reactions. To probe themechanistic aspects of DNA photocleavage, various additiveswere used as the reactive oxygen species quenchers/scavengers(Figure 11, b). By using singlet oxygen quenchers such as TEMP,DABCO (1,4-diazabicyclo[2.2.2]octane), and L-histidine did notsuppress the photocleavage property of complex 2. Thehydroxyl and superoxide radical scavengers, that is, KI, DMSO,superoxide dismutase (SOD), and catalase, inhibited the photo-cleavage by more than 50 %. Thus, hydroxyl and superoxideradicals are established as the ROS in the DNA photocleavagereactions.

Figure 11. (a) Bar diagram showing the percentage of nicked circular DNAformation in presence of the complexes 1–4 (25 µM) after photo-exposurefor 2 h using a CW diode laser (785 nm, 100 mW, bars 1–6): bar 1, DNA alone;bar 2, DNA + 2 (under argon); bars 3–6, DNA + 1–4, respectively. (b) Gelelectrophoresis diagram showing the percentage of NC DNA formation withcomplex 2 in the presence of various ROS quenching/scavenging agents asadditives: lane 1, DNA alone; lane 2, DNA + 2 + TEMP; lane 3, DNA + 2 +DABCO; lane 4, DNA + 2 + L-histidine; lane 5, DNA + 2 + KI; lane 6, DNA + 2+ DMSO (4 µL); lane 7, DNA + 2 + catalase (4 units); lane 8, DNA + 2 + SOD(4 units).

Conclusions

We have been successful in synthesizing mitochondria-target-ing iron(III) catecholates by suitable ligand modification. Theyshow remarkable PDT activity under red light with low toxicityin the dark. The complexes are based on suitable modificationof the previously reported iron(III) catecholates that were seento localize primarily in the nucleus of different cells.[17,18] As NERmechanisms are operative in the nucleus to rectify and revive

damaged DNA, it has been hypothesized that delivering cyto-


toxins to other cellular organelles may have a better impact forthe development of more active anticancer agents. Moreover,mitochondrial dysfunction is implicated in tumorigenesis thatrenders it a popular target for the development of various cyto-toxins. Appendage of a lipophilic triphenylphosphonium (TPP)cation has enabled the effective internalization of the com-

plexes 1 and 2 primarily into the mitochondria of the HeLa andHaCaT cells rather than the nuclei. Thus, in the presence of light,the complexes can photocleave mitochondrial DNA and not nu-clear DNA. It has further been substantiated from the confocalmicroscopy studies that in the absence of the TPP group, ageneral diffused distribution of the complexes takes placethroughout the cell without showing any selectivity. The an-thracene or pyrene groups, conjugated to the dopamine unit,have served as fluorescent markers for tracing the complexesinside the cells by cellular imaging studies. The generation of ROS precisely inside the mitochondria of the cells could be thecause of the apoptotic cell death. Photocytotoxicity studieshave shown that the iron(III) catecholates bearing either an an-

thracene or a pyrene photoactive moiety behave similarly andcould be activated by using red light owing to the presenceof an intense LMCT-type catecholate to iron(III) charge transferabsorption band near 750 nm. Such examples of 3d metal com-plexes that can be selectively activated by low energy light arerare in the literature.[47–49] The present series of near-IR lightactive photocytotoxic iron(III) complexes with their good DNAbinding, DNA photocleavage, and cellular organelle targetingproperties opens up the relatively unexplored area of metal-based PDT agents for cancer management and cure.

Experimental Section

Materials and Methods

The chemicals were purchased from S. D. Fine Chemicals, India, andSigma Aldrich, USA. The solvents were procured from commercialsources and dried by using standard procedures. Dulbecco's modi-fied eagle medium (DMEM), Dulbecco's phosphate buffered saline(DPBS), fetal bovine serum (FBS), 2′,7′-dichlorofluorescein diacetate(DCFDA), propidium iodide (PI), and annexin-V-FITC/PI kits were ob-tained from Sigma Aldrich (USA). Mitotracker® Deep Red (MTR) FM(Cat. no. M22426) was purchased from Invitrogen (USA).

A Thermo Finnigan Flash EA 1112 CHNS analyzer was used for ele-mental analysis. Electrospray ionization mass spectrometry (ESI-MS)data were obtained from an Agilent 6538 Ultra High Definition

(UHD) Accurate Mass-Q-TOF (LC-HRMS) instrument. 1

H NMR spectrawere obtained with a Bruker 400 MHz NMR spectrometer. The infra-red and UV/Visible spectra were obtained with a Bruker Alpha orPerkin–Elmer Spectrum 750 spectrophotometer, respectively. Emis-sion spectra were recorded by using a Perkin–Elmer LS 55 spectro-photometer. The molar conductivity of the complexes was meas-ured with a Control Dynamics (India) conductivity meter. Electro-chemical studies were performed by using an EG&G PAR Model 253VersaStat potentiostat/galvanostat with electrochemical analysissoftware 270. A three-electrode setup consisting of a glassy carbonelectrode (working electrode), a platinum wire (auxiliary electrode),and a saturated calomel electrode (SCE) as reference was used. Tet-rabutylammonium perchlorate (TBAP, 0.1 M) was used as a support-ing electrolyte in DMF. Magnetic susceptibility measurements of the

complexes at 300 K were recorded with solid samples by using


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MPMS SQUID VSM (Quantum Design, USA). Light irradiation wasperformed by using Waldmann 1200L PDT instrument as a red lightsource (600–720 nm) or a Luzchem Photoreactor (Model LZC-1, On-tario, Canada, fitted with eight fluorescent Sylvania white tubes,cool white, 4100 K) as a source for broad band visible light (400–700 nm). The photoreactor consists of an inbuilt cooling device tomaintain an ambient temperature inside the chamber. MTT assay

readings were obtained by using a Molecular Devices Spectra MaxM5 plate reader. Fluorescence assorted cell sorting experimentswere performed by using a FACS Verse instrument (BD Biosciences).Confocal microscopy images were acquired by using a Leica micro-scope (TCS, SP5) with oil immersion lens with a magnification of 63×.

Syntheses

The ligands and the complexes were synthesized by using the pro-cedures detailed below.

2-{TPP-N ,N -bis[(pyridin-2-yl)methyl]ethanamine} chloride

(TPPbpa)[50–52]

Sodium tris-acetoxyborohydride (16.9 g, 80 mmol) and glacial acetic

acid (3.4 mL, 60 mmol) were added to a mixture of 2-aminoethanol(1.2 g, 20 mmol) and pyridine-2-carbaldehyde (4.3 g, 40 mmol) indry tetrahydrofuran (THF, 50 mL) and stirred under a nitrogen at-mosphere at room temperature for 72 h. The solvent was removed;the residue was dissolved in dichloromethane and neutralized bythe addition of saturated sodium hydrogen carbonate solution. Theorganic fractions were separated and dried with sodium sulfate. Theremoval of the solvent under reduced pressure afforded a yellowoil as the desired product, that is N ,N -bis(2-pyridylmethyl)-2-amino-ethanol (yield 70 %), which was used without further purificationfor the next step.

N ,N -Bis(2-pyridylmethyl)-2-aminoethanol (2.4 g, 10 mmol) was dis-solved in dry dichloromethane (50 mL) and thionyl chloride (3.5 g,30 mmol) was added slowly over a time span of 2 h. The mixture

was stirred and heated at reflux for 1 h. The resulting solution wascooled and excess thionyl chloride was destroyed by dropwise addi-tion of methanol. The solvent was evaporated under reduced pres-sure to afford a yellow oil as the desired product, that is, 2-chloro-N ,N -bis[(pyridin-2-yl)methyl]ethanamine (yield 65 %).

2-Chloro-N ,N -bis[(pyridin-2-yl)methyl]ethanamine (2.6 g, 10 mmol)was dissolved in n-butanol (25 mL) and triphenylphosphine (PPh3,5.2 g, 20 mmol) was added. A pinch of KI was added and the reac-tion mixture was heated at 100 °C for 4 d. A brown precipitateformed, which was washed thoroughly with chloroform and ether,dried in vacuo, and used without further purification (yield 30 %).The compound was characterized from the 1H NMR spectrum inD2O.

1H NMR (D2O, 400 MHz): δ = 8.89 (d, J = 8.0 Hz, 2 H), 8.74–

8.60 (m, 2 H), 8.46 (d, J = 8.0 Hz, 2 H), 8.23–8.18 (m, 2 H), 7.91–7.41(m, 15 H), 4.08–4.02 (m, 4 H), 3.24 (t, J = 8.0 Hz, 2 H), 1.84 ppm (t, J = 8.0 Hz, 2 H) (Figure S1 in the Supporting Information).

4-{2-[(Anthracen-9-yl)methylamino]ethyl}benzene-1,2-diol (an-

dopa) and 4-{2-[(pyren-1-yl)methylamino]ethyl}benzene-1,2-

diol (pydopa)[53]

An ethanol solution of anthracene-9-carbaldehyde (2.0 g, 10 mmol)or pyrenecarbaldehyde (2.3 g, 10 mmol) was added to 2-(3,4-di-methoxyphenyl)ethanamine (1.8 g, 10 mmol) dissolved in ethanol(20 mL) and stirred at room temperature for 2 h. The resulting pre-cipitate of the respective Schiff base was filtered, dried in air, andrecrystallized from ethanol. The Schiff base (5 mmol) was dissolvedin methanol (50 mL) and cooled in an ice bath. Sodium borohydride

(1 g, excess) was slowly added and the reaction mixture was stirred


at room temperature for 10 h. The solvent was evaporated andwater (100 mL) was added to dissolve the inorganic side products.The product was extracted with CH2Cl2, dried with sodium sulfate,and evaporated to obtain an oily product. Drying in a desiccatorafforded a dark-yellow solid of 2-(3,4-dimethoxyphenyl)-N -[(anthra-cene-9-yl)methyl]ethanamine or 2-(3,4-dimethoxyphenyl)-N -[(pyren-1-yl)methyl]ethanamine as the product.

The reduced Schiff base (5 mmol) was dissolved in CH2Cl2 (previ-ously dried with P4O10) under a nitrogen atmosphere and cooledin an ethyl acetate/liquid nitrogen bath. Boron tribromide (BBr3, 1 Min CH2Cl2, 20 mmol) was added and the reaction mixture waswarmed to room temperature. The solution was stirred for 24 hunder a nitrogen atmosphere with occasional sonication. The result-ing suspension was again cooled in an ethyl acetate/liquid nitrogenbath and carefully quenched with methanol. The solution waswarmed to room temperature, followed by addition of water(10 mL). The solvent CH2Cl2 was evaporated and more water(50 mL) was added to the resulting suspension. Stirring for an hourafforded the desired product (andopa or pydopa) as a dark-yellowsolid, which was filtered, washed thoroughly with water, and driedin air, yield ca. 60 %. 1H NMR for andopa ([D6]DMSO, 400 MHz): δ =

8.53 (s, 1 H), 8.37 (d, J = 8.0 Hz, 2 H), 8.07 (d, J = 8.0 Hz, 2 H), 7.55–7.48 (m, 4 H), 6.83–6.71 (m, 3 H), 4.65 (s, 2 H), 3.00 (t, J = 8.0 Hz, 2H), 2.75 ppm (t, J = 8.0 Hz, 2 H) (Figure S2 in the Supporting Infor-mation). 1H NMR for pydopa ([D6]DMSO, 400 MHz): δ = 8.58–8.13(m, 9 H, pyrene), 6.70–6.50 (m, 3 H, aromatic), 4.97 (s, 2 H), 3.05 (t, J = 8.0 Hz, 2 H), 2.84 ppm (t, J = 8.0 Hz, 2 H) (Figure S3 in theSupporting Information).

Complexes 1–4

The metal complexes were synthesized by a general reaction start-ing from anhydrous ferric chloride (0.16 g, 1.0 mmol) in methanol(5 mL) and TPPbpa (0.52 g, 1.0 mmol for 1, 2) or phbpa (0.29 g,1.0 mmol for 3, 4) also in methanol (10 mL) and stirred for an hour.A methanol solution of andopa (0.34 g, 1.0 mmol for 1, 3) or pydopa

(0.37 g, 1.0 mmol for 2, 4) previously deprotonated with triethyl-amine (0.2 g, 2.0 mmol) was added to the reaction mixture. Stirringfor 3 h afforded a violet precipitate of the complex. The solid waswashed thoroughly with ether and dried with P4O10 in vacuo.

[Fe(TPPbpa)(andopa)Cl]Cl (1): Yield ca. 60 %. Elemental analysiscalcd (%) for C55H50Cl2FeN4O2P (956.75): C 69.05, H 5.27, N 5.86;found: C 68.83, H 5.52, N 5.62. ESI-MS in MeOH (m/ z ): 442.1195 [M– 2Cl]2+. FTIR: ν̃ = 3040 (w, C–H, aromatic), 2950 (w, C–H, alkyl), 1610(m), 1575 (m), 1480 (s), 1425 (w), 1260 (m, C–O), 1080 (m), 1040 (s),950 (m), 870 (m), 855 (m), 790 (m), 740 (s), 620 (m), 600 (m), 515(m), 480 (m), 410 cm–1 (m) (s, strong; m, medium; w, weak). UV/Visible spectra in DMF: λmax (ε) = 748 (1030), 468 (1480), 398(12,540), 369 (13,920), 351 (10,700), 337 nm (7450 M–1 cm–1). Emis-sion spectrum in DMSO: λem ( λex, Φ) = 425 nm (390 nm, 0.03). Molar

conductance in DMF ( ΛM): 147 S m2 M–1. Magnetic moment µeff :5.81 µB at 298 K.

[Fe(TPPbpa)(pydopa)Cl]Cl (2): Yield ca. 50 %. Elemental analysiscalcd (%) for C57H50Cl2FeN4O2P (980.77): C 69.80, H 5.14, N 5.71;found: C 71.09, H 5.86, N 5.80. ESI-MS in MeOH (m/ z ): 454.3191 [M– 2Cl]2+. FTIR: ν̃ = 3060 (w, C–H, aromatic), 2600 (w, C–H, alkyl), 1670(m), 1495 (s), 1335 (s), 1260 (m, C–O), 1050 (w), 820 (w), 750 (w),725 (w), 605 (w), 500 cm–1 (w). UV/Visible spectrum in DMF: λmax(ε) = 685 (3850), 462 (5110), 340 (25480), 325 (20680), 274 (26390),264 nm (19960 M–1 cm–1). Emission spectrum in DMSO: λem ( λex, Φ):395 n m (340 n m, 0 .04) . Mo lar c o n d uc t an c e i n DMF ( ΛM):153 S m2 M–1. Magnetic moment µeff : 5.83 µB at 298 K.

[Fe(phbpa)(andopa)Cl] (3): Yield ca. 80 %. Elemental analysis calcd

(%) for C42H38ClFeN4O2 (722.09): C 69.86, H 5.30, N 7.76; found: C


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70.03, H 5.22, N 7.68. ESI-MS in MeOH (m/ z ): 686.1936 [M – Cl]+.FTIR: ν̃ = 3080 (w, C–H, aromatic), 2955 (w, C–H, alkyl), 1630 (m),1575 (m), 1480 (s), 1435 (w), 1280 (m, C–O), 1060 (m), 940 (m), 875(m), 800 (m), 745 (s), 620 (m), 600 (m), 510 (m), 460 (m), 420 cm–1

(m). UV/Visible spectra in DMF: λmax (ε): 745 (885), 467 (1280), 389(12340), 369 (13375), 351 (10610), 336 nm (7260 M–1 cm–1). Emissionspectrum in DMSO: λem ( λex, Φ): 420 nm (390 nm, 0.04). Molar con-

ductance in DMF ( ΛM): 67 S m2 M–1. Magnetic moment µeff : 5.82 µBat 298 K.

[Fe(phbpa)(pydopa)Cl] (4): Yield 75 %. Elemental analysis calcd (%)for C44H38ClFeN4O2 (746.11): C 70.83, H 5.13, N 7.51; found: C 71.09,H 5.05, N 7.59. ESI-MS in MeOH (m/ z ): 710.2404 [M – Cl]+. FTIR: ν̃ =3040 (w, C–H, aromatic), 2650 (w, C–H, alkyl), 1650 (m), 1485 (s),1340 (s), 1270 (m, C–O), 1040 (w), 840 (w), 770 (w), 725 (w), 600 (w),505 cm–1 (w). UV/Visible spectra in DMF: λmax (ε): 675 (4850),469 (6440), 340 (31910), 324 (25490), 274 (32980), 264 nm(24950 M–1 cm–1). Emission spectrum in DMSO: λem ( λex, Φ): 400 nm(340 nm, 0.04). Molar conductance in DMF ( ΛM): 73 S m

2M–1. Mag-

netic moment µeff : 5.85 µB at 298 K.

Theoretical Calculations: The geometries of the complexes 1 and2 were optimized by density functional theory (DFT) methods usingthe B3LYP/LanL2DZ level as implemented in the Gaussian 09 pro-gram.[35–37] The electronic transitions and transition probabilitieswere obtained from linear response time-dependent density func-tional theory (TD-DFT). Selected electronic transitions for the com-plexes 1 and 2 are listed in Table S1.

MTT Assay: Cytotoxicity experiments were done to measure theanti-proliferative activities of complexes 1–4 in four different celllines, that is, HeLa (human cervical carcinoma), HaCaT (human kerat-inocyte), MCF-7 (human breast cancer), and A549 (human lung ade-nocarcinoma) cells. They were maintained in reconstituted Dul-becco's Modified Eagle's Medium (DMEM) supplemented with fetalbovine serum (FBS, 10 %), penstrep (100 µg mL–1), and glutamax

(2 mM) at 37 °C in an incubator at a CO2 level of 5 %. The adherentcultures were grown as a monolayer and were passaged once in5 d by trypsinizing with trypsin–EDTA (0.25 %). Approximately 8000cells were used for plating. Different concentrations of the com-plexes (dissolved in DMSO) were added to the cells, making thefinal DMSO concentration as 1 %. The cell plates were covered withaluminum foil to avoid light exposure and incubated for 4 h. Thiswas followed by irradiation with red or visible l ight in DPBS medium(200 µL). A dark control was also used where the media was dis-carded and replaced with fresh media (200 µL). After light treat-ment, the DPBS was removed and replaced with fresh media(200 µL). Incubation was continued for another 20 h, after whichMTT (5 mg mL–1 in DPBS) was added. After 3–4 h, the media wasremoved carefully and DMSO (200 µL) was added into each of the

wells. The absorbance reading of formazan was made at 540 nm.The cytotoxicity of the complexes was measured as the percentageratio of the absorbance of the treated cells to the untreated con-trols. The IC50 values of the complexes were determined by a non-linear regression analysis by using GraphPad Prism 5.1.

DCFDA Assay: The generation of ROS by complex 2 was studiedby a dichlorofluorescein diacetate (DCFDA) assay in HeLa and HaCaTcells. Cells were treated with complex 2 (10 µM) and incubated for4 h in the dark. Subsequently, one of the plates was exposed to redlight (600–720 nm, 50 J cm–2) for 30 min. Cells were washed withDPBS, quickly trypsinized, and harvested. DCFDA (1 µM) was addedto the cell suspensions, incubated for 15 min in ice, and the fluores-cence distribution was studied by using a FACS Verse machine (BD

Biosciences).


DNA Fragmentation Experiment: About 3 × 105 HeLa cells weregrown in 6-well tissue culture plates for 24 h in DMEM with 10 %FBS. Cells were treated with complex 2 for 4 h and one of the plateswas irradiated with red light (600–720 nm) for 30 min in the DPBSmedium. After irradiation, DPBS was removed and the cells werefurther allowed to grow for another 10 h in fresh DMEM supple-mented with 10 % FBS. Eventually, the media was discarded; cells

were washed with DPBS, trypsinized, and suspended in lysis buffer(0.4 mL, 10 mM Tris-HCl, pH 8.0, 20 mM EDTA, 0.2 % triton-X 100)for 20 min on ice. The cells were centrifuged for 20 min at13000 rpm and the supernatant was collected. It was treated withphenol and chloroform to remove all protein content. The superna-tant was precipitated by using sodium acetate (3 mM, pH = 5.8) andethanol at –20 °C overnight. The DNA pellet formed was washedwith 70 % ethanol and suspended in Tris-EDTA (TE) containingRNAse (1X Tris-EDTA with 100 mg mL–1 RNAse) for 2 h at 37 °C. Thesamples were loaded in 1.5 % agarose gel and run for about 3 h at70 V. A 100 bp marker was used as a control. They were photo-graphed in UV light.

Annexin-V-FITC/PI Assay: This assay was performed in HeLa cells

by using 1–4 to quantify the cell population showing features of apoptosis. About 3 × 105 HeLa cells were cultured in 6-well platesfor 24 h in 10 % FBS/DMEM medium. The cells were treated with1–4 for 4 h in the dark. Controls including untreated cells and cellstreated with the dyes alone were also used. One of the plates wasexposed to red light (600–720 nm, 50 J cm–2) for 30 min in DPBSmedium. Following irradiation, DPBS was removed and the cellswere allowed to grow for another 10 h in 10 % FBS/DMEM medium.The media was discarded; the cells were washed with DPBS andtrypsinized. They were suspended in binding buffer (400 µL, 1X).Annexin-V-FITC (1 µL) and PI (2 µL) were added to the cell suspen-sions and incubated for 20 min in the dark. The fluorescence wasimmediately recorded by using a flow cytometer and gating of thecell population was performed based on the cells stained by the

dyes alone.Cell Cycle Analysis Experiment: This was done to investigate theeffect of the complexes 1–4 on the cell cycle progress. Approxi-mately 1.0 × 106 HeLa or HaCaT cells were plated per well in 6-welltissue culture plates with DMEM reconstituted with 10 % FBS. Theywere allowed to grow for 24 h at 37 °C in a CO2 incubator. DMSOsolutions of the complexes (10 µM) were added to the cells andincubation was continued in the dark for 4 h. Subsequently, DMEMwas replaced with DPBS in one of the plates, which was exposedto red light irradiation (600–720 nm) for 30 min. After irradiation,DPBS was again replaced with fresh DMEM containing 10 % FBS,and the cells were incubated for another 10 h. The other plate wastreated with the complexes similarly for 4 h, after which the mediawas replaced with fresh reconstituted DMEM. Eventually, they were

processed as reported earlier.[29] Flow cytometry analysis was per-formed by using a FACS Verse machine (BD Biosciences) at FL2channel (595 nm) and the distribution of cells in the various phasesof the cell cycle was assessed from the histogram generated by“Cell Quest Pro” software (BD Biosciences). Data analysis for thepercentage of cells in each cell cycle phase was performed by usingWinMDI 2.9.

Cellular Internalization Experiments: To estimate the cell popula-tion that internalize the complexes, flow cytometry analysis wasperformed using 1–4. HeLa or HaCaT cells (1.0 × 106) were platedper well in a 6-well tissue culture plate with DMEM containing 10 %FBS. After 24 h of incubation at 37 °C in a CO2 incubator, complexes1–4 (10 µM) were added to the cells and incubated for 4 h in the

dark. The media was discarded and cells were washed three times


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with DPBS. They were trypsinized, collected in eppendorf vials, andcentrifuged. The supernatant was discarded and the cells were re-suspended in DPBS. Flow cytometry analysis was performed by us-ing a FACS Verse machine (BD Biosciences) using the Pacific blue-Afilter.

Confocal Microscopy: The localization of the complexes insideHeLa and HaCaT cells was studied by using a Leica microscope (TCS,SP5) with oil immersion lens with a magnification of 63×. Cell plat-ing was done in 12-well tissue culture plates with cover slips at aseeding density of 1 × 105 cells in 1.5 mL of the culture mediumfor 24 h. Complexes 1–4 (10 µM) were added to the cells and incu-bated in the dark for 4 h. The cells were fixed and permeabilizedby using chilled methanol for 5 min at –20 °C. After discarding themethanol, the cells were washed with DPBS and incubated withpropidium iodide (1 mg mL–1). The cover slips were mounted onslides, attached with nail enamel, and the cells were visualized withthe microscope. To probe the sub-cellular localization, a similartreatment was performed. However, the cells were not fixed but livecells were stained with either Mitotracker® Deep Red (MTR) or ERTracker® Green (250 nM) and incubated for 30 min at room temper-ature. To investigate the precise locale of ROS generation insideHeLa or HaCaT cells, confocal microscopy experiments were con-ducted by using complexes 1 and 2 along with MTR and DCFDAdyes. Cells were plated in 12-well tissue culture plates on cover slipsas described earlier. The cells were subsequently treated with thecomplexes for 4 h in the dark. One of the plates was exposed tored light (600–720 nm) irradiation for 40 min, the media was re-moved and cells were washed with DPBS. This was followed bytreatment with MTR for 30 min at room temperature and DCFDAfor 20 min. The cover slips were mounted on slides, attached withnail enamel, and visualized under a Leica microscope (TCS, SP5)with oil immersion lens with a magnification of 63×. Images wereprocessed by using LAS AF Lite software.

DNA Binding and Cleavage Experiments

To calculate the intrinsic binding constants of the complexes 1–4to calf thymus (ct)-DNA, UV/Visible titrations were performed in Tris-HCl buffer medium (5 mM, pH 7.2) with known concentrations of the complexes (30 µM) and increasing the concentrations of ct-DNA.DNA (250 µM) in the buffer medium gave a ratio of UV absorbanceat 260 and 280 nm of 2:1, indicating its protein-free nature. Theconcentration of ct-DNA in nucleobases was calculated from its ab-sorption intensity at 260 nm with ε = 6600 M–1 cm–1. The data fittingwas done by using the McGhee–von Hippel equation. [44,45] Visco-metric titrations were carried out by using a Schott AVS. 310 Auto-mated Viscometer. Initially, the concentration of ct-DNA was keptat 150 µM while the flow times were measured with an automatedtimer. Data were represented as (η/η0)

1/3 versus [complex]/[DNA],where η is the viscosity of DNA in the presence of the complex andη0 is that of ct-DNA alone. Viscosity values were calculated from theobserved flow time of DNA containing solutions (t ) of the com-plexes corrected for the flow time of the buffer alone (t 0), whereη = (t – t 0)/t 0.

DNA cleavage experiments were performed by using supercoiledpUC19 DNA (0.2 µg, 30 µM) and the complexes (20 µM) under vari-ous experimental conditions. A diode laser of 785 nm wavelength[100 mW, Model: LQC785–100C, Newport Corporation, LD module,continuous wave (CW) circular beam, power 100 mW] was used. Ina typical experiment, the total volume of 20 µL, consisting of DNA(1 µL), NaCl (50 mM, 1 µL), complex (20 µM), and Tris-HCl buffer(50 mM, pH 7.2), was incubated for 1 h at 37 °C followed by irradia-tion for 2 h at room temperature. Following further incubation for

1 h, the samples were loaded in 1 % agarose gel containing ethid-


ium bromide (1 µg mL–1) using loading dye (25 % bromophenolblue, 0.25 % xylene cyanol, and 30 % glycerol, 2 µL). The gel wasrun in TAE (Tris-acetate-EDTA) buffer for 2 h at 60 V. The gel wasphotographed by using a UVITECH Gel documentation system. Themechanistic aspects of DNA photocleavage studies were conductedby using various additives, that is, KI (0.5 mM), DMSO (4 µL), SOD (4units), catalase (4 units), TEMP (0.5 mM), DABCO (0.5 mM), and L-

histidine (0.5 mM), which were added prior to addition of the com-plex. The procedures followed are described elsewhere.[29]

Acknowledgments

The authors thank the Indian Institute of Science (IISc), Depart-ment of Science and Technology (DST), Government of India(grant number SR/S5/MBD-02/2007 and J. C. Bose Fellowship toA. R. C.) and the Council of Scientific and Industrial Research(CSIR), New Delhi (grant number 01(2559)/12/EMR-II/2012) forfinancial support. The authors also thank the Alexander vonHumboldt (AvH) Foundation, Germany, for an electrochemicalsystem. The authors thank Ms. Koushambi Mitra for helping

with the theoretical calculations, Mr. Vashista K. for the FACSdata, and Dr. Santosh Poddar for the confocal microscopy im-ages.

Keywords: Bioinorganic chemistry · Medicinal chemistry ·Anticancer agents · Iron · Mitochondrial localization

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Received: September 28, 2015Published Online: February 2, 2016

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