Synthesis, characterization, and DNA binding and cleavage properties of copper(II)-tryptophanphenylalanine-1,10-phenanthroline/2,2′-bipyridine complexes

Article abstract of DOI:10.1002/cbdv.200900388

The mononuclear dipeptide-based Cu^II complexes [Cu ^II(trp-phe)(phen)(H₂O)] ClO₄ (1) and [Cu^II(trp-phe)(bpy)(H₂O)] ClO₄ (2) (trp-phe=tryptophanphenylalanine, phen=1,10-phenanthrol

Full text of DOI:10.1002/cbdv.200900388

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Synthesis, Characterization, and DNA Binding and Cleavage Properties of
Copper(II)-tryptophanphenylalanine-1,10-phenanthroline/2,2’-bipyridine
Complexes
by Pulimamidi R. Reddy*, Nomula Raju, and Battu Satyanarayana
Department of Chemistry, Osmania University, Hyderabad – 500007, India
(phone: þ91-40-27171664; fax: þ91-40-27090020; e-mail: rabi_pr@rediffmail.com)
The mononuclear dipeptide-based CuII complexes [CuII(trp-phe)(phen)(H2O)] · ClO4 (1) and
[CuII(trp-phe)(bpy)(H2O)] · ClO4 (2) (trp-phe¼tryptophanphenylalanine, phen¼1,10-phenanthroline,
bpy¼2,2’-bipyridine) were isolated, and their interaction with DNA was studied. They exhibit
intercalative mode of interaction with DNA. The intercalative interaction was quantified by
SternꢀVolmer quenching constant (Ksq ¼0.14 for 1 and 0.08 for 2). The CuII complexes convert
supercoiled plasmid DNA into its nicked circular form hydrolytically at physiological conditions at a
concentration as low as 5 mm (for 1) and 10 mm (for 2). The DNA hydrolysis rates at a complex
concentration of 50 mm were determined as 1.74 hꢀ1 (R¼0.985) for 1 and 0.65 hꢀ1 (R¼0.965) for 2. The
rate enhancement in the range of 2.40–4.10ꢁ107-fold compared to non-catalyzed double-stranded DNA
is significant. This was attributed to the presence of a H2O molecule in the axial position of the Cu
complexes.
Introduction. – During the last decade, the interaction between transition metal
complexes and DNA has received much attention [1–14], since binding studies of small
molecules to DNA are important in the development of new therapeutic reagents and
DNA probes [15–19]. CuIIꢀPeptideꢀdiimine complexes were shown to bind to DNA in
a nonspecific (noncovalent) mode of interaction such as electrostatic, groove [20][21],
intercalative, and partial intercalative binding [22]. Synthesis, structure, and nuclease
properties of ternary CuII-peptides with 1,10-phenanthroline were reported and
structurally characterized by X-ray crystallography [23]. Most of the applications of
these complexes require that the complexes bind to DNA through an intercalative
mode. Since aromatic moieties are known to play an important role in DNA binding
through intercalation, we have isolated a peptide, trp-phe, with aromatic side chains
and synthesized its CuII complexes with phen and bpy ligands. The DNA-binding
properties of these CuII complexes were explored by spectroscopic and viscosity
measurements. Their catalytic activity towards pUC19 DNA was also investigated. The
rate of hydrolysis of DNA catalyzed by the complexes was determined by kinetics
experiments. The rate enhancement in the range of 2.40–4.10ꢁ107-fold compared to
non-catalyzed double-stranded DNA is impressive.
Results and Discussion. – Synthesis and Solubility. The synthesis of 1 and 2 is
schematically presented in Scheme 1. The complexes are partially soluble in MeOH and
EtOH, but they show good solubility in aqueous MeOH/EtOH. They were non-
ꢀ 2011 Verlag Helvetica Chimica Acta AG, Zꢁrich

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hygroscopic and stable in both solid and solution phases. The analytical data for the CuII
complexes are in good agreement with the molecular formulae of the complexes. The
molar conductance values are too low to account for any dissociation of the complexes
in MeOH, indicating the non-electrolytic nature of the complexes (Table 1).
Scheme 1. Syntheses of Complexes 1 and 2
Table 1. Physicochemical Data for Complexes 1 and 2
Complex
meff (BM) ESI-MS UV/VIS
[nm]
LM
[Wꢀ1 cm2 molꢀ1
]
[CuII(trp-phe)(phen)(H2O)]·ClO4 (1) 2.01
[CuII(trp-phe)(bpy)(H2O)]·ClO4 (2)
1.69
593
587
206, 222, 268, 626 1.60
244, 298, 310, 613 2.10
In spite of trying different conditions and solvents, we could not succeed in isolating
single crystals. However, the complexes were fully characterized. The geometries were
determined by UV/VIS and EPR studies. The molecular-modeling structures were also
obtained to determine the bond distances. They are comparable with those obtained by
X-ray crystallography for the analogous [Cu(gly-trp)(phen)] system [23].
Characterization of CuIIꢀtrp-pheꢀphen/bpy Complexes. In the IR spectra of free
dipeptide, the NꢀH and C¼O stretching vibrations were observed at 3316 and 1684 cmꢀ1
respectively, the NꢀH stretching was merged with OꢀH stretching vibration of H2O
molecule and appeared as a broad band at 3420 cmꢀ1 for 1, 3446 cmꢀ1 for 2 (NH2 þ

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133
H2O), and the C¼O band was shifted to 1654 cmꢀ1 upon complexation in both the
complexes. The nas (COOꢀ) and ns (COOꢀ) stretching vibrations for 1 and 2 were
observed at 1654 and 1384 cmꢀ1, respectively. The large difference between the two
values indicates that the COOꢀ groups are coordinated to the metal ion in a
monodentate fashion [24]. In phen, the peaks corresponding to the ring stretching
frequencies (n(C¼C) and n(C¼N)) at 1502 and 1421 cmꢀ1 were shifted to higher
frequencies (ca. 1518 and 1426 cmꢀ1, resp.); in bpy, the peaks corresponding to the ring
stretching frequencies (n(C¼C) and n(C¼N)) at 1557 and 1452 cmꢀ1 were shifted to
1603 and 1474 cmꢀ1, respectively, indicating the coordination of the heterocyclic N-
atoms to the Cu ion for 1 and 2, respectively. The characteristic out-of-plane H-bending
modes of free phen observed at 853 and 738 cmꢀ1 (for bpy at 780 and 714 cmꢀ1) were
shifted to 848 and 722 cmꢀ1 (for bpy 768 and 703 cmꢀ1), respectively, upon metal
complexation. The very strong bands at 1086 cmꢀ1 for 1 and 1094 cmꢀ1 for 2 were
assigned to n(ClꢀO) of ClOꢀ4 . The rocking and wagging modes of MꢀOH2 were
observed at 920 cmꢀ1 and 623 cmꢀ1, respectively, for both the complexes [25]. The new
non-ligand peaks at 454 and 512 cmꢀ1 were assigned to n(CuꢀO) and n(CuꢀN)
stretching vibrations, respectively [26]. These values are comparable with those
observed for analogous CuII complexes [23]. However, the slight variations may be
attributed to the presence of an aromatic side chain in trp-phe.
The observed magnetic moment values (meff ¼2.01 and 1.69 BM for 1 and 2, resp.)
are indicative of paramagnetic nature of Cu complexes with spin (1/2) system [27].
The electronic spectra for the CuII complexes were recorded in MeOH. Four
absorption bands with varied intensity were observed for 1 (Fig. 1) and 2 (Fig. S1)1).
The intense bands at 206 and 222 nm of 1, and 244 and 298 nm of 2 were assigned to
intra-ligand (pꢀp*) transition of phen and bpy ligands, respectively, while the less
intense bands at 268 nm of 1 and at 310 nm of 2 were assigned to the LMCT transitions.
Besides, a broad band observed at 626 nm for 1 and 613 nm for 2 in the spectra matches
2
2
well with the reported B1g! B2gꢀ(n1) ðdx2ꢀy2 ! dyzÞ transition of the five-coordi-
nated CuII in an approximate square-pyramidal geometry [28]. They agree well with the
reported analogous systems [23].
The active chemical species of the complexes were identified by means of
electrospray ionization mass spectrometry (ESI-MS). The peaks observed at m/z 593
and 587 were due to the formation of species [Cu(trp-phe)(phen)]þ for 1 and [Cu(trp-
phe)(bpy)]þNaþ for 2.
The solid-state EPR spectra of 1 and 2 at room temperature was indicative of S¼
1/2, which also showed four g-tensor viz. g1, g2, g3, and g4 values at 2.04, 2.09, 2.17, and
2.45, respectively, for 1, and 2.08, 2.15, 2.31, and 2.45, respectively for 2, where g4 >g3 >
g2 >g1 (Table 2). It was observed earlier that if the value of g4 is higher than 2.3, it is
indicative of covalent character of the MꢀL bonds [29].
The EPR spectra of the complexes provided good evidence for distinguishing the
two ground states of five-coordinated structure, i.e., dx2 ꢀ dy2 (square-pyramidal) and
dz2 (trigonal-bipyramidal). For the system with g3 >g2 > g1, the ratio of (g2 ꢀg1)/(g3 ꢀ
g2), hereafter called parameter R, will give information regarding the ground-state
orbital. If R>1, predominantly the ground-state is dz2, and if R<1, the ground-state will
1
)
Supplementary material (Figs. S1–S4) for complex 2 is available upon request from the authors.

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Fig. 1. UV/VIS Absorption spectra of complex 1 (inset: d!d transition).
Table 2. EPR Parameters of Complexes 1 and 2 (at room temperature)
Complex
g1
g2
g3
g4
Ra)
[CuII(trp-phe)(phen)(H2O)]·ClO4 (1)
[CuII(trp-phe)(bpy)(H2O)]·ClO4 (2)
2.04
2.08
2.09
2.15
2.17
2.31
2.45
2.45
0.625
0.437
a) R¼(g2 –g1)/(g3 –g2).
be dx2 ꢀ dy2 , which corresponds to square-pyramidal structure. The R (R<1) values for
1 and 2 (Table 2) indicate a dx2 ꢀ dy2 ground state, which is consistent with a distorted
square-pyramidal structure [30].
The thermal decomposition stoichiometries of the analytically characterized
complexes were determined by TGA (thermogravimetric analyses). The spectra
showed no mass loss in the temperature range of 25–1808. However, in the range of
180–2408, a mass loss of 3.2% (calc. 3.1%; 18 mass units) was observed. This is due to
the loss of one coordinated H2O molecule from 1 and 2.
The minimum-energy conformers for the Cu complexes were calculated for the
corresponding axial coordination of H2O above and below the plane (Fig. 2 for 1;
Fig. S2, for 21)). The energy differences between the two planes (H2O coordination)
were found to be 14.28 kcal/mol for 1 and 16.34 kcal/mol for 2, respectively. The extra
stability found in below-the-plane systems may be attributed to an intramolecular H-
bonding network between the axial H2O molecule and the C¼O O-atom of COOꢀ, NH

CHEMISTRY & BIODIVERSITY – Vol. 8 (2011)
135
Fig. 2. Proposed geometry and energy-minimized molecular structures, relative energies, and bond lengths
of 1 with H2O molecule a) above and b) below the plane
of amide group, and the stacking interaction between the indole ring of peptide and
phenyl ring of phen/bpy.
Based on the above results, a structure is proposed with an approximate square-
pyramidal geometry where four equatorial positions are occupied by NN and NO
donor atoms of phen/bpy and peptide, respectively, and the axial position is occupied by
a H2O molecule. This concurs with the X-ray structure of CuIIꢀgly-trpꢀphen complex
[23] where an amide N-atom was coordinated in the axial position instead of a H2O
molecule.
DNA Binding. The interaction of small molecules with double-helical DNA may
increase or decrease Tm, the temperature at which the double helix is cleaved into a
single-stranded DNA. While increase in Tm values indicates an intercalative or
phosphate binding, decrease is typical for base binding. The thermal denaturation
profile of DNA in the absence and presence of complexes is provided in Fig. 3. As can
be seen, an increase of ca. 48 for 1 and ca. 38 for 2 was observed in the Tm profile of
complexes as compared to free DNA. This provides a clear evidence for an
intercalative/phosphate binding of Cu complexes with DNA.
Hydrodynamic measurements that are sensitive to length change (i.e., viscosity and
sedimentation) are regarded as the least ambiguous and the most critical tests of a
binding model in solution in the absence of crystallographic structural data [31]. A
classical intercalation model demands that the DNA helix must lengthen as base pairs

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CHEMISTRY & BIODIVERSITY – Vol. 8 (2011)
Fig. 3. Thermal denaturation profiles of a) free CT DNA, and b) and c) after addition of 1 and 2,
respectively. The concentrations of 1 or 2 and DNA were 30 mm. Inset: differential melting curves.
are separated to accommodate the binding ligand/complex leading to an increase of
DNA viscosity. In contrast, a partial intercalation of ligand/complex could bend (or
kink) the DNA helix, and reduce its effective length and concomitantly its viscosity.
The effect of 1 and 2 on the viscosity of DNA at room temperature is shown in Fig. 4.
The viscosity of the DNA increases steadily with an increasing amounts of the
complexes. Such behaviour is consistent with other intercalators (i.e., EB), which
increase the relative specific viscosity for the lengthening of the DNA double helix
resulting from an intercalation. The results clearly emphasize that both of the
complexes intercalate between adjacent DNA base pairs, causing an extension in the
helix thereby increasing the viscosity of DNA [32].
Ethidium bromide (EB) emits intense fluorescence light in the presence of DNA,
due to its strong intercalation between the adjacent DNA base pairs. It was reported
earlier that the enhanced fluorescence can be quenched by the addition of a second
molecule [33][34]. The extent of quenching fluorescence EB bound to DNA was used
to determine the extent of binding between the second molecule and DNA. The
emission spectra of EB bound to DNA in the absence and the presence of the complex
1 is shown in Fig. 5 (Fig. S3, for 21)). The addition of complexes to DNA pretreated
with EB caused appreciable reduction in emission intensity, indicating that the
complexes efficiently compete with EB in binding to DNA. According to the
SternꢀVolmer equation, I0/I¼1þKsqr, where I0 and I are the fluorescence intensities in

CHEMISTRY & BIODIVERSITY – Vol. 8 (2011)
137
&
*
Fig. 4. Effect of increasing amounts of 1 ( ) and 2 ( ) on the relative viscosities of CT DNA at room
temperature in 5 mm Tris ·HCl buffer. Conditions: [DNA]¼200 mm, [complex]¼0–200 mm.
the absence and the presence of complexes, respectively, Ksq is linear Stern–Volmer
quenching constant dependent on the ratio of concentration of bound EB to the
concentration of DNA, and r is the concentration ratio of the complex to DNA. The
quenching plots (Insets of Fig. 5 and Fig. S31)) illustrate that the quenching of EB
bound to DNA by the complexes is in good agreement with the linear SternꢀVolmer
equation, which indicates that the complexes bind to DNA. In the plot of I0/I vs.
[complex]/[DNA], Ksq is given by the ratio slope/intercept, the Ksq values for 1 and 2
are 0.14 and 0.08, respectively.
Fluorescence Scatchard plots for the binding of EB to CT DNA in the presence of
complexes were obtained as described by Lepecq and Paoletti [35]. The binding
isotherms of EB and DNA in the absence and presence of the complexes were
determined experimentally, and are presented in Fig. 6. As can be seen from the plots, a
decrease of slope upon the addition of the complexes 1 and 2 confirm an exclusive
intercalative mode of binding with DNA.
DNA Cleavage. The cleavage activity was demonstrated by gel-electrophoresis
experiments using supercoiled (SC) plasmid pUC19 DNA in a medium Tris·HCl/NaCl
buffer. These experiments were monitored by the addition of varying concentrations of
the complex (0–100 mm) to 2 ml of pUC19 DNA, and the total volume was increased to
16 ml by adding 5 mm Tris·HCl/5 mm NaCl buffer. When DNA was incubated with
increasing concentrations of the complexes, SC DNA was degraded to nicked circular

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Fig. 5. Emission spectra of EB bound to DNA in the absence (· · ·) and presence (—) of 1. [1]/[DNA]¼0,
0.8, 1.60, 2.41, 3.19, 3.97, and 4.78, lex ¼540 nm. Inset: SternꢀVolmer quenching plot.
(NC) form. The catalytic activities of 1 and 2 are depicted in Fig. 7. The activity of 1
starts at a concentration as low as 5 mm (Fig. 7,a). At 50 mm, a complete conversion of
SC plasmid DNA into the NC form was observed, and at 100 mm of 1, the DNA was
completely smeared (Fig. 7,b). In contrast, only 40% cleavage was achieved with 2
(Fig. 7,c). This may be due to the efficient binding of 1 with DNA compared to that of 2
and may also be due to the generation of stable [Cu(phen)2]þ species which could be
related to the presence of an indole ring of trp-phe moiety, which is known to stabilize
the radical species. Similar observations were made by Garcꢀa-Raso et al. [23].
To ensure that the Cu complexes are solely responsible for the cleavage, several
control experiments were performed under identical conditions. No cleavage of DNA
was observed with free Cu (100 mm) and free ligands (100 mm; Fig. 7,a; Lanes 5, 6, and 7,
resp.). In a control experiment with DMSO (1 mm), a known radical scavenger, only
slight inhibition (ca. 2%) of DNA cleavage was observed (Fig. 7,a, Lane 8 for 1; and
.
Fig. 7,c, Lane 7 for 2), ruling out the possibility of DNA cleavage via OH -based
depurination pathway and also a possible oxidative cleavage [36] (dioxygenþCu
complexþtraces of DNA). This observation indicates that the DNA cleavage was the
result of hydrolysis of a phosphodiester bond [37–42]. Thus, the present study reveals
that no diffusable radical is involved in the DNA cleavage, as no external agents like
H2O2, MPA, ascorbic acid, or light were employed in the cleavage experiments.

CHEMISTRY & BIODIVERSITY – Vol. 8 (2011)
139
~
&
Fig. 6. Fluorescence Scatchard plots for the EB to CT DNA in the absence ( ) and presence of 1 ( ) and 2
*
( ). The term rEB is the concentration ratio of bound EB to total DNA, and cf is the concentration of
free EB.
The cleavage of pUC19 DNA by 1 and 2 was kinetically monitored by
quantification of SC and NC forms of DNA. The observed distribution of SC and
NC DNA in agarose gel provides a measure of the extent of hydrolysis of the
phosphodiester bond in each plasmid DNA, and the data were used to perform simple
kinetics. The kinetic plots (Fig. 8 for 1 and Fig. S4, for 21)), which show a decrease of SC
and an increase of NC forms of DNA vs. time, follow the pseudo-first-order kinetics,
and both forms fitted well to a single exponential curve. From these curve fits, the DNA
hydrolysis rates were determined as 1.74 (R¼0.985) and 0.65 hꢀ1 (R¼0.963) for 1 and
2, respectively. The enhancement of DNA hydrolysis rate constant by metal complexes
in the range of 0.09–0.25 hꢀ1 was considered impressive [43][44]. With the rate
constants of 1.74 (for 1) and 0.65 hꢀ1 (for 2), which amounts to 2.40–4.10ꢁ107-fold rate
enhancement compared to non-catalysed double stranded DNA, is significant
considering the type of ligands and experimental conditions.
Based on the above conclusions, a tentative mechanism for the hydrolytic cleavage
of DNA may be as follows: i) activation of the Cu-coordinated H2O molecule, and
formation of a nucleophile CuꢀOH species; ii) phosphodiester coordination with
amide and amine N-atoms through an electrostatic interaction; iii) intramolecular OH
transfer from CuII to phosphate. There are literature reports on an intramolecular
pathway for the hydrolysis of phosphate diester [45]. A hydrolytic cleavage mechanism
was established by enzymatic religation of the termini with T4 DNA ligase which

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Fig. 7. Agarose gel-electrophoresis pattern for the cleavage of supercoiled pUC19 DNA by 1 and 2 at 378 in
a buffer containing 5 mm Tris ·HCl/5 mm aqueous NaCl. a) Lane 1: DNA control, Lane 2: DNAþ1
(5 mm), Lane 3: DNAþ1 (10 mm), Lane 4: DNAþ1 (25 mm), Lane 5: DNAþCu(NO3)2 (100 mm), Lane
6: DNAþphen (100 mm), Lane 7: DNAþtrp–phe (100 mm), Lane 8: DNAþ1 (25 mm)þDMSO
(1 mm). b) Lane 1: DNA control, Lane 2: DNAþ1 (25 mm), Lane 3: DNAþ1 (50 mm), Lane 4: DNAþ1
(100 mm). c) Lane 1: DNA control, Lane 2: DNAþ2 (10 mm), Lane 3: DNAþ2 (25 mm), Lane 4: DNAþ
2 (50 mm), Lane 5: DNAþ2 (75 mm), Lane 6: DNAþ2 (100 mm), Lane 7: DNAþ2 (100 mm)þDMSO
(1 mm).
requires 5’-phosphate and 3’-OH groups. Based on the efficiency of religation, it was
concluded that hydrolysis gave equally probable 3’- and 5’-products [46].
Accordingly, a LꢀMꢀOH form, which is the active species in the hydrolysis of
phosphate backbone in an enzyme/model, is depicted in Scheme 2.
Conclusions. – Two new CuIIꢀpeptideꢀphen/bpy complexes were successfully
synthesized, and their structures were established. Their DNA binding and cleavage
abilities were investigated. The CuII complexes bind to CT DNA in an exclusive
intercalative mode. They show good catalytic activity in the absence of any external
agents or light under physiological conditions.
P. R. R. thanks the Council of Scientific and Industrial Research (CSIR) New Delhi for financial
assistance through major research project. He also thanks UGC for the award of Emeritus Fellowship.

CHEMISTRY & BIODIVERSITY – Vol. 8 (2011)
141
Fig. 8. Disappearance of supercoiled (SC) DNA and formation of nicked circular form (NC) in the
presence of complex 1. Conditions: [complex]¼50 mm; in aqueous buffer (pH 7.2) at 378.
Scheme 2. A Proposed Mechanism for the DNA Cleavage
Experimental Part
General. Boc-tryptophan, phenylalanine methyl ester, Cu(NO3)2 ·3 H2O, 2,2’-bipyridine, 1,10-
phenanthroline, and ethidium bromide (EB) were obtained from Sigma (99.99% purity), USA, and
were of anal. grade. The peptide trp-phe was synthesized by the known procedure [47]. The calf thymus

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CHEMISTRY & BIODIVERSITY – Vol. 8 (2011)
(CT) DNA was obtained from Fluka (Switzerland); and pUC 19 DNA, agarose, and Tris·HCl were
obtained from Bangalore Genei (India). All the chemicals were used as supplied. UV/VIS Spectra:
Shimadzu 160A spectrophotometer (800–200 nm) and JASCO V-530 UV/VIS spectrophotometer; with
1-cm quartz micro-cuvettes. IR Spectra: Perkin-Elmer FT-IR spectrometer, in KBr pellets, in the 4000–
400 cmꢀ1 range. ESI-MS: Quattro Lc (Micro mass, Manchester, UK) triple quadruple mass spectrometer
with MassLynx software. Elemental analyses: Heraeus Carlo Erba 1108 elemental analyzer. Molar
conductivity: Digisun digital conductivity bridge (model: DI-909) with a dip-type cell. Magnetic
susceptibilities of complexes: at r.t. on a Faraday balance (CAHN-7600) using Hg[Co(NCS)4] as the
standard; diamagnetic corrections were made by using Pascalꢂs constants [48]. EPR Spectra: JEOL
(JES-FA200) X-band spectrometer at r.t. TGA Analyses: METTLER TOLEDO (TGA/SDTA 851e)
thermo-analyzer at 25–3008, heating rate of 58/min. The molecular-modeling calculations were carried
out with semi-empirical PM3 Hamiltonian as implemented in hyperchem software programme package.
Gel pattern of electrophoresis was photographed by Alpha-Innotec gel documentation system (USA).
Concentrated CT DNA stock soln. was prepared in 5 mm Tris·HCl/50 mm NaCl in double
dist. H2O at pH 7.5, and the concentration of DNA soln. was determined by UV absorbance at 260 nm
(e¼6600 mꢀ1 cmꢀ1) [49]. Soln. of CT DNA in 5 mm Tris·HCl/50 mm NaCl (pH 7.5) gave a ratio of UV
absorption at 260 and 280 nm, A260/A280, of ca. 1.8–1.9, indicating that the DNA was sufficiently free of
protein [50]. All stock solns. were stored at 48 and were used within 4 d. The concentration of EB was
determined spectrophotometrically at 480 nm (e¼5680 mꢀ1 cmꢀ1) [51].
Syntheses of Peptide and Complexes. The dipeptide trp-phe was synthesized by conventional soln.-
phase method [47] using EDCI and HOBt as coupling agents and dry CH2Cl2 as solvent. While (tert-
butoxy)carbonyl (Boc) group was used for N-protection, the C-terminal was protected as methyl ester
(COOMe). Deprotection of Boc was achieved with TFA/CH2Cl2 1:1, and saponification of the methyl
ester required LiOH in THF/MeOH/H2O 3 :1:1. 1H-NMR (200 MHz, CDCl3): 2.00 (br., 2 H); 2.93–3.95
(m, 5 H); 4.85 (m, 1 H); 6.82–7.20 (m, 8 H); 7.21 (m, 2 H); 8.00 (m, 1 H); 10.10 (d, 2 H). ESI-MS: 351.
To a mixture of trp-phe (0.351 g, 1.0 mmol) and KOH (0.056 g, 1.0 mmol) in H2O, an aq. soln. of
Cu(NO3)2 ·3 H2O (0.241 g, 1.0 mmol) was added with stirring, followed by the addition of phen (0.198 g,
1.0 mmol) or bpy soln. (0.156 g, 1.0 mmol) in MeOH (2 ml). The stirring was continued for ca. 3 h at 608.
After cooling to r.t., an aq. soln. of NaClO4 ·H2O (0.181 g, 1.0 mmol) was added, and stirring was
continued for another 30 min. The resulting violet-blue precipitate was filtered and washed with EtOH
and cyclohexane, and dried in vacuo to afford 72 and 65% of 1 and 2, resp.
Data of [CuII(trp-phe)(phen)(H2O)]ClO4 (1). UV/VIS (MeOH): 626, 268, 222, 206. IR (KBr): 3420
(br.), 3263s, 2953m, 1684vs, 1654s, 1576m, 1518s, 1426s, 1384s, 1146m, 1086m, 920m, 848s, 722m, 623w,
512m, 454m. ESI-MS: 593 ([Cu(trp-phe)(phen)]þ ). Anal. calc. for C32H30ClCuN5O8 (711.61): C 54.01, H
4.25, N 9.84; found: C 53.01, H 4.02, N 9.95. meff ¼2.01 BM. LM (10ꢀ3 m in MeOH, 258)¼1.60 Wꢀ1 cm2
molꢀ1
.
Data of [CuII(trp-phe)(bpy)(H2O)]ClO4 (2). UV/VIS (MeOH): 613, 310, 298, 244. IR (KBr): 3446
(br.), 3253s, 2954m, 1684vs, 1654s, 1603m, 1474s, 1445vs, 1382s, 1250m, 1094m, 1032s, 768vs, 731m, 703m,
623w, 512m, 454m. ESI-MS: 587 ([[Cu(trp-phe)(bpy)]þNa]þ ). Anal. calc. for C30H30ClCuN5O8
(687.59): C 52.40, H 4.40, N 10.19; found: C 52.01, H 4.25, N 10.56. meff ¼1.69 BM. LM (10ꢀ3 m in
MeOH, 258)¼2.10 Wꢀ1 cm2 molꢀ1
.
DNA Binding. Thermal-Denaturation Studies. The thermal-denaturation studies were performed on
Shimadzu 160A spectrophotometer equipped with a thermostatic cell holder. DNA (30 mm) was treated
with complexes 1 or 2 (30 mm) in a ratio of 1:1 in buffer (5 mm Tris·HCl/50 mm NaCl) at pH 7.5. The
samples were continuously heated at the rate of 18/min while monitoring the absorption changes at
260 nm. Melting-temp. (Tm) and the melting-interval (DTm) data were obtained according the procedure
reported in [52].
Viscometric Studies. Viscometric titrations were performed with an Ostwald viscometer at r.t. The
concentration of DNAwas 200 mm, complex concentration was varied from 0–200 mm, and the flow times
were measured with a digital timer. Each sample was measured three times for accuracy, and an average
flow time was determined. Data is presented as (h/h0)1/3 vs. [complex]/[DNA], where h is the viscosity of
DNA in the presence of complex and h0 that of DNA alone. Viscosity values were calculated from the

CHEMISTRY & BIODIVERSITY – Vol. 8 (2011)
143
observed flow time of DNA containing solns. (t) corrected for that buffer alone (5 mm Tris·HCl/50 mm
NaCl) (t0), h¼(t–t0).
Fluorescence-Spectroscopic Studies. Fluorescence spectra were recorded with SPEX-Fluorolog
0.22 m fluorimeter equipped with a 450-W Xe lamp. The slit widths were 2ꢁ2ꢁ2ꢁ2, and the emission
spectral range was 550–650 nm. All fluorescence titrations were carried out in 5 mm Tris·HCl/50 mm
NaCl (pH 7.5). Soln. containing DNA and EB was titrated with varying concentrations of 1 and 2. The
concentration of DNA and EB was maintained at 41 mm, and the concentration of complexes was in the
range of 0–196 mm. The solns. containing DNA and EB were mixed (equimolar ratio), and the complexes
1 and 2 were added and allowed to equilibrate. They were excited at 540 nm (lmax for EB), and the
fluorescence emissions at 598 nm (lmax) were recorded.
Fluorescence spectra were also utilized to obtain Scatchard plots. For this, titration of DNA against
EB in the absence and presence of Cu complexes were performed. Initial concentration of DNA in 5 mm
Tris·HCl/50 mm NaCl was 20 mm, and the concentration of 1 and 2 were kept at 50 mm. After each
addition of EB to the solns. containing DNA and Cu complexes, the emission spectra were recorded in
the range of 550–650 nm with excitation at 540 nm at 258. Corrections were made to the data for the
volume changes during the course of titrations. The data were analyzed by the method of Lepecq and
Paoletti [35] to obtain bound (cb) and free (cf) concentration of EB, and Scatchard plots were obtained by
plotting rEB/cf vs. rEB (where r¼cb/conc. of DNA).
DNA Cleavage. The DNA cleavage experiments were performed with supercoiled (SC) plasmid
pUC19 DNA at pH 7.2 in Tris·HCl buffer. DNA Cleavage experiments were monitored by the addition
of varying concentrations of the complexes 1 and 2 (0–100 mm) to 2 ml of pUC19 DNA, the total volume
was increased to 16 ml by adding 5 mm Tris·HCl/5 mm NaCl buffer. After mixing, DNA solns. were
incubated for 1 h at 378. The samples were subjected to electrophoresis in TAE (Tris·AcOH EDTA
buffer) for 2 h at 60 V. The gel was stained with 0.5 mg/ml EB and photographed under UV light.
Blank experiments were carried out using free Cu(NO3)2 ·3 H2O and free ligands (trp-phe, phen) at
100 mm. Experiments were also carried out with known radical scavenger DMSO. Kinetic analyses were
carried out at a fixed concentration of DNA and 1 or 2 (50 mm), with different incubation times under
identical experimental conditions.
REFERENCES
[1] M. J. Clarke, F. Zhu, D. R. Frasca, Chem. Rev. 1999, 99, 2511.
[2] J. Liu, W.-J. Mei, A.-W. Xu. C.-P. Tan, S. Shi, L.-N. Ji, Antiviral Res. 2004, 66, 65.
[3] J. Liu, W.-J. Mei, A.-W. Xu, C.-P. Tan, L.-N. Ji, Transition Met. Chem. (Dordrecht, Neth.) 2003, 28,
500.
[4] D. S. Sigman, A. Mazumder, D. M. Perrin, Chem. Rev. 1993, 93, 2295.
[5] K. E. Erkkila, D. T. Odom, J. K. Barton, Chem. Rev. 1999, 99, 2777.
[6] M. Narra, P. Elliott, S. Swavey, Inorg. Chim. Acta 2006, 359, 2256.
[7] D. Lawrence, V. G. Vaidyanathan, B. U. Nair, J. Inorg. Biochem. 2006, 100, 1244.
[8] H. Chao, L.-N. Ji, Bioinorg. Chem. Appl. 2005, 3, 15.
[9] L.-F. Tan, H. Chao, Y.-J. Liu, H. Li, B. Sun, L.-N. Ji, Inorg. Chim. Acta 2005, 358, 2191.
[10] J.-Z. Wu, L. Yuan, Y. Yu, Inorg. Chim. Acta 2006, 359, 718.
[11] A. E. Friedman, J. C. Chambron, J. P. Sauvage, N. J. Turro, J. K. Barton, J. Am. Chem. Soc. 1990, 112,
4960.
[12] L.-F. Tan, H. Chao, H. Li, Y.-J. Liu, B. Sun, W. Wei, L.-N. Ji, J. Inorg. Biochem. 2005, 99, 513.
[13] P. J. Dandliker, R. E. Holmlin, J. K. Barton, Science 1997, 275, 1465.
[14] I. Ortmans, C. Moucheron, A. Kirsch-De Mesmaeker, Coord. Chem. Rev. 1998, 168, 233.
[15] L.-N. Ji, X.-H. Zou, J.-G. Liu, Coord. Chem. Rev. 2001, 216–217, 513.
[16] H. Xu, K.-C. Zheng, L.-J. Lin, H. Li, Y. Gao, L.-N. Ji, J. Inorg. Biochem. 2004, 98, 87.
[17] Y.-J. Liu, H. Chao, L.-F. Tan, Y.-X. Yuan, W. Wei, L.-N. Ji, J. Inorg. Biochem. 2005, 99, 530.
[18] G. Yang, J. Z. Wu, L. Wang, L. N. Ji, X. Tian, J. Inorg. Biochem. 1997, 66, 141.
[19] G. Yang, L. Wang, L.-N. Ji, J. Inorg. Biochem. 1997, 67, 289.

144
CHEMISTRY & BIODIVERSITY – Vol. 8 (2011)
[20] Y. Jin, M. A. Lewis, N. H. Gokhale, E. C. Long, J. A. Cowan, J. Am. Chem. Soc. 2007, 129, 8353.
[21] R. Nagane, T. Koshigoe, M. Chikira, E. C. Long, J. Inorg. Biochem. 2001, 83, 17.
[22] R. S. Kumar, S. Arunachalam, Eur. J. Med. Chem. 2009, 44, 1878.
´
[23] A. Garcꢃa-Raso, J. J. Fiol, B. Adrover, V. Moreno, I. Mata, E. Espinosa, E. Molins, J. Inorg. Biochem.
2003, 95, 77.
[24] L. H. Abdel-Rehman, L. P. Battaglia, M. R. Mahmoud, Polyhedron 1996, 15, 327.
[25] K. Nakamoto, ꢄInfrared and Raman Spectra of Inorganic and Coordination Compoundsꢂ, 3rd edn.,
John Wiley & Sons, New York, p. 228.
[26] G. G. Mohamed, M. M. Omar, A. M. M. Hindy, Spectrochim. Acta, Part A 2005, 62, 1140.
[27] R. N. Patel, N. Singh, V. L. N. Gundla, Indian J. Chem., Sect. A 2006, 45, 614.
[28] T. Murakami, S. Hatakeyama, S. Igarashi, Y. Yukawa, Inorg. Chim. Acta 2000, 310, 96.
[29] D. Kivelson, R. Neiman, J. Chem. Phys. 1961, 35, 149.
´
ˇ
[30] P. Kopel, Z. Travnicek, J. Marek, M. Korabik, J. Mrozinski, Polyhedron 2003, 22, 411.
[31] S. Satyanarayana, J. C. Dabrowiak, J. B. Chaires, Biochemistry 1993, 32, 2573.
[32] S. Satyanarayana, J. C. Dabrowiak, J. B. Chaires, Biochemistry 1992, 31, 9319.
[33] B. C. Baguley, M. Le Bret, Biochemistry 1984, 23, 937.
[34] J. R. Lakowicz, G. Webber, Biochemistry 1973, 12, 4161.
[35] J.-B. Lepecq, C. Paoletti, J. Mol. Biol. 1967, 27, 87.
[36] R. Ren, P. Yang W. Zheng, Z. Hua, Inorg. Chem. 2000, 39, 5454.
[37] L. A. Basile, A. L. Raphael, J. K. Barton, J. Am. Chem. Soc. 1987, 109, 7550.
[38] T. Itoh, H. Hisada, T. Sumiya, M. Hosono, Y. Usui, Y. Fujii, Chem. Commun. 1997, 677.
[39] K. M. Deck, T. A. Tseng, J. N. Burstyn, Inorg. Chem. 2002, 41, 669.
[40] L. Zhu, O. dos Santos, C. W. Koo, M. Rybstein, L. Pape, J. W. Canary, Inorg. Chem. 2003, 42, 7912.
´
[41] L. Bonfa, M. Gatos, F. Mancin, P. Tecilla, U. Tonellato, Inorg. Chem. 2003, 42, 3943.
[42] F. Mancin, P. Tecilla, New. J. Chem. 2007, 31, 800.
[43] J. Rammo, R. Hettich, A. Roigk, H.-J. Schneider, Chem. Commun. 1996, 105.
[44] A. Roigk, R. Hettich, H.-J. Schneider, Inorg. Chem. 1998, 37, 751.
[45] J.-H. Li, J.-T. Wang, P. Hu, L.-Y. Zhang, Z.-N. Chen, Z.-W. Mao, L.-N. Ji, Polyhedron 2008, 27, 1898.
[46] R. Ott, R. Krꢅmer, Appl. Microbiol. Biotechnol. 1999, 52, 761.
[47] M. Bodanszky, A. Bodanszky, ꢄThe Practices of Peptide Synthesisꢂ, Springer, New York, 1984.
[48] B. N. Figgiꢂs, J. Lewis, ꢄModern Coordination Chemistryꢂ, Eds. J. Lewis, R. C. Wilkinson, Wiley
Interscience, NewYork, 1967, pp. 403.
[49] J. Marmur, J. Mol. Biol. 1961, 3, 208.
[50] M. E. Reichmann, C. A. Rice, C. A. Thomas, P. Doty, J. Am. Chem. Soc. 1954, 76, 3047.
[51] F. Garland, D. E. Graves, L. W. Yielding, H. C. Cheung, Biochemistry 1980, 19, 3221.
[52] ꢄMethods in Molecular Biologyꢂ, Eds. W. D. Wilson, F. A. Tanious, M. Fernandez-Saiz, C. T. Rigl,
K. R. Fox, Humana, Clifton, New Jersey, 1997, pp. 90.
Received November 21, 2010