Photoinduced DNA and Protein Cleavage Activity of Ferrocene

Organometallics 2009, 28, 1495–1505

1495

Photoinduced DNA and Protein Cleavage Activity of Ferrocene-Conjugated Ternary Copper(II) Complexes Basudev Maity, Mithun Roy, Sounik Saha, and Akhil R. Chakravarty* Department of Inorganic and Physical Chemistry, Indian Institute of Science, Bangalore 560012, India ReceiVed October 29, 2008

Ferrocene-conjugated ternary copper(II) complexes [Cu(L)(B)](ClO4)2, where L is FcCH2N(CH2Py)2 (Fc ) (η5-C5H4)FeII(η5-C5H5)) and B is a phenanthroline base, viz., 2,2′-bipyridine (bpy, 1), 1,10phenanthroline (phen, 2), dipyrido[3,2-d:2′,3′-f]quinoxaline (dpq, 3), and dipyrido[3,2-a:2′,3′-c]phenazine (dppz, 4), have been synthesized and characterized by various spectroscopic and analytical techniques. The bpy complex 1, as its hexafluorophosphate salt, has been structurally characterized by X-ray crystallography. The molecular structure shows the copper(II) center having an essentially square-pyramidal coordination geometry in which L with a pendant ferrocenyl (Fc) moiety and bpy show respective tridentate and bidentate modes of binding to the metal center. The complexes are redox active, showing a reversible cyclic voltammetric response of the Fc+-Fc couple near 0.5 V vs SCE and a quasi-reversible Cu(II)-Cu(I) couple near 0.0 V. Complexes 2-4 show binding affinity to calf thymus (CT) DNA, giving binding constant (Kb) values in the range of 4.2 × 104 to 2.5 × 105 M-1. Thermal denaturation and viscometric titration data suggest groove binding and/or a partial intercalative mode of binding of the complexes to CT DNA. The complexes show good binding propensity to the bovine serum albumin (BSA) protein, giving KBSA values of ∼104 M-1 for the bpy and phen complexes and ∼105 M-1 for the dpq and dppz complexes. Complexes 2-4 exhibit efficient chemical nuclease activity in the presence of 3-mercaptopropionic acid (MPA) as a reducing agent or hydrogen peroxide (H2O2) as an oxidizing agent. Mechanistic studies reveal formation of hydroxyl radicals as the reactive species. The dpq and dppz complexes are active in cleaving supercoiled (SC) pUC19 DNA on photoexposure to visible light of different wavelengths including red light using an argon-krypton mixed gas ion laser. Mechanistic investigations using various inhibitors reveal the formation of hydroxyl radicals in the DNA photocleavage reactions. The dppz complex 4, which shows efficient photoinduced BSA cleavage activity, is a potent multifunctional model nuclease and protease in the chemistry of photodynamic therapy (PDT) of cancer. Introduction Bioorganometallic chemistry is an emerging area in chemical biology due to varied biological applications of organometallic compounds.1-10 Among different organometallic systems, those based on ferrocene and its conjugates are of particular importance for their medicinal activity due to excellent stability of the ferrocene moiety in biological media, its lipophilicity, and its reversible redox chemistry. In addition, ferrocene could readily form a variety of stable conjugate species. For example, anticancer and antimalarial drugs, namely, ferrocifen (ferroceneconjugated tamoxifen) and ferroquine (ferrocene-attached chlo* To whom correspondence should be addressed. Fax: 91-80-23600683. E-mail: [email protected]. (1) Guo, Z.; Sadler, P. J. Angew. Chem., Int. Ed. 1999, 38, 1512–1531. (2) (a) Fish, R. H.; Jaouen, G. Organometallics 2003, 22, 2166–2177. (b) Jaouen, G.; Vessie’res, A.; Butler, I. S. Acc. Chem. Res. 1993, 26, 361– 369. (3) Ryabov, A. D. Angew. Chem., Int. Ed. Engl. 1991, 30, 931–941. (4) Metzler-Nolte, N. Angew. Chem., Int. Ed. 2001, 40, 1040–1043. (5) Bergamo, A.; Sava, G. Dalton Trans. 2007, 1267–1272. (6) Van Staveren, D. R.; Metzler-Nolte, N. Chem. ReV. 2004, 104, 5931– 5985. (7) Melchart, M.; Sadler, P. J. In Bioorganometallics; Jaouen, G., Ed.; Wiley-VCH: Weinheim, 2006; pp 39-64. (8) Fish, R. H. Coord. Chem. ReV. 1999, 185, 569–584. (9) Severin, K.; Bergs, R.; Beck, W. Angew. Chem., Int. Ed. 1998, 37, 1634–1654. (10) Strohfeldt, K.; Tacke, M. Chem. Soc. ReV. 2008, 37, 1174–1187.

roquine), are known to enhance the activity of the drugs.11,12 It has been reported that the cytotoxicity of tamoxifen is greatly enhanced due to the presence of a ferrocenyl moiety (Fc ) (η5C5H4)FeII(η5-C5H5)) showing activity toward estrogen-dependent and estrogen-independent breast cancer cells.11 In the case of chloroquine, the ferrocenyl moiety strongly influences the antimalarial activity.12 Moreover, bioconjugates of metallocenes with protein, peptide, DNA, RNA, and sugar are now being used in various biochemical studies.13-16 Besides ferroceneconjugated species, other bioorganometallic compounds such as arene-ruthenium complexes form a class of stable and watersoluble complexes showing anticancer activity under in Vitro and in ViVo reaction conditions.17-20 Such complexes are known (11) Jaouen, G.; Top, S.; Vessires, A.; Leclercq, G.; McGlinchey, M. J. Curr. Med. Chem. 2004, 11, 2505–2517. (12) (a) Biot, C. Curr. Med. Chem. Anti-Infect. Agents 2004, 3, 135– 147. (b) Biot, C.; Glorian, G.; Maciejewski, L. A.; Brocard, J. S.; Domarle, O.; Blampain, G.; Millet, P.; Georges, A. J.; Abessolo, H.; Dive, D.; Lebibi, J. J. Med. Chem. 1997, 40, 3715–3718. (13) Moriuchi, T.; Fujiwara, T.; Hirao, T. J. Organomet. Chem. 2007, 692, 1353–1357. (14) Noor, F.; Wu¨stholz, A.; Kinscherf, R.; Metzler-Nolte, N. Angew. Chem., Int. Ed. 2005, 44, 2429–2432. (15) Landells, J. S.; Kerr, J. L.; Larsen, D. S.; Robinson, B. H.; Simpson, J. J. Chem. Soc., Dalton Trans. 2000, 1403–1409. (16) McReynolds, L.; O’Malley, B. W.; Nisbet, A. D.; Fothergill, J. E.; Givol, D.; Fields, S.; Robertson, M.; Brownlee, G. G. Nature 1978, 273, 723–728. (17) Liu, H.-K.; Berners-Price, S. J.; Wang, F.; Parkinson, J. A.; Xu, J.; Bella, J.; Sadler, P. J. Angew. Chem., Int. Ed. 2006, 45, 8153–8156.

10.1021/om801036f CCC: $40.75  2009 American Chemical Society Publication on Web 02/12/2009

1496 Organometallics, Vol. 28, No. 5, 2009

to control tumor malignancy as antimetastasis agents.21,22 This is an important development in anticancer therapy, in which the organometallic complexes act as inhibitors to protein kinases that are overexpressed in the cancer cells causing malignancy.23-25 Transition metal complexes as synthetic nucleases have found varied applications in nucleic acid chemistry, namely, sequencespecific DNA-damaging agents, as foot printing and therapeutic agents.26-30 Among different modes of DNA cleavage, oxidative cleavage of DNA in visible light is of importance in the chemistry of photodynamic therapy (PDT) of cancer.31-35 PDT is a noninvasive therapeutic treatment of cancer in which photoactivation of a drug is selectively done at the cancer cells, leaving the healthy cells unaffected. The FDA-approved PDT drug is Photofrin. It is a porphyrin species that is active on photoactivation of its lowest energy Q-band at 630 nm red light, generating a 1ππ* state with subsequent formation of a triplet state that activates molecular oxygen from its stable triplet to the reactive singlet state (1O2).31 Porphyrin-based drugs are known to show dark toxicity and hepatotoxicity due to bilirubin formation on oxidation.36,37 This has led to the current surge in research activity on metal-based drugs as substitutes for porphyrin-based PDT drugs.38-40 Although metal-based chemotherapeutic agents such as cisplatin and its analogues have been

(18) Hambley, T. V. Dalton Trans. 2007, 4929–4937. (19) Yan, Y. K.; Melchart, M.; Habtemariam, A.; Sadler, P. J. Chem. Commun. 2005, 4764–4776. (20) Auzias, M.; Therrien, B.; Suss-Fink, G.; Sy¨tepˇnickˇa, P.; Ang, W. H.; Dyson, P. J. Inorg. Chem. 2008, 47, 578–583. (21) Patricia, S. S. Nat. Med. 2006, 12, 895–904. (22) Dyson, P. J.; Sava, G. Dalton Trans. 2006, 1929–1933. (23) Atilla-Gokcumen, G. E.; Williams, D. S.; Bregman, H.; Pagano, N.; Meggers, E. ChemBioChem 2006, 7, 1443–1450. (24) Schatzschneider, U.; Metzler-Nolte, N. Angew. Chem., Int. Ed. 2006, 45, 1504–1507. (25) Smalley, K. S. M.; Contractor, R.; Haass, N. K.; Kulp, A. N.; AtillaGokcumen, G. E.; Williams, D. S.; Bregman, H.; Flaherty, K. T.; Soengas, M. S.; Meggers, E.; Herlyn, M. Cancer Res. 2007, 67, 209–217. (26) (a) Sigman, D. S. Acc. Chem. Res. 1986, 19, 180–186. (b) Sigman, D. S.; Mazumder, A.; Perrin, D. M. Chem. ReV. 1993, 93, 2295–2316. (27) (a) Pogozelski, W. K.; Tullius, T. D. Chem. ReV. 1998, 98, 1089– 1107. (b) McMillin, D. R.; McNett, K. M. Chem. ReV. 1998, 98, 1201– 1219. (28) Erkkila, K. E.; Odom, D. T.; Barton, J. K. Chem. ReV. 1999, 99, 2777–2795. (29) De Oliveira, M. C. B.; Scarpellini, M.; Neves, A.; Terenzi, H.; Bortoluzzi, A. J.; Szpoganics, B.; Greatti, A.; Mangrich, A. S.; de Souza, E. M.; Fernandez, P. M.; Soares, M. R. Inorg. Chem. 2005, 44, 921–929. (30) Christianson, D. W.; Lipscomb, W. N. Acc. Chem. Res. 1989, 22, 62–69. (31) Bonnett, R. Chemical Aspects of Photodynamic Therapy; Gordon & Breach: London, U.K., 2000. (32) Detty, M. R.; Gibson, S. L.; Wagner, S. J. J. Med. Chem. 2004, 47, 3897–3915. (33) Henderson, B. W.; Busch, T. M.; Vaughan, L. A.; Frawley, N. P.; Babich, D.; Sosa, T. A.; Zollo, J. D.; Dee, A. S.; Cooper, M. T.; Bellnier, D. A.; Greco, W. R.; Oseroff, A. R. Cancer Res. 2000, 60, 525–529. (34) (a) Sternberg, E. D.; Dolphin, D.; Bru¨ckner, C. Tetrahedron 1998, 54, 4151–4202. (b) De Rosa, M. C.; Crutchley, R. J. Coord. Chem. ReV. 2002, 233-234, 351–371. (35) Sessler, J. L.; Hemmi, G.; Mody, T. D.; Murai, T.; Burrell, A.; Young, S. W. Acc. Chem. Res. 1994, 27, 43–50. (36) Ochsner, M. J. Photochem. Photobiol. B 1996, 32, 3–9. (37) Moriwaki, S. I.; Misawa, J.; Yoshinari, Y.; Yamada, I.; Takigawa, M.; Tokura, Y. Photodermatol. Photoimmunol. Photomed. 2001, 17, 241– 243. (38) Chifotides, H. T.; Dunbar, K. R. Acc. Chem. Res. 2005, 38, 146– 156. (39) Angeles-Boza, A. M.; Chifotides, H. T.; Aguirre, J. D.; Chouai, A.; Fu, P. K.-L.; Dunbar, K. R.; Turro, C. J. Med. Chem. 2006, 49, 6841– 6847. (40) Detty, M. R.; Gibson, S. L.; Wagner, S. J. J. Med. Chem. 2004, 47, 3897–3915.

Maity et al.

successfully used, they show toxicity and drug resistivity.41,42 To circumvent the problems, new methodologies have been developed. Sadler and co-workers have recently reported a Pt(IV) diazido complex that shows photocytotoxicity to human ovarian carcinoma cells.43 Lippard and co-workers have reported a Pt(IV) prodrug that on cellular reduction generates cisplatin using an effective targeted delivery system.44 In contrast, the chemistry of metal-based compounds showing photocytotoxicity is relatively unexplored.38 Dirhodium(II,II) complexes are known to show photocytotoxicity in visible light.38 There is a recent report on the use of ruthenium nitrosyl complexes showing cellular activity on photorelease of NO.45 The use of organometallic complexes in the chemistry of PDT is relatively unknown. The present work stems from our interest in exploring the chemistry of ferrocene-conjugated 3d metal complexes as multifunctional model nucleases and antimetastasis agents for their potential use in PDT. Earlier reports from our laboratory have shown that 3d metal complexes having planar phenanthroline bases cleave DNA in visible light in the absence of any external agents.46-50 We have planned to use ferroceneconjugated copper(II) mixed-metal binuclear complexes to explore photoinduced DNA cleavage activity over a broad visible spectral window involving both metal centers. In addition, the complexes having two redox-active metal ions are likely to show multifunctional chemical and photonuclease activity. We are also interested in studying the protein binding and photoinduced protein cleavage activity of the Fc-conjugated copper(II) complexes. Although there are only a few reports available on synthetic organometallic photonucleases, such complexes show DNA cleavage activity in the mercury arc lamp and are thus not suitable for any PDT applications.51,52 Herein, we report the synthesis, structure, DNA and protein binding, and DNA and protein photocleavage activity of the ferroceneconjugated copper(II) complexes [Cu(L)(B)](ClO4)2 (1-4), where L is ferrocenylmethylbis(2-pyridylmethylamine) (FcCH2N(CH2Py)2) and B is a N, N-donor heterocyclic base, namely, 2,2′-bipyridine (bpy, 1), 1,10-phenanthroline (phen, 2), dipyrido[3,2-d:2′,3′-f]quinoxaline (dpq, 3), and dipyrido[3,2-a: 2′,3′-c]phenazine (dppz, 4) (Chart 1). Complex 1, as the (41) (a) Zhang, C. X.; Lippard, S. J. Curr. Opin. Chem. Biol. 2003, 7, 481–489. (b) Lippard, S. J. Chem. ReV. 1999, 99, 2467–2498. (c) Rosenberg, B.; Vancamp, L.; Trosko, J. E.; Mansour, V. H. Nature 1969, 222, 385– 386. (42) (a) Fricker, S. P. Dalton Trans. 2007, 4903–4917. (b) Reisner, E.; Arion, V. B.; Keppler, B. K.; Pombeiro, A. J. L. Inorg. Chim. Acta 2008, 361, 1569–1583. (c) Galanski, M.; Jakupec, M. A.; Keppler, B. K. Curr. Med. Chem. 2005, 12, 2075–2079. (43) Mackay, F. S.; Woods, J. A.; Heringova´, P.; Kasˇpa´rkova´, J.; Pizarro, A. M.; Moggach, S. A.; Parsons, S.; Brabec, V.; Sadler, P. J. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 20743–20748. (44) Dhar, S.; Liu, Z.; Thomale, J.; Dai, H.; Lippard, S. J. J. Am. Chem. Soc. 2008, 130, 11467–11476. (45) Rose, M. J.; Fry, N. L.; Marlow, R.; Hinck, L.; Mascharak, P. K. J. Am. Chem. Soc. 2008, 130, 8834–8846. (46) Dhar, S.; Senapati, D.; Das, P. K.; Chattopadhyay, P.; Nethaji, M.; Chakravarty, A. R. J. Am. Chem. Soc. 2003, 125, 12118–12124. (47) Dhar, S.; Senapati, D.; Reddy, P. A. N.; Das, P. K.; Chakravarty, A. R. Chem. Commun. 2003, 2452–2453. (48) Roy, M.; Saha, S.; Patra, A. K.; Nethaj, M.; Chakravarty, A. R. Inorg. Chem. 2007, 46, 4368–4370. (49) Sasmal, P. K.; Patra, A. K.; Nethaji, M.; Chakravarty, A. R. Inorg. Chem. 2007, 46, 11112–11121. (50) Patra, A. K.; Dhar, S.; Nethaji, M.; Chakravarty, A. R. Chem. Commun. 2003, 1562–1563. (51) Herebian, D.; Sheldrick, W. S. J. Chem. Soc., Dalton Trans. 2002, 966–974. (52) (a) Hurley, A. L.; Mohler, D. L. Org. Lett. 2000, 2, 2745–2748. (b) Mohler, D.; Barnhardt, E. K.; Hurley, A. L. J. Org. Chem. 2002, 67, 4982–4984.

DNA CleaVage by Ferrocene Copper(II) Complexes

Organometallics, Vol. 28, No. 5, 2009 1497

Chart 1. Schematic Drawing of the Complexes [Cu(L)(B)](ClO4)2 (B ) bpy, 1; phen, 2; dpq, 3; dppz, 4) and the Heterocyclic Bases Used

Figure 1. Electronic spectra of [Cu(L)(B)](ClO4)2 (B ) bpy, 1; phen, 2; dpq, 3; dppz, 4) in water containing 10% DMF. The arrows indicate the wavelengths of light used for photoinduced DNA cleavage studies. Table 1. Selected Physicochemical Data for the Complexes [Cu(L)(B)](ClO4)2 (B ) bpy, 1; phen, 2; dpq, 3; dppz, 4) IRa/cm-1 [ν(ClO4-)] visible spectral band:b λmax/nm (ε/M-1 cm-1) cyclic voltammetry E1/2/V (∆Ep/mV)c molar conductance ΛMd/S m2 M-1 magnetic moment µeffe/µB

1

2

3

4

1102, 1089 450 (340), 612 (190) 0.535 (75) 0.04 (170) 155

1102, 1088 448 (320), 588 (230) 0.53 (85) -0.02 (140) 161

1090 453 (435), 578 (345) 0.52 (85) -0.02 (120) 150

1101, 1091 441 (620), 560 (345) 0.51 (80) -0.01 (145) 156

1.81

1.79

1.77

1.81

a In KBr phase. b In aqueous DMF (9:1 v/v). The bands near 450 and 600 nm are ferrocene-based (Fc) and Cu(II)-based, respectively. c Fe(III)/Fe(II) and Cu(II)/Cu(I) couple in MeCN-0.1 M TBAP, E1/2 ) 0.5(Epa + Epc), ∆Ep ) (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. Ligand reduction peak: Epc ) -1.49 V for complex 1, Epc ) -1.61V for complex 2, Epc ) -1.19 and -1.39 for complex 3, and E1/2/V (∆Ep/mV) ) -1.16 (120) and -1.69 (220) for complex 4. d Molar conductivity in DMF. e Magnetic moment at 298 K using solid powdered samples of the complexes.

hexafluorophosphate salt, has been structurally characterized by X-ray crystallography. The significant results include photoinduced DNA cleavage activity of the dpq and dppz complexes in red light and the UV-A light induced protein cleavage activity of the dppz complex. A preliminary report on the dppz complex 4 has been made.53

Results and Discussion Synthesis and General Aspects. The ferrocene-conjugated ternary copper(II) complexes [Cu(L)(B)](ClO4)2 (1-4) having heterocyclic bases (B) were prepared in high yield (∼65%) from a reaction of the ferrocenyl tripodal ligand FcCH2N(CH2Py)2 (L) with copper(II) perchlorate hexahydrate and the respective heterocyclic base in methanol. The complexes were characterized by various spectroscopic and analytical techniques (Table 1). All the complexes displayed the mass peak corresponding to [M - ClO4]+ species in their ESI-MS spectra (Figures S1-S4, Supporting Information). The IR spectra showed two characteristic perchlorate anion stretching bands near 1100 cm-1 for complexes 1, 2, and 4 and one broad band for 3. The complexes may have a ClO4- weakly bound to the Cu(II) in (53) Maity, B.; Roy, M.; Chakravarty, A. R. J. Organomet. Chem. 2008, 693, 1395–1399.

the solid state. The complexes were 1:2 electrolytic, showing molar conductance values of 150-161 S m2 M-1 in DMF at 25 °C. The magnetic moment values of ∼1.8 µB at 25 °C suggested the presence of one-electron paramagnetic 3d9-Cu(II) in the complexes having a diamagnetic Fe(II) center in the Fc unit. The UV-visible spectra of 1-4 were recorded in aqueous DMF (10% DMF). The complexes showed a broad and weak copper-centered d-d band within 560-610 nm (Figure 1). A blue shift in the d-d band wavelength was observed for 1 to 4. A very intense ferrocene-centered electronic transition appeared at ∼440 nm.54 The ligand-centered electronic transitions were observed in the UV region of the electronic spectra. Electrochemistry. Cyclic voltammetric experiments were carried out in MeCN-0.1 M TBAP. All the complexes (1-4) showed redox activity (Table 1). The Fe(III)-Fe(II) couple of the ferrocenyl moiety was observed as a reversible response near 0.52 V vs SCE. The conjugation of the {-CH2N(CH2Py)2Cu(B)} unit to Fc has caused a ∼100 mV positive shift of the Fe(III)-Fe(II) potential from 0.42 V of ferrocene ([(η5-C5H5)2Fe]). The complexes also showed a quasireversible cyclic voltammetric response near 0.0 V assignable to the Cu(II)-Cu(I) redox couple (Figure 2). Ligand reduction peaks were observed at -1.16 and -1.69 V for the dppz complex 4 and at -1.19 and -1.39 V for the dpq complex 3. Crystal Structure. The bipyridyl complex as its hexafluorophosphate salt was structurally characterized by single-crystal X-ray diffraction technique. The complex crystallized in the P1j space group in the triclinic crystal system with two molecules in the unit cell. The ORTEP view of the cationic complex is shown in Figure 3. The unit cell packing diagram is shown in Figure S5 (Supporting Information). Selected bond distances and angles data are given in Table 2. The structure consists of a discrete binuclear heterobimetallic complex having Cu(II) and Fe(II). The Fc moiety is covalently linked to CH2N(CH2Py)2, forming the tripodal ligand L, which shows tridentate coordination to copper(II). The five-membered rings in the Fc moiety show an eclipsed conformation. The dihedral angle between the η5-C5H5 and η5-C5H4 rings is 2.67°. The average Fe-C bond distance is 2.038 Å. The copper(II) center has an axially elongated square-pyramidal CuN5 coordination geometry (τ ) (54) Sohn, Y. S.; Hendrikson, D. N.; Gray, H. B. J. Am. Chem. Soc. 1971, 933, 3603–3612.


Maity et al.

Figure 2. Cyclic voltammograms showing the Fe(III)-Fe(II) and Cu(II)-Cu(I) redox processes in the complexes [Cu(L)(B)](ClO4)2 [B ) bpy, 1 (s); phen, 2 (- · -); dpq, 3 ( · · · ); dppz, 4(---)] in MeCN-0.1 M TBAP (scan rate: 50 mV s-1).

Figure 4. (a) Spectral traces showing the effect of gradual addition of CT DNA (360 µM NP) to a 50 µM solution of [Cu(L)(dpq)](ClO4)2 (3) in DMF-Tris-HCl buffer medium. The inset shows the plot of ∆εaf/∆εbf vs [DNA]. (b) Emission spectral traces of BSA (2 µM) in the presence of complex 3 with the inset showing the plot of (I0/I) vs [complex]. Table 3. DNA and BSA Binding Data for the Complexes [Cu(L)(B)](ClO4)2 (B ) bpy, 1; phen, 2; dpq, 3; dppz, 4) 1

2

3

4

3.7 × 104 0.2 1.6 × 104

4.2 ((0.6) × 104 [0.15] 3.1 × 105 2.2 3.8 × 104

1.3 ((0.7) × 105 [0.21] 9.1 × 105 4.4 1.2 × 105

2.5 ((0.4) × 105 [0.2] 1.6 × 106 5.8 2.8 × 105

Kba/M-1 [s] Kappb/M-1 ∆Tmc/°C KBSAd/M-1

a Intrinsic equilibrium DNA binding constant from UV-visible experiment. b Apparent DNA binding constant from ethidium bromide displacement assay. c Change in DNA melting temperature of CT DNA. d BSA (bovine serum albumin) binding constant from fluorescence spectral measurements.

Figure 3. ORTEP view of the cationic complex in [Cu(L)(bpy)](PF6)2 · H2O showing 50% probability thermal ellipsoids and the atom-numbering scheme for the metal and the heteroatoms. The hydrogen atoms are not shown for clarity. Table 2. Selected Bond Distances (Å) and Bond Angles (deg) for [Cu(L)(bpy)](PF6)2 · H2O Cu(1)-N(1) Cu(1)-N(2) Cu(1)-N(3) N(3)-Cu(1)-N(4) N(3)-Cu(1)-N(5) N(4)-Cu(1)-N(5) N(3)-Cu(1)-N(2) N(4)-Cu(1)-N(2)

2.239(4) 2.063(4) 1.994(4) 162.14(15) 82.53(14) 81.92(14) 96.20(15) 95.53(15)

Cu(1)-N(4) Cu(1)-N(5) Fe(1)-C01 [Fe(1)-C02]a N(5)-Cu(1)-N(2) N(3)-Cu(1)-N(1) N(4)-Cu(1)-N(1) N(5)-Cu(1)-N(1) N(2)-Cu(1)-N(1)

1.999(4) 2.062(3) 1.649 [1.657] 160.51(14) 98.02(15) 97.75(14) 122.71(15) 76.76(15)

a C01 and C02 are two centroids of the rings comprising atoms C(24) to C(28) and C(29) to C(33), respectively. The Fe(1)-C distances are in the range 2.028(6) to 2.052(5) Å, giving an average value of 2.038[5] Å.

0.03).55 The 2,2′-bipyridine ligand shows bidentate axialequatorial mode of binding to the copper(II) center. The Cu-N bond distances are in the range 1.994(4) to 2.239(4) Å. There are two lattice PF6 anions and a solvent water molecule in the crystallographic asymmetric unit. Two water molecules in the unit cell are involved in a strong hydrogen-bonding interaction (Figure S5, Supporting Information). The structural features of the ternary bpy complex are similar to those of the reported binary complex [Cu(L)Cl2].56 DNA Binding Property. UV-visible absorption spectral measurements were performed to determine the equilibrium (55) Addison, A. W.; Rao, T. N.; Reedijk, J. V.; Verschoor, G. C. J. Chem. Soc., Dalton Trans. 1984, 1349–1356.

binding constant (Kb) and the binding site size (s) of the complexes to CT DNA by monitoring the change in the absorption intensity of the ligand-centered band at ∼260 nm for the phenanthroline base complexes 2-4. The complexes showed significant hypochromicity at 260 nm with a minor bathochromic shift of ∼2 nm, suggesting mainly the groove binding nature of the complexes to the CT DNA in Tris-HCl buffer medium (Figure 4a, Figure S6 in the Supporting Information). Strong intercalative binding of small molecules to the DNA are known to cause much larger bathochromic shifts and hypochromism of the spectral bands.57 The Kb values of the complexes 2-4 are in the range 4.2 × 104 to 2.5 × 105 M-1 with s (fitting parameter) values of 0.15-0.2 (b.p.), giving an order 4 > 3 > 2 (Table 3). The Kb values of the complexes compare well with other known transition metal complexes of phenanthroline bases.58 The higher DNA binding propensity of the dppz complex than its dpq analogue could be due to the presence of the extended aromatic phenazine ring, which might facilitate partial intercalation of the base through noncovalent π-π interaction with the DNA bases. The binding site size (s) for the complexes is small, and a low value of s (s e 1) suggests a dominant groove binding nature or surface aggregation of the complex on DNA in preference to intercalation.59 The ethidium bromide (EB) displacement assay was used to determine the apparent DNA binding constants (Kapp) of the complexes to CT DNA. The emission intensity of EB was used as a spectral probe, as it is known to show reduced emission (56) Evans, A. J.; Watkins, S. E.; Craig, D. C.; Colbran, S. B. J. Chem. Soc., Dalton Trans. 2002, 983–994. (57) Nair, R. B.; Tang, E. S.; Kirkland, S. L.; Murphy, C. J. Inorg. Chem. 1998, 37, 139–141. (58) Patra, A. K.; Dhar, S.; Nethaji, M.; Chakravarty, A. R. Dalton Trans. 2005, 896–902. (59) Angeles-Boza, A. M.; Bradley, P. M.; Fu, P. K.-L.; Wicke, S. E.; Bacsa, J.; Dunbar, K. R.; Turro, C. Inorg. Chem. 2004, 43, 8510–8519.


Figure 5. (a) DNA melting plots for CT DNA (260 µM NP) alone and in the presence of 50 µM complex [Cu(L)(B)](ClO4)2 (B ) bpy, 1; phen, 2; dpq, 3; dppz, 4) in 5 mM phosphate buffer (pH ) 6.8). (b) Effect of addition of increasing amount of complexes 1 (O), 2 (b), 3 ([), and 4 (2) on the relative viscosity of CT DNA at 37.0 ((0.1) °C in 5 mM Tris-HCl buffer (pH ) 7.2) ([DNA] ) 260 µM, [complex] ) 0-200 µM).

intensity in buffer solution because of solvent quenching, and an enhancement of the emission intensity is observed when EB intercalatively binds to DNA.60,61 The competitive binding of complexes 1-4 to CT DNA could lead to the displacement of the EB, exposing it for solvent quenching of the emission intensity. The binding propensity of the complexes to DNA was measured from the extent of reduction of the EB emission intensity (Figure S7 in the Supporting Information). The Kapp values for 1-4 are 3.7 × 104 to 1.6 × 106 M-1, giving an order 4 > 3 > 2 > 1 (Table 3). The dppz and dpq complexes showed significantly higher Kapp values than the phen complex. The bpy complex is a poor binder to DNA. The DNA melting experiments were carried out to see the effect of DNA duplex stability on binding to the complexes. The duplex DNA at its melting temperature unwinds to give single strand DNA, thus increasing the absorbance at 260 nm. A classical intercalator such as EB stabilizes the duplex DNA, causing the DNA to melt at a higher temperature.62 The present complexes 2-4 gave low values of ∆Tm, ranging from 2.2 to 5.8 °C, indicating primarily DNA groove binding propensity of the complexes (Figure 5, Table 3). The dppz complex with a relatively high ∆Tm value of 5.8 °C could have a partial intercalative mode of DNA binding with the phenazine ring of dppz intercalating to the DNA base pairs. The bpy complex is a poor binder to DNA, giving a ∆Tm value of only 0.2 °C. We have done viscometric titration experiments to determine the relative specific viscosity of CT DNA in the presence of the complexes. A classical intercalator such as EB shows a significant increase in relative viscosity of the DNA solution on intercalation due to an increase in overall DNA contour length on binding to DNA.63,64 In contrast, groove binding or partially intercalating molecules cause little or no effect on the relative viscosity of DNA solution. The plots of relative viscosity (η/η0)1/3 (where η and η0 are the specific viscosity of DNA in the presence and absence of complex, respectively) versus [complex]/[DNA] ratio for 1-4 show only a minor change, indicating groove binding for the phen complex 2 and groove binding along with minor intercalative binding for the dpq (3) and dppz (4) complexes with no apparent binding of the bpy complex (1) to CT DNA (Figure 5). (60) Waring, M. J. J. Mol. Biol. 1965, 13, 269–282. (61) LePecq, J.-B.; Paoletti, C. J. Mol. Biol. 1967, 27, 87–106. (62) Kelly, J. M.; Tossi, A. B.; McConnell, D. J.; OhUigin, C. Nucl. Acid Res. 1985, 13, 6017–6034. (63) Eichhorn, G. L.; Shin, Y. A. J. Am. Chem. Soc. 1968, 90, 7323– 7328. (64) Satyanarayana, S.; Dabrowiak, J. C.; Chaires, J. B. Biochemistry 1992, 31, 9319–9324.


Figure 6. Gel electrophoresis diagram showing the chemical nuclease activity of the complexes [Cu(L)(B)](ClO4)2 (B ) bpy, 1; phen, 2; dpq, 3; dppz, 4) (15 µM) using SC pUC19 DNA (0.2 µg, 30 µM b.p.) in the presence of 500 µM 3-mercaptopropionic acid (MPA) as a reducing and 200 µM H2O2 as oxidizing agent: lane 1, DNA control; lane 2, DNA + MPA; lane 3, DNA + H2O2; lane 4, DNA + 1 + MPA; lane 5, DNA + 2 + MPA; lane 6, DNA + 3 + MPA; lane 7, DNA + 4 + MPA; lane 8, DNA + 1 + H2O2; lane 9, DNA + 2 + H2O2; lane 10, DNA + 3 + H2O2; lane 11, DNA + 4 + H2O2; lane 12, DNA + distamycin (100 µM) + 2 + MPA; lane 13, DNA + distamycin (100 µM) + 3 + MPA; lane 14, DNA + distamycin (100 µM) + 4 + MPA; lane 15, DNA + methyl green (100 µM) + 4 + MPA.

Chemical Nuclease Activity. The DNA cleavage activity of complexes 1-4 (15 µM) was studied in the presence of hydrogen peroxide (H2O2, 200 µM) as an oxidizing and 3-mercaptopropionic acid (MPA, 500 µM) as a reducing agent using supercoiled (SC) pUC19 DNA (0.2 µg, 30 µM) in 50 mM Tris-HCl/50 mM NaCl buffer (pH 7.2). The choice of H2O2 and MPA is based on the observation of Fe(III)-Fe(II) and Cu(II)-Cu(I) redox couples in 1-4 in the cyclic voltammetric studies. While H2O2 is used to oxidize the Fe(II) center of the ferrocenyl moiety, MPA is used to reduce the copper(II) center to the reactive copper(I) species. Complexes 2-4 showed significant chemical nuclease activity in the presence of H2O2 or MPA, while the precursor species, L, and phenanthroline bases alone did not show any cleavage of DNA under similar experimental conditions (Figure 6, Figure S8 in the Supporting Information). H2O2 or MPA controls, in the absence of the complexes, did not show any apparent cleavage of SC DNA. The bpy complex 1 showed poor chemical nuclease activity. The chemical nuclease activity of the complexes in the presence of oxidizing or reducing agent suggests the involvement of both the metal centers in the oxidative DNA cleavage process. The chemical nuclease activity follows the order 4 (dppz) > 3 (dpq) > 2 (phen) . 1 (bpy). The nuclease activity order follows the DNA binding propensity of the complexes. To investigate the mechanistic aspects of the chemical nuclease activity, several additives such as hydroxyl radical scavengers (catalase, DMSO, KI), singlet oxygen quenchers (NaN3, TEMP), and superoxide scavenger (superoxide dismutase, SOD) were used in the DNA cleavage reactions of complex 3 (Figure 7). Hydroxyl radical scavengers were found to inhibit the DNA cleavage activity of 3, while the singlet oxygen quenchers showed no apparent inhibition. SOD also did not show any inhibition of the DNA cleavage. The mechanistic data indicate the involvement of reactive hydroxyl radicals in the DNA cleavage reactions. The DNA groove binding preference of the complexes was determined by using DNA major groove binder methyl green and DNA minor groove binder distamycin. We observed significant inhibition of the chemical nuclease activity of 2 and 3 in the presence of distamycin (100 µM). There was no apparent effect on the chemical nuclease activity of 2 and 3 in the presence of methyl green (100 µM). Complex 4 showed inhibition in the chemical nuclease activity only in the presence of methyl green. The results suggest DNA minor groove binding preference for complexes 2 and 3 and DNA major groove binding preference for the dppz complex 4. Photonuclease Activity. Photoinduced DNA cleavage activity of the complexes was studied using SC pUC19 DNA (30


Maity et al. Table 4. Selected SC pUC19 DNA (0.2 µg, 30 µM) Cleavage Data for the Complexes [Cu(L)(B)](ClO4)2 (B ) bpy, 1; phen, 2; dpq, 3; dppz, 4)

Figure 7. Bar diagram showing the chemical nuclease activity of [Cu(L)(dpq)](ClO4)2 (3) in the presence of different singlet oxygen quenchers and hydroxyl radical scavengers: (a) in the presence of H2O2 and (b) in the presence of MPA.

Figure 8. (a) Gel electrophoresis diagram showing the photoinduced DNA cleavage activity of the complexes [Cu(L)(B)](ClO4)2 (B ) bpy, 1; phen, 2; dpq, 3; dppz, 4) (5 µM) using SC pUC19 DNA (0.2 µg, 30 µM b.p.) on exposure to monochromatic UV-A light of 365 nm for 2 h: lane 1, DNA control (light); lane 2, DNA + 2 (20 µM, dark); lane 3, DNA + 3 (20 µM, dark); lane 4, DNA + 4 (20 µM, dark); lane 5, DNA + 1 (light); lane 6, DNA + 2 (light); lane 7, DNA + 3 (light); lane 8, DNA + 4 (light); lane 9, conversion of NC (extracted after DNA + 4 photocleavage at 365 nm) to SC form on treatment with T4 DNA ligase (4 U). (b) Mechanistic data for complex 3 in UV-A light of 365 nm for an exposure time of 2 h using different additives as singlet oxygen quenchers and hydroxyl radical/superoxide anionic radical scavengers: lane 1, DNA control; lane 2, DNA + 3; lane 3, DNA + DMSO (4 µL) + 3; lane 4, DNA + KI (200 µM) + 3; lane 5, DNA + catalase (2 units) + 3; lane 6, DNA + NaN3 (200 µM) + 3; lane 7, DNA + TEMP (200 µM) + 3; lane 8, DNA + SOD (2 units) + 3.

µM, 0.2 µg) in a medium of Tris-HCl/NaCl (50 mM, pH, 7.2) buffer on irradiation with monochromatic UV-A light of 365 nm (6 W) and visible light of 458, 568, and 647 nm (100 mW). The extent of DNA photocleavage at 365 nm is shown in the gel electrophoresis diagram in Figure 8a. Selected DNA cleavage data are given in Table 4. The DNA photocleavage activity of the complexes in UV-A light follows the order 4 (dppz) > 3 (dpq) . 2 (phen) > 1 (bpy). Complexes 3 and 4 (5 µM), having respectively a photoactive quinoxaline and phenazine moiety, showed efficient DNA cleavage activity (∼90%) at 365 nm for an exposure time of 2 h. The phen complex 2 is a poor photocleaver of SC DNA under similar experimental conditions. This could be due to the photoinactive nature of phen in metal-bound form. The bpy complex 1 did not show any photocleavage of SC DNA.

sl. no.

reaction condition

λa/nm ([complex]/ µM)

% NCb

1 2

DNA control DNA + 1

3

DNA + 2

4

DNA + 3

5

DNA + 4

365 365 (5) 458 (20) 365 (5) 458 (20) 568 (20) 647 (20) 365 (5) 458 (20) 568 (20) 647 (20) 365 (5) 458 (20) 568 (20) 647 (20)

2 6 5 17 23 17 4 88 90 80 59 96 95 81 60

Wavelength used for DNA cleavage, λ ) 365 nm (6W, UV-A), λ ) 458 (30 mW), 568 and 647 nm (CW Ar-Kr laser, 100 mW). Exposure time (t) ) 2 h. b NC is the nicked circular form of DNA obtained from the cleavage of SC DNA. a

Figure 9. Gel electrophoresis diagram showing visible light induced DNA cleavage activity of the complexes [Cu(L)(B)](ClO4)2 (B ) bpy, 1; phen, 2; dpq, 3; dppz, 4) (20 µM) using SC pUC19 DNA (0.2 µg, 30 µM b.p.) for an exposure time of 2 h: lane 1, DNA control (458 nm); lane 2, DNA + 1 (458 nm); lane 3, DNA + 2 (458 nm); lane 4, DNA + 3 (458 nm); lane 5, DNA + 4 (458 nm); lane 6, DNA + 2 (568 nm); lane 7, DNA + 3 (568 nm); lane 8, DNA + 4 (568 nm); lane 9, DNA + 2 (647 nm); lane 10, DNA + 3 (647 nm); lane 11, DNA + 4 (647 nm).

We investigated the visible light induced DNA cleavage activity of 1-4 using light of different wavelengths, namely, 458, 568, and 647 nm, from a tunable continuous-wave (CW) Ar-Kr mixed-gas ion laser (Figure 9, Table 4). The wavelengths chosen for photoirradiations were based on the presence of the metal-centered bands near 450 and 560 nm for the complexes. DNA control experiments using DNA alone showed no apparent photocleavage of DNA under the experimental conditions used. The bpy complex 1 (20 µM) did not show any apparent visible light induced DNA cleavage activity at these wavelengths. The phen complex 2 also showed poor DNA cleavage activity in visible light. Both the dpq (3) and dppz (4) complexes displayed significantly high DNA photocleavage activity in visible light. The cleavage activity follows the order 4 g 3 . 2. Control experiments done using copper(II) perchlorate, dpq, dppz, ferrocene, and L did not show any apparent DNA photocleavage activity under similar reaction conditions. The dpq and dppz complexes showed significant DNA cleavage activity in red light of 647 nm that falls within the PDT spectral window. The complexes 2-4 did not show any DNA cleavage activity in the dark. This excluded the possibility of any hydrolytic cleavage of DNA involving the phosphodiester bond (Figure 8a). We also carried out religation experiments using T4 DNA ligase treated with the nicked circular DNA that was formed as the photoproduct of supercoiled DNA in the presence of the complexes. We did not observe any recombination of NC DNA to its SC form, indicating the oxidative nature of the DNA cleavage (Figure 8a). The mechanistic aspects of the DNA photocleavage reactions were explored using the dpq complex 3 and different additives



Figure 10. Gel electrophoresis diagram showing the mechanistic data for the visible light induced DNA cleavage activity of [Cu(L)(dpq)](ClO4)2 (3, 20 µM) at 458 and 568 nm using SC pUC19 DNA (0.2 µg, 30 µM b.p.) for an exposure time of 2 h: lane 1, DNA control (458 nm); lane 2, DNA + 3 (458 nm); lane 3, DNA + DMSO (4 µL) + 3 (458 nm); lane 4, DNA + KI (200 µM) + 3 (458 nm); lane 5, DNA + catalase (2 units) + 3 (458 nm); lane 6, DNA + NaN3 (200 µM)+ 3 (458 nm); lane 7, DNA + TEMP (200 µM) + 3 (458 nm); lane 8, DNA + 3 (568 nm); lane 9, DNA + DMSO + 3 (568 nm); lane 10, DNA + KI (200 µM) + 3 (568 nm); lane 11, DNA + catalase (2 units) + 3 (568 nm); lane 12, DNA + NaN3 (200 µM) + 3 (568 nm); lane 13, DNA + TEMP (200 µM) + 3 (568 nm).

in UV-A light of 365 nm and visible light of 458 and 568 nm (Figure 8b, Figure 10). The complexes did not show any DNA photocleavage activity under an argon atmosphere, suggesting the involvement of reactive oxygen species (ROS) as the DNAdamaging agent. Singlet oxygen quenchers such as TEMP or NaN3 did not show any inhibitory effect, thus excluding the possibility of a type-II pathway forming singlet oxygen (1O2). Hydroxyl radical scavengers such as KI, DMSO, and catalase showed significant inhibition in the photocleavage activity in both UV and visible light. The mechanistic data suggest the formation of hydroxyl radical via a photoredox pathway.65,66 Such a pathway is known for redox-active copper(II) complexes.47 The visible light induced DNA photocleavage reaction is believed to be metal-assisted involving the visible bands of ferrocene and copper(II).46 Protein Binding. The interaction between bovine serum albumin (BSA) and the complexes 1-4 was investigated by tryptophan emission-quenching experiments. The fluorescence property of BSA in buffer medium is due to the presence of tryptophan residues. The fluorescence intensity primarily depends on the extent of exposure of the two tryptophan residues to the proximity of the immediate polar environment and to quenching groups such as protonated carbonyl, protonated imidazole, and tyrosinate anions through molecular interaction.67 The emission intensity of BSA was found to quench gradually on increasing the complex concentration (Figure 4b, Figure S6 in the Supporting Information). This might be due to changes in the secondary/tertiary structure of BSA in phosphate buffer medium, affecting the orientation of the tryptophan residues of BSA. To understand this spectral change, we initially measured the fluorescent intensity of BSA solution. The fluorescence intensities were subsequently measured on gradual addition of the complex, resulting in reduction of the fluorescence intensity to 50% of its original value. The resulting solution was dialyzed for 12 h to free it from the complex, and then the fluorescence intensity was measured. Dialysis was found to result in partial recovery (∼30%) of the fluorescence intensity of BSA. A control experiment using only BSA solution kept at 4 °C in the absence of the complex showed complete retention of its fluorescence even after 12 h (Figure S11 in the Supporting Information). The protein binding results indicate a partial irreversible conformational change in the protein secondary/tertiary structure (65) Burrows, C. J.; Muller, J. G. Chem. ReV. 1998, 98, 1109–1151. (66) Roy, M.; Pathak, B.; Patra, A. K.; Jemmis, E. D.; Nethaji, M.; Chakravarty, A. R. Inorg. Chem. 2007, 46, 11122–11132. (67) Peters, T. AdV. Protein Chem. 1985, 37, 161–245.

Figure 11. SDS-PAGE diagram showing the photocleavage of bovine serum albumin (BSA, 4 µM) protein in UV-A light of 365 nm by the complexes [Cu(L)(B)](ClO4)2 (B: phen, 2; dpq, 3; dppz, 4) in 50 mM Tris-HCl buffer having 10% DMF (pH 7.2) for an exposure time of 2 h. Panels a-c are for complexes 2-4, respectively: lane 1, molecular marker; lane 2, BSA control; lane 3, BSA + complex (100 µM); lane 4, BSA + complex (150 µM); lane 5, BSA + complex (200 µM); lane 6, BSA + complex (250 µM); lane 7, BSA + complex (250 µM, in dark). Panel d is the MALDI-TOF MS of BSA: (i) BSA and (ii) BSA + 4 after 2 h photoirradiation at 365nm.

of BSA.68 The extent of quenching of the fluorescence intensity of BSA gives the measure of binding propensity of the complexes to BSA. The binding constant for the complexes was determined quantitatively by using the Stern-Volmer equation.69 The linear plot of I0/I vs [complex] gave the binding constant (KBSA). The dppz complex 4 gave the highest KBSA value of 2.8 × 105 M-1 (Table 3). The KBSA values follow the order 4 > 3 > 2. The high binding propensity of the complexes to BSA could be due to the lipophilic character of the ferrocenyl moiety. Complex 4 has a dppz ligand with a phenazine moiety that could have increased the hydrophobic nature of the molecule more than its dpq and phen analogues.70 Photoinduced Protein Cleavage. Dose-dependent photoinduced protease activity of complexes 1-4 against 4 µM BSA in Tris-HCl buffer medium was studied by SDS-PAGE (Figure 11 and Figure S10 in the Supporting Information). The extent of protein photocleavage was compared with the untreated BSA band. Complexes 1-3 were found to be inactive in cleaving BSA on photoexposure to UV-A light (6 W) of 365 nm. We have not observed any fading or smearing of the BSA band for complexes 1-3. We did the control experiments for BSA alone in the dark and in the presence of light. We did not observe apparent cleavage for BSA under these conditions (Figure S10). The dppz complex, however, showed significant smearing or fading of the BSA band on photoirradiation in UV-A light of 365 nm, indicating photocleavage of BSA under physiological reaction conditions. The fading out of the band suggests possible nonspecific binding of the complex to BSA, leading to the cleavage of BSA into very small fragments. A similar type of UV-A light induced photocleavage of BSA has recently been reported by Toshima and co-workers using porphyrin com(68) Ribou, A.-C.; Vigo, J.; Viallet, P.; Salmon, J.-M. Biophys. Chem. 1999, 81, 179–189. (69) Lakowicz, J. R. Principles of Fluorescence Spectroscopy, 2nd ed.; Plenum Press: New York, 1999. (70) Sarkar, B. Biol. Trace Elem. Res. 1989, 21, 137–144.


pounds.71 We performed MALDI-TOF MS analysis for the free BSA (4 µM) and BSA photocleaved by complex 4 (250 µM) (Figure 11d). We observed a significantly intense peak with an m/z value of ∼66 kD, corresponding to BSA alone after UV-A light exposure for 2 h. There was no measurable peak in the mass spectrum when 4 µM of BSA was irradiated with 4 (250 µM) at 365 nm. Complex 4 did not show any protein cleavage activity in the dark, thus ruling out any hydrolytic protein damage. We did not observe any significant protein cleavage activity of the complex in the visible light. The involvement of reactive oxygen species in the cleavage reaction was investigated by carrying out the photoinduced protein cleavage experiments in the presence of singlet oxygen quenchers such as DABCO (3 mM) and NaN3 (3 mM) and hydroxyl radical scavengers such as KI (3 mM) and DMSO (20 µL). DABCO or NaN3 did not show any apparent effect on photoinduced protein cleavage activity of 4 excluding the possibility of a 1O2 cleavage pathway. KI and DMSO showed significant reduction in the protease activity of 4, suggesting the involvement of hydroxyl radicals as the reactive species (Figure S9 in the Supporting Information).

Conclusion We present here the chemistry of ferrocene-conjugated copper(II) complexes showing efficient chemical nuclease activity and photoinduced DNA and protein cleavage activity. The complexes are designed to have a multiutility ferrocenyl moiety conjugated to a copper(II) center that is bound to photoactive and DNA binding phenanthroline bases. The dipyridoquinoxaline (dpq) and dipyridophenazine (dppz) complexes show multifunctional oxidative DNA cleavage activity. The bpy complex 1, as its PF6 salt, has been structurally characterized by X-ray crystallography. The complexes having planar phenanthroline bases show efficient groove binding propensity to the CT DNA. The dppz complex, in addition, shows a partial intercalative mode of binding to DNA. The redox-active complexes show dual chemical nuclease activity in the presence of oxidizing (H2O2) or reducing (MPA) agent involving Fe(II) and Cu(II) centers. The mechanistic pathway involves the formation of hydroxyl radicals as the cleavageactive species. While the dppz complex shows binding at the major groove of DNA, the other complexes display DNA minor groove binding propensity. The dpq and dppz complexes exhibit efficient photoinduced DNA cleavage activity in UV-A and visible light. The oxidative DNA cleavage occurs via a metalassisted photoexcitation process forming hydroxyl radicals in a photoredox pathway. The dpq and dppz complexes show significant red light induced DNA cleavage activity at 647 nm within the photodynamic therapy spectral window. The complexes having phenanthroline bases show efficient binding to bovine serum albumin protein. The dppz complex 4 exhibits significant photocleavage of BSA in UV-A light of 365 nm. While complexes 3 and 4 are excellent model photonucleases in both UV-A and visible light, complex 4 is a model photoprotease in UV-A light. The results presented here on 3dmetal-based bioorganometallic complexes are unprecedented in the chemistry of PDT. This work is expected to presage further development on designing new photoactive 3d-metal-based bioorganometallic complexes as nucleolytic and antimetastasis agents for potential cellular applications in PDT. (71) (a) Tanimoto, S.; Matsumura, S.; Toshima, K. Chem. Commun. 2008, 3678–3680. (b) Suzuki, A.; Hasegawa, M.; Ishii, M.; Matsumura, S.; Toshima, K. Bioorg. Med. Chem. Lett. 2005, 15, 4624–4627.

Maity et al.

Experimental Section Materials. The reagents and chemicals were obtained from commercial sources (Sigma-Aldrich, USA; SD Fine Chemicals, India). The supercoiled (SC) pUC19 DNA (cesium chloride purified), T4 ligase, 5X ligation buffer, and protein molecular weight marker were procured from Bangalore Genie (India). Ferrocene carboxaldehyde, 2-picolylamine, calf thymus (CT) DNA, agarose (molecular biology grade), acrylamide, N,N′-methylenebisacrylamide, distamycin, methyl green, catalase, superoxide dismutase, bovine serum albumin (BSA), and ethidium bromide (EB) were purchased from Sigma-Aldrich (USA). Tris(hydroxymethyl)aminomethane-HCl (Tris-HCl) buffer was prepared using deionized and sonicated triple-distilled water. Ferrocenylmethylbis(2-pyridylmethylamine) (L), dipyrido[3,2-d:2′,3′-f]quinoxaline (dpq), and dipyrido[3,2-a:2′,3′-c]phenazine (dppz) were prepared following literature procedures.56,72,73 The solvents were purified by standard literature procedures.74 General Methods. The elemental analysis was done using a Thermo Finnigan FLASH EA 1112 CHNS analyzer. The infrared, electronic, and fluorescence spectra were recorded on Perkin-Elmer Lambda 35, Perkin-Elmer spectrum one 55, and Perkin-Elmer LS 50B spectrophotometers, respectively. Magnetic susceptibility data at 298 K for the polycrystalline samples of the complexes were obtained using a model 300 Lewis-coil force magnetometer, George Associates Inc. (Berkeley, CA). Hg[Co(NCS)4] was used as a standard. Diamagnetic corrections were done on the experimental susceptibility data.75 Molar conductivity measurement was done using a Control Dynamics (India) conductivity meter. Cyclic voltammetric measurements were made at 25 °C on an EG&G PAR 253 VersaStat potentiostat/galvanostat using a three-electrode configuration consisting of a glassy carbon working, a platinum wire auxiliary, and a saturated calomel reference (SCE) electrode. Ferrocene (E1/2 ) 0.42 V) was used as a standard in MeCN/ 0.1 M [Bun4N](ClO4) (TBAP). DNA binding studies by UV-visible and DNA melting method were done using a Cary 300 Bio UV-visible spectrometer with a Cary temperature controller. Electrospray ionization mass spectral measurements were done using a Bruker Daltonics Esquire 300 Plus ESI Model mass spectrometer. MALDI-TOF mass spectra were recorded using Bruker Daltonics Ultraflex MALDI-TOF instrument. Synthesis of [Cu(L)(B)](ClO4)2 (B ) bpy, 1; phen, 2; dpq, 3; dppz, 4). Complexes 1-4 were prepared by following a general synthetic procedure in which a methanolic solution (7 mL) of [Cu(H2O)6](ClO4)2 (0.18 g, 0.5 mmol) was initially reacted with a methanolic solution (7 mL) of ferrocenylmethylbis(2-pyridylmethylamine) (L, 0.2 g, 0.5 mmol), and the resulting solution was stirred for 1 h under a dinitrogen inert atmosphere followed by cooling in an ice bath. To the cold solution was added a methanolic solution (10 mL) of the heterocyclic base B (bpy, 0.078 g; phen, 0.1 g; dpq, 0.12 g; dppz, 0.14 g; 0.5 mmol) with continuous stirring for 45 min. The solid thus obtained was isolated and washed with cold diethyl ether-methanol mixture until the washings were no longer colored, and the product was finally dried under vacuum over P4O10 (Yield: 284 mg, 70% for 1; 273 mg, 65% for 2; 298 mg, 67% for 3; 301 mg, 64% for 4). Anal. Calcd for C33H31Cl2CuFeN5O8 (1): C, 48.54; H, 3.88; N, 8.58. Found: C, 48.25; H, 3.53; N, 8.34. FT-IR (KBr phase): 3495w(br), 3078w(br), 1613m, 1488w, 1442s, 1353w, 1320w, 1277w, 1102vs (ClO4-), 1089vs(sh) (ClO4-), 1000w, 830m, 768s, 623s, 545w, 477w, 422w cm-1 (vs, very strong; s, strong; m, medium; w, weak; br, broad; sh, shoulder). ESI-MS in MeCN: m/z 716 (M - ClO4)+. UV-visible (72) Collins, J. G.; Sleeman, A. D.; Aldrich, J. R.; Greguric, I.; Hambly, T. W. Inorg. Chem. 1998, 37, 3133–3141. (73) Amouyal, E.; Homsi, A.; Chambron, J.-C.; Sauvage, J.-P. J. Chem. Soc., Dalton Trans. 1990, 1841–1845. (74) Perrin, D. D.; Armarego, W. L. F.; Perrin, D. R. Purification of Laboratory Chemicals; Pergamon Press: Oxford, 1980. (75) Khan, O. Molecular Magnetism; VCH: Weinheim, 1993.

DNA CleaVage by Ferrocene Copper(II) Complexes in aqueous DMF (10% DMF) [λmax, nm (ε, M-1 cm-1)]: 254 (19 800), 281 (15 100), 309 (7150), 450 (340), 612 (190). ΛM, S m2 M-1 in DMF at 25 °C: 155. µeff ) 1.81 µB at 298 K. Anal. Calcd for C35H31Cl2CuFeN5O8 (2): C, 50.05; H, 3.72; N, 8.34. Found: C, 49.87; H, 3.55; N, 8.43. FT-IR (KBr phase): 3470w(br), 3070w(br), 1613m, 1519w, 1483w, 1429m, 1345w, 1314w, 1282w, 1102vs (ClO4-), 1088vs (ClO4-), 993w, 852m, 768m, 729m, 623s, 472w, 421w cm-1. ESI-MS in MeCN: m/z 739 (M - ClO4)+. UV-visible in aqueous DMF (10% DMF) [λmax, nm (ε, M-1 cm-1)]: 265 (39 500), 292 (12 900), 448 (320), 588 (230). ΛM, S m2 M-1 in DMF at 25 °C: 161. µeff ) 1.79 µB at 298 K. Anal. Calcd for C39H31Cl2CuFeN7O8 (3): C, 49.82; H, 3.50; N, 10.99. Found: C, 49.62; H, 3.71; N, 10.78. FT-IR (KBr phase): 3436w(br), 3079br, 1612m, 1580w, 1529w, 1483m, 1447m, 1406m, 1345w, 1309w, 1278w, 1090vs (ClO4-), 998w, 815m, 763m, 736m, 623s, 545w, 488w, 442w cm-1. ESI-MS in MeCN: m/z 793 (M - ClO4)+. UV-visible in aqueous DMF (10% DMF) [λmax, nm (ε, M-1 cm-1)]: 253 (54 200), 293 (18 200), 324 (8300), 338 (6880), 453 (435), 578 (345). ΛM, S m2 M-1 in DMF at 25 °C: 150. µeff ) 1.77 µB at 298 K. Anal. Calcd for C41H33Cl2CuFeN7O8 (4): C, 52.28; H, 3.53; N, 10.41. Found: C, 51.92; H, 3.36; N, 10.30. FT-IR (KBr phase): 3450w(br), 3079w(br), 1610m, 1496m, 1447w, 1416w, 1360w, 1284w, 1101vs(sh) (ClO4-), 1091vs (ClO4-), 1028w, 819w, 766m, 732w, 623s, 421w cm-1. ESI-MS in MeCN: m/z 841 (M - ClO4)+. UV-visible in aqueous DMF (10% DMF) [λmax, nm (ε, M-1 cm-1)]: 266 (47 100), 361 (10 000), 381 (9700), 441 (620), 560 (345). ΛM, S m2 M-1 in DMF at 25 °C: 156. µeff ) 1.81 µB at 298 K. Solubility and Stability. The complexes were highly soluble in DMSO, DMF, and MeCN and moderately soluble in MeOH and CH2Cl2. The complexes were soluble and stable in aqueous DMF. CAUTION! The perchlorate salts are potentially explosiVe; only a small quantity of the complex was handled with precautions. X-ray Crystallographic Procedure. The crystal structure of [Cu(L)(bpy)](PF6)2 · H2O was obtained by single-crystal X-ray diffraction technique. Single crystals of the complex were grown by a diffusion technique in which hexane was layered over the dichloromethane solution of the complex. The source of solvent water could be the commercially available solvents used for crystallization. A bluish-green needle-shaped crystal was mounted on a glass fiber using epoxy cement. X-ray diffraction data were measured in frames with increasing ω (width of 0.3 deg per frame) and with a scan speed at 15 s/frame using a Bruker SMART APEX CCD diffractometer, equipped with a fine focus 1.75 kW sealedtube X-ray source. Empirical absorption corrections were made using a multiscan program.76 The structure was solved by heavyatom methods and refined by full matrix least-squares using the SHELX system of programs.77 All non-hydrogen atoms were refined anisotropically. All hydrogen atoms belonging to the complex were in their calculated positions and refined using a riding model. The perspective view of the complex was obtained using the ORTEP program.78 Selected crystallographic data are summarized in Table 5. DNA Binding Experiments. The experiments were carried out in 5 mM Tris-HCl buffer (pH 7.2) at ambient temperature. The DNA melting experiment was performed using phosphate buffer (pH 6.8). The ratio of the absorbance values of CT DNA at 260 and 280 nm in Tris-HCl buffer was found to be 1.9:1, indicating DNA free from any protein impurities. The DNA concentration in base pair was determined by absorption spectroscopy using the molar absorption coefficient of 6600 M-1 cm-1 at 260 nm for CT DNA.79 In the UV-visible absorption titration experiment, the complex solution (50 µM) in 5 mM Tris-HCl buffer containing 5% DMF (76) Walker, N.; Stuart, D. Acta Crystallogr. 1983, 39, 58–166. (77) Sheldrick, G. M. SHELX-97, Programs for Crystal Structure Solution and Refinement; University of Go¨ttingen: Go¨ttingen, Germany. (78) Johnson, C. K. ORTEP-III, Report ORNL-5138; Oak Ridge National Laboratory: Oak Ridge, TN, 1976.

Organometallics, Vol. 28, No. 5, 2009 1503 Table 5. Selected Crystallographic Data for [Cu(L)(bpy)](PF6)2 · H2O empirical formula fw, g M-1 cryst syst space group a, Å b, Å c, Å R, deg β, deg γ, deg V, Å3 Z T, K Fcalcd, g cm-3 λ, Å (Mo KR) µ, cm-1 data/restraints/params F(000) goodness-of-fit R (Fo)a, I > 2σ(I) [wR (Fo)b] R (all data) [wR (all data)] largest diff peak and hole (e Å-3)

C33H33N5OP2F12CuFe 924.97 triclinic P1j 9.093(2) 11.438(3) 18.614(5) 105.532(17) 91.927(15) 93.493(17) 1859.3(9) 2 293(2) 1.652 0.71073 11.44 7293/0/496 922 1.064 0.0563 [0.1431] 0.0952 [0.1674] 0.752, -0.546

a R ) ∑|Fo|-|Fc|/∑|Fo|. b wR ) {∑[w(Fo2 - Fc2)2]/∑[w(Fo)2]}1/2; w ) [σ2(Fo)2 + (AP)2 + BP]-1, where P ) (Fo2 + 2Fc2)/3, A ) 0.0817; B ) 0.7755.

was titrated with 360 µM CT DNA. The band at ∼260 nm was monitored for the complexes during the absorption titration experiment. The concentration of the CT DNA was varied within 20-500 µM. 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 CT DNA. The intrinsic equilibrium binding constant (Kb) and the binding site size (s, b.p.) of the complexes to CT DNA were obtained by the McGhee-von Hippel (MvH) method using the expression of Bard and co-workers: ∆εaf/∆εbf ) (b - (b2 - 2Kb2Ct[DNA]/s)1/2)/2Kb, where b ) 1 + KbCt + Kb[DNA]/2s; Kb is the microscopic binding constant for each site; Ct is the total concentration of the metal complex; s is the binding site size (in base pairs) of the metal complex interacting with the DNA; εf, εa, and εb are respectively the molar extinction coefficients of the free complex in solution, complex bound to DNA at a definite concentration, and the complex in completely bound form with CT DNA.80,81 The nonlinear least-squares regression analysis was done using Origin Laboratory, version 6.1. The fluorescence spectral measurements were done using ethidium bromide (EB) bound CT DNA solution (260 µM) in 5 mM Tris-HCl buffer (pH 7.2) at 25 °C. EB itself did not show any fluorescence in Tris-HCl buffer due to fluorescence quenching by the solvent. It showed enhanced fluorescence emission due to its intercalation to the CT DNA that made EB inaccessible to the solvent molecules. The fluorescence intensities of EB-bound CT DNA at 601 nm with increasing concentration of the complex were recorded. The addition of metal complex to CT DNA could result in the competitive displacement of EB, and hence there was a decrease in the emission intensity. The apparent binding constants for the complexes, Kapp, were determined by using the equation Kapp × [complex] ) KEB × [EB], where [complex] is the concentration of the complex at 50% reduction of the fluorescence intensity, KEB ) 1.0 × 107 M-1, and [EB] ) 1.3 µM.82 DNA thermal denaturation studies were carried out by monitoring the absorption intensity of CT DNA (260 µM) at 260 nm varying the (79) Reichman, M. E.; Rice, S. A.; Thomas, C. A.; Doty, P. J. Am. Chem. Soc. 1954, 76, 3047–3053. (80) McGhee, J. D.; Von Hippel, P. H. J. Mol. Biol. 1974, 86, 469– 489. (81) Carter, M. T.; Rodriguez, M.; Bard, A. J. J. Am. Chem. Soc. 1989, 111, 8901–8911. (82) Lee, M.; Rhodes, A. L.; Wyatt, M. D.; Forrow, S.; Hartley, J. A. Biochemistry 1993, 32, 4237–4245.

1504 Organometallics, Vol. 28, No. 5, 2009 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 in 5 mM phosphate buffer (pH 6.8). The DNA melting experiment was carried out with the complex to a CT DNA molar ratio of 1:5.2. 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 260 µM. The flow time was measured with an automated timer. The data were 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 CT DNA containing solutions (t) corrected for that of the buffer alone (t0), η ) (t - t0). DNA Cleavage Experiments. The photoinduced cleavage of SC pUC19 DNA (0.2 µg, 30 µM, 2686 base pairs) by the complexes was studied by agarose gel electrophoresis. The reactions were carried out under illuminated conditions using a UV-A lamp of 365 nm (6 W, sample area of illumination ) 45 mm2) and visible light of 568 and 647 nm using a CW Ar-Kr laser (100 mW, laser beam diameter ) 1.8 mm, beam divergence ) 0.70 mrad, Spectra Physics water-cooled mixed-gas ion laser stabilite 2018-RM). In addition, the DNA photocleavage experiment was performed at 458 nm using the same laser source but with a laser power of 30 mW. The power of the laser beam was measured using a Spectra Physics CW laser power meter (model 407A). Eppendorf and glass vials were used for respective UV and visible light experiments in a dark room at 25 °C using SC DNA (1 µL, 30 µM) in 50 mM tris(hydroxymethyl)methane-HCl (Tris-HCl) buffer (pH 7.2) containing 50 mM NaCl and the complex (2 µL). The DNA cleavage reaction was performed in the presence of various external additives such as KI (200 µM), NaN3 (200 µM), TEMP (200 µM), DMSO (4 µL), catalase (2 units), and SOD (2 units) for the mechanistic investigation. The concentration of the complexes in DMF or the additives in buffer corresponded to the quantity in 2 µL stock solution used prior to dilution to the 20 µL final volume using 50 mM Tris-HCl buffer. The solution path length used for illumination in the sample vial was ∼5 mm. The reaction mixture was incubated for 1 h at 37 °C prior to the photoexposure. After the photoexposure, loading dye containing 25% bromophenol blue, 0.25% xylene cynol, and 30% glycerol (3 µL) was added to the reaction mixture, and the solution was finally loaded on 1% agarose gel containing 1.0 µg mL-1 ethidium bromide (EB). The electrophoresis was carried out in a dark room for 2 h at 45 V in 1X TAE (Tris-acetate-EDTA) buffer. The bands were visualized by UV light and photographed. The extent of cleavage of SC DNA was determined by measuring the intensities of the bands using a UVITECH gel documentation system. Corrections were made for the low level of nicked circular (NC) form present in the original supercoiled (SC) DNA sample and for the low affinity of EB binding to SC compared to NC form of DNA.83 The chemical nuclease studies on the complexes were carried out using hydrogen peroxide (200 µM) as an oxidizing agent and 3-mercaptopropionic acid (500 µM) as a reducing agent, complexes of 15 µM, and SC pUC19 DNA of 30 µM under dark reaction conditions. The observed error in measuring the band intensities was ∼5%. The religation experiment for complex 4 was carried out using T4 DNA ligase. The NC form obtained from the photoinduced DNA cleavage was removed from the agarose gel using a gel extraction kit with subsequent addition of 5× ligation buffer and T4 DNA ligase (1 µL, 4 U). The solution was incubated for 10 h at 16 °C before subjecting it to gel electrophoresis.84 (83) Bernadou, J.; Pratviel, G.; Bennis, F.; Girardet, M.; Meunier, B. Biochemistry 1989, 28, 7268–7275. (84) Gupta, T.; Dhar, S.; Nethaji, M.; Chakravarty, A. R. Dalton Trans. 2004, 1896–1900.

Maity et al. BSA Binding Experiments. The protein binding study was performed by tryptophan fluorescence quenching experiments using bovine serum albumin (BSA, 2 µM) as the substrate in phosphate buffer (pH 6.8). Quenching of the emission intensity of tryptophan residues of BSA at 344 nm (excitation wavelength at 295 nm) was monitored using complexes 1-4 as quenchers with increasing complex concentration.85 Stern-Volmer I0/I vs [complex] plots using the corrected fluorescence data taking into account the effect of dilution were done. Linear fit of the Stern-Volmer equation (I0/I ) 1 + KBSA[Q]), where I0 and I are the emission intensity of BSA in the absence of quencher and emission intensity of BSA in the presence of the quencher of concentration [Q], gave the binding constant (KBSA) values using Origin 6.1. The fluorescence intensity of BSA (5 µM) as well as BSA in the presence of complex 4 (5 µM) was measured. The equilibrium dialysis (12 h) of the BSA solution containing the complex was done to free BSA from the complex. The fluorescence intensity was subsequently measured. Dialysis tubing with a cutoff of 3 kD molecular mass was used for dialysis after heating it three times in water. BSA Cleavage Experiments. Photoinduced protein cleavage experiments were carried out according to the literature procedure described by Kumar and co-workers.86 Freshly prepared solution of BSA (MW 66 600) in 50 mM Tris-HCl buffer (pH 7.2) was used for the photochemical protein cleavage studies. The protein solutions (4 µM) in Tris-HCl buffer medium containing complexes 1-4 with varied concentration from 100 to 250 µM were photoirradiated at 365 nm (6 W) for 2 h. Eppendorf and glass vials were used for the UV-A and visible light induced protein cleavage studies, respectively. The visible light induced protein cleavage activity of complex 4 was studied using the CW Ar-Kr laser with wavelengths of 458 nm (30 mW), 568 nm (50 mW), and 647 nm (50 mW). The BSA solution in the presence of the complexes was incubated at 37 °C for 1 h prior to the photoexposure. Protein samples were evaporated under vacuum by using an EYELA centrifugal vaporizer (model CVE-200D) for gel electrophoresis experiments. The irradiated samples (50 µL) were dried in a centrifugal vaporizer, and the samples were dissolved in the loading buffer (24 µL) containing SDS (7% w/v), glycerol (4% w/v), TrisHCl buffer (50 mM, pH 6.8), mercaptoethanol (2% v/v), and bromophenol blue (0.01% w/v). The protein solutions were then denatured on heating to a boil for 3 min. The polyacrylamide gel was prepared by following a literature procedure.87 Samples were loaded on a 3% polyacrylamide (stacking) gel. The gel electrophoresis was done at 60 V until the dye passed into the separating gel from the stacking (3%) gel, and then the voltage was increased to 120 V. The gels were run for 1.5 h, stained with Coomassie Brilliant Blue R-250 solution (acetic acid/methanol/water ) 1:2:7 v/v), and destained with a water/methanol/acetic acid mixture (5: 4:1 v/v) for 4 h. The gels, after destaining, were scanned with a HP Scanjet G3010 scanner, and the images were further processed using Adobe Photoshop 7.0 software package. Molecular weight markers were used in each gel to calibrate the molecular weights of the BSA. The presence of reactive oxygen species was investigated by carrying out the photoinduced protein cleavage experiments. Various singlet oxygen quenchers such as DABCO (3 mM) and NaN3 (3 mM) and hydroxyl radical scavengers such as KI (3 mM) and DMSO (20 µL) were used for mechanistic studies.

Acknowledgment. We thank the Department of Science and Technology (DST), Government of India, and the Council of Scientific and Industrial Research (CSIR), New (85) Hua, Y. J.; Liua, Y.; Wangb, J. B.; Xiaob, X. H.; Qua, S. S. J. Pharm. Biomed. Anal. 2004, 36, 915–919. (86) Kumar, C. V.; Buranaprapuk, A.; Sze, H. C.; Jockusch, S.; Turro, N. J. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 5810–5815. (87) Scha¨gger, H.; von Jagow, G. Anal. Biochem. 1987, 166, 368–379.


Delhi, for financial support [SR/S5/MBD-02/2007 and 01(2081)/06/EMR-II]. We are thankful to DST for the CCD diffractometer facility and the Alexander von Humboldt Foundation, Germany, for donation of an electroanalytical system. S.S. is thankful to the CSIR, New Delhi, for a research fellowship. A.R.C. thanks DST for a J. C. Bose National Fellowship. Supporting Information Available: CIF file giving crystallographic data for the bpy complex and figures showing mass

Organometallics, Vol. 28, No. 5, 2009 1505 spectra of the complexes (Figures S1-S4), unit cell packing diagram (Figure S5), UV-visible DNA binding and fluorescence BSA binding (Figure S6), ethidium bromide displacement assay (Figure S7), gel electrophoresis diagrams on DNA cleavage experiments (Figure S8), mechanistic data on protein cleavage (Figure S9), protein cleavage (Figure S10), and protein dialysis data (Figure S11). This material is available free of charge via the Internet at http://pubs.acs.org. OM801036F

Photoinduced DNA and Protein Cleavage Activity of Ferrocene

Recommend Documents