← Back
Polypyridyl Complexes of Ruthenium(II): Stabilization of G‐quadruplex DNA and Inhibition of Telomerase Activity
DOI: 10.1002/cplu.201200039
Polypyridyl Complexes of Ruthenium(II): Stabilization of
G-quadruplex DNA and Inhibition of Telomerase Activity
Du Liu,[a] Yanan Liu,[a] Chuan Wang,[a] Shuo Shi,[c] Dongdong Sun,[a] Feng Gao,*[b]
Qianling Zhang,*[d] and Jie Liu*[a]
Two ruthenium(II) complexes [Ru(phen)2(tip)](ClO4)2 (1) and
[Ru(bpy)2(tip)](ClO4)2 (2; phen = 1,10-phenanthroline, bpy = 2,2’bipyridine, tip = 2-thiophenimidazo[4,5-f][1,10]phenanthroline)
were synthesized and characterized by elemental analysis,
1
H NMR spectroscopy, and electrospray ionization-mass spectrometry to explore the role of metal complexes as novel telomeric quadruplex stabilizers. The different quadruplex binding
properties of these compounds were evaluated by absorption
and emission analyses, circular dichroism spectroscopy, fluorescence resonance energy transfer (FRET) melting assay, NMR
spectroscopy, and molecular modeling. The results show that
both complexes can well induce and stabilize different G-quad-
ruplex structures using a 1:1 [quadruplex]/[complex] binding
mode ratio. Complex 1 exhibits higher interaction ability at
1.43 106 m1 binding affinity and superior G-quadruplex selectivity over duplex DNA through multiple interaction (mainly intercalating) with the G-quadruplex at the 3’-terminal face. Furthermore, polymerase chain reaction (PCR)-stop assay, electrophoretic mobility shift assay, telomerase repeat amplification
protocol, and MTT assay demonstrate that complex 1 not only
can stabilize dimer forms of the G-quadruplex at low concentrations but also exhibit better inhibitory activity for telomerase and cancer cells. The results suggest that complex 1 may
be a potential telomerase inhibitor for cancer chemotherapy.
Introduction
G-quadruplexes (or G-tetrads) are functionally useful secondary
DNA structures containing G-quartets stabilized through
Hoogsteen hydrogen bonding.[1] These structures are found
throughout the human genome and are currently being considered as potential anticancer targets.[2] Telomeric DNA consists of tandem repeats of sequence d[(TTAGGG)n] and is the
most studied DNA sequence. This sequence can cap the ends
of chromosomes and protect them from deleterious processes
during replication steps.[3] A previous study has indicated that
telomeric DNA may fold into G-quadruplex structures in the
presence of metal ions, such as K + or Na + .[4] The formation of
G-quadruplex by telomeric DNA inhibits the activity of telomerase,[5] an enzyme not found in most normal somatic cells,
but present in 85–90 % of cancer cells and contributes to the
immortality of these cells.[6] Therefore, the design of drugs that
target and stabilize the telomeric G-quadruplex is a rational
and promising approach to interfere with telomerase activity
in tumor cells and to act as potential anticancer agents.[7]
Recently, several research groups have synthesized
a number of small-molecule ligands for G-quadruplex structure
stabilization and telomerase activity inhibition.[8] Several metal
complexes, which generally have a positively charged center
or substituents and p-delocalized system, have been reported
to interact with G-quadruplex.[9]
Octahedral metal complexes can also be designed to bind
G-quadruplex using a large planar aromatic intercalating
ligand for DNA binding and ancillary ligands for shape and
functional group recognition within the major groove.[10]
Ruthenium(II) complexes with polypyridyl ligands as typical
octahedral metal complexes have prominent DNA binding
ChemPlusChem 2012, 77, 551 – 562
properties resulting from a combination of easily constructed
rigid chiral structures spanning all three spatial dimensions
and a rich photophysical repertoire.[11] Some of these complexes have been investigated as nucleic acid probes, synthetic
restriction enzymes, anticancer drugs, and DNA footprinting
agents, among others.[12] To date, a few RuII complexes have
been found to promote the formation and stabilization of Gquadruplexes.[13] Shi et al. have reported the remarkable ability
of a novel dinuclear complex to promote an antiparallel Gquadruplex formation.[14] Ruthenium(II) complexes containing
the dppz ligand can serve as a prominent molecular “light
switch” for both G-quadruplexes and i-motif, but it preferentially binds to G-quadruplexes over the i-motif.[15] Thomas and
co-workers investigated the binding preferences of a dinuclear
[a] Dr. D. Liu,+ Y. Liu,+ C. Wang, D. Sun, Prof. J. Liu
Department of Chemistry, Jinan University
Guangzhou 510632 (P. R. China)
Fax: (+ 86) 20-8522-1263
E-mail: tliuliu@jnu.edu.cn
[b] Dr. F. Gao
School of Chemistry and Chemical Engineering, Sun Yat-Sen University
Guangzhou 510275 (P. R. China)
[c] Prof. S. Shi
Department of Chemistry, Tongji University
Shanghai 200092 (P. R. China)
[d] Prof. Q. Zhang
Chemistry and Chemical Engineering, Shenzhen University
Shenzhen 518061 (P. R. China)
[+] These authors contributed equally to this work.
Supporting information for this article is available on the WWW under
http://dx.doi.org/10.1002/cplu.201200039.
2012 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
551
J. Liu et al.
ruthenium(II) complexes with
different quadruplex DNA structures. It was found that the differences in quadruplex binding
affinity and optical signature are
rationalized through a consideration of the structural features
of the quadruplexes.[16] In 2011,
they found that polypyridyl complexes of RuII display sequence
selectivity and high-affinity binding to duplex DNA through
groove binding.[17] However,
they have not determined
whether the inhibition of telomerase activity is relevant to the
stabilization of this G-quadru- Scheme 1. Synthetic routes for the ligand (tip) and ruthenium complexes [Ru(phen) (tip)](ClO ) (1) and [Ru(bpy) 2
4 2
2
plex, or even if there are further (tip)](ClO4)2 (2). HAC = CH3COOH.
effects of the antitumor activity.
Herein, two RuII complexes
[Ru(phen)2(tip)](ClO4)2 (1) and
[Ru(bpy)2(tip)](ClO4)2 (2; phen = 1,10-phenanthroline, bpy = 2,2’
-bipyridine, tip = 2-thiophenimidazo[4,5-f][1,10]phenanthroline),
which show some interesting properties through interaction
with G-quadruplex DNA, were designed and synthesized. The
spectroscopic, biochemical, and cellular properties of these
compounds were examined to reveal their interactions with
the telomeric G-quadruplex DNA (HTG21) as well as the relationship between the inhibition of telomerase and antitumor
activities. The title complexes acting as G-quadruplex stabilizers showed effective inhibition of telomerase and antitumor
activities, thus suggesting that telomerase G-quadruplex may
act as a potential antitumor agent. The synthetic route and
structure of complex 1 and 2 are shown in Scheme 1.
Results and Discussion
Fluorescence behavior of G-quadruplex and duplex DNA
(ds26)
Zhou and co-workers have reported fluorescence selectivity as
an approach to detect the selectivity between G-quadruplex
and other DNA structures.[18] Here, the fluorescence behavior
of Ru complexes with different DNA is reported. Human telomeric DNA (HTG21) was selected to form the G-quadruplex
structures in the presence of K + , whereas ds26 was selected as
the duplex DNA structure. The results of the different fluorescence spectra are shown in Figure 1. Both complexes 1 and 2
emit luminescence at ambient temperature, with a maximum
appearing at 598 nm.
Upon the addition of different DNA, it was clear that there
was a more remarkable fluorescence enhancement in the presence of HTG21 than ds26. This observation maybe implies that
the complexes are more inaccessible to water molecules and
that there is a greater overlap between the aromatic surfaces
of the metal complexes and the bases when bound to quadruplex as opposed to duplex DNA.[16, 19] In particular, complex 1
552
www.chempluschem.org
Figure 1. Emission comparison of 5 mm solution of complexes 1 (A) and 2 (B)
in the presence of HTG21 and ds26 using Tris-KCl buffer (100 mm KCl,
10 mm Tris-HCl, pH 7.4, [Ru]/[DNA] = 2:1) at lex = 460 nm. C) Relative fluorescence strength of complexes 1 and 2. Results are the mean values of at least
three independent experiments.
2012 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemPlusChem 2012, 77, 551 – 562
Bioinorganic Chemistry
promoted a bigger fluorescence intensity of the G-quadruplex,
more than 1.2 times larger than duplex DNA, whereas complex 2 only has 0.7-fold increase in selectivity for G-quadruplex.
The different fluorescence behaviors between the two RuII
complexes and telomeric quadruplex inspired us to further
study their different DNA-binding properties by a variety of research methods.
Emission spectra analyses and binding affinities
Given that fluorescence spectra can reflect the information
about the local environmental changes in a chromophore,
then these spectra can be used to probe the interaction between the fluorophore and its environment.[20] In the current
experiment, the binding of the Ru complexes to G-quadruplex
DNA (HTG21) was investigated using fluorescence titration.
The results are illustrated in Figure 2. Upon the addition of
r=C f ¼ nK b rK b
ð1Þ
r ¼ C b =C DNA
C b ¼ C t ðFF 0 Þ=ðF max F 0 Þ
where Ct is the total compound concentration, F is the observed fluorescence emission intensity at given DNA concentration, F0 is the intensity in the absence of DNA, and Fmax is
the fluorescence of the totally bound compound. Binding data
were cast into the form of a Scatchard plot of r/Cf versus r,
where r is the binding ratio Cb/[DNA], and Cf is the free ligand
concentration.[22] The values of the binding constants for complexes 1 and 2 with G-quadruplexes were 1.43 106 and 6.33
105 m1, respectively. These observations imply that the interaction between complex 1 and quadruplex DNA is stronger compared with that between complex 2 and the quadruplex DNA.
Absorption spectroscopy studies and binding ability
Figure 2. Emission spectral traces of complexes 1 (A) and 2 (B) in Tris-KCl
buffer (100 mm KCl, 10 mm Tris-HCl, pH 7.4) at increasing ratios of [HTG21]/
[Ru] = 0–2.0, [Ru] = 5 mm.
HTG21, the emission intensities of complexes 1 and 2 increased to approximately 1.32 and 0.8 times larger than the
original intensities, respectively. The enhanced fluorescence in
these complexes implies that these complexes can interact
with HTG21 and be protected by DNA efficiently, because the
hydrophobic environment inside the DNA helix decreases the
accessibility of solvent water molecules to the complex and
thus the complex mobility is restricted at the binding site, thus
leading to the decrease of vibrational modes of relaxation.[17, 21]
Based on the emission enhancement, the intrinsic binding constant was obtained according to the Scatchard Equation (1):
ChemPlusChem 2012, 77, 551 – 562
Electronic spectra of the complexes in the absence and presence of quadruplex were obtained to gain insight into the
binding ability between the RuII complexes and G-quadruplex
DNA (Figure S1 in the Supporting Information). Both RuII complexes were characterized using a metal-to-ligand-chargetransfer (MLCT) transition band and an intraligand (IL) absorption band at approximately 457 and 288 nm, respectively. However, another narrow separated band at approximately 263 nm
was present in complex 1. Upon the addition of HTG21, both
the MLCT and IL absorption bands of these RuII complexes exhibited obvious hypochromisms (H) and red shifts (Dl), thus
indicating that both complexes can intercalate the G-quadruplex.[23]
The addition of HTG21 to the solution of complex 1 led to
a red shift of 6 nm at 457 nm and hypochromism of 25.1 % for
the band at 288 nm. However, the addition of HTG21 to the
solution of complex 2 in the same buffer led to only a red shift
of 3 nm and 11.1 % hypochromism of the band at 457 nm
(Table 1). The absorption titration is well consistent with the
result of the emission spectroscopic analysis. The higher binding affinity of complex 1 is probably results from the greater
planar area of the ancillary ligand, intercalating with the base
pairs or entering into the grooves within DNA.[17, 24]
Table 1. Absorption spectra (lmax) and DNA-binding date of complexes 1
and 2.
Complex
lmax (free) [nm]
lmax (bound) [nm]
Dl [nm]
DH [%]
1
263
288
457
288
457
264
290
463
289
460
1
2
6
1
3
24.8
25.1
11.8
19.2
11.1
2
2012 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chempluschem.org
553
J. Liu et al.
Continuous variation analysis and binding stoichiometric
ratios
Continuous variation analysis using the luminescence intensities was performed to further validate the meaningful binding
stoichiometries of the Ru complexes with quadruplex DNA
(Figure 3). The point of intersection for complexes 1 and 2
with telomeric G-quadruplex is X = 0.51 and 0.56, respectively.
These data are consistent with the 1:1 [quadruplex]/[complex]
binding mode, suggesting a specific ruthenium–quadruplex interaction with a single guanine tetrad.[25]
Figure 3. The result of continuous variation analysis for complexes 1 and 2
with HTG21 in Tris-KCl buffer (100 mm KCl, 10 mm Tris-HCl, pH 7.4).
Stability of G-quadruplex by the fluorescence resonance
energy transfer (FRET) melting assay
Thermodynamic stability of the Ru complexes to G-quadruplex
DNA was determined using the melting temperature of the Gquadruplex DNA by a FRET melting assay. Change in melting
temperature (DTm) values were calculated by subtracting the
Tm of the nucleic acid with the complex from the Tm of the free
fluorescent-labeled oligonucleotide F21T. The DTm values of
the F21T DNA treated with the complexes are calculated, and
their concentration-dependent melting curves are shown in
Figure 4 A,B. All Tm values of the samples incubated with the
complexes increased compared with the control value
(51.5 8C), thus indicating that the Ru complexes could enhance
the thermodynamic stability of this oligomer. The 12.8 8C increase in the melting temperature (DTm) of complex 1 at
[Ru]/[F21T] = 10 ratio shows its high degree of stabilization for
G-quadruplex DNA. Hence, complex 1 is a more effective stabilizer of G-quadruplex DNA than complex 2 (DTm = 8.8 8C).
These activity differences are in accordance with the result of
the above studies.
Thermal denaturation analyses by circular dichroism (CD)
spectroscopy were also conducted to determine further the
complex-induced stabilization of a folded quadruplex. As the
ratio of Ru complexes to HTG21 equaled 10 (i.e. [HTG21] =
2 mm, [Ru] = 20 mm), thermal denaturation curves from the CD
signal at 295 nm are shown in Figure 4 C. Complex 2 has
a weak stabilizing effect on G-quadruplex, with only approximately 8.6 8C increase in the Tm value, whereas complex 1
could increase the Tm value from 60.2 to 73.4 8C (Table S1). This
554
www.chempluschem.org
Figure 4. Plot of DTm versus complex concentrations. FRET melting profiles
of 0.2 mm F21T with complexes 1 (A) and 2 (B) in Tris-KCl buffer. C) Thermal
denaturation curves from a CD signal at 295 nm with 2 mm HTG21 and in
the presence of complexes 1 and 2 (20 mm) in Tris-KCl buffer.
result is consistent with the FRET assay results, further implying
that complex 1 possesses a higher stabilizing ability than complex 2.
Furthermore, the G-quadruplex selectivity of complexes was
assessed by a competition FRET experiment where different
ratios of nonfluorescent duplex DNA (ds26) were added to the
classic FRET experiment with the telomeric sequence F21T
(Figure 5).[26] In the presence of various amounts of competitor
ds26, the thermal stabilization of F21T enhanced by the complexes was slightly affected, whereas the addition of a 50
molar excess of base pairs induces a decrease in stabilization
(Figure 5 C).[27] The results suggest that the binding of Ru complexes to a quadruplex is at least 40-fold higher than that to
a duplex. The combined results of FRET competition assay,
which well verifies the fluorescence selectivity, demonstrate
that the title complexes can be considered as a new class of
highly selective G-quadruplex binding ligands.
2012 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemPlusChem 2012, 77, 551 – 562
Bioinorganic Chemistry
Figure 6. 1H NMR analysis of titration of the G4 A3-quadruplex with complex 1 at various [Ru]/[G4 A3-quadruplex] ratios in 90 % H2O/10 % D2O with
150 mm KCl, 25 mm KH2PO4, 1 mm EDTA (pH 7.40). Only the proton signals
corresponding to the imino group (400 MHz, 25 8C) are shown.
identical to those exhibited by [PtII(dppz-COOH)(NC)]CF3SO3
(dppz-COOH = 11-carboxydipyrido[3,2-a:2’,3’-c]phenazine)[28]
and Hoechst 33258,[30] both of which are known to bind to
a G-quadruplex. Combined with the optical experiments and
the NMR analysis, we predict that complex 1 prefers intercalating with the G-tetrad at 3’-terminal face and/or multiple stacking on the G-tetrad.
Molecular modeling and binding mode
Figure 5. Competitive FRET melting curves of F21T with 1 mm of complexes 1 (A) and 2 (B) and duplex competitor ds26 in Tris-KCl (100 mm KCl,
10 mm Tris-HCl, pH 7.4) F21T = 0.2 mm. C) Competition FRET experiment of
complexes for the G-quadruplex DNA sequence over duplex DNA. Results
are the mean values of at least three independent experiments.
To provide insight into the binding mode(s), four different
binding sites of 1 with intermolecular G-quadruplex d(T2AG3T)4
(PDB code 1NP9) were investigated by molecular modeling
studies. For this calculation by software, every possible binding
angle (0–3608, including both major and minor grooves), surrounding the axis of quadruplex has been estimated, and the
results shown in Figure 7 are geometries with the lowest
energy among all these possible binding modes between each
adjacent G4 planes. The estimated interaction energy changes
for each model are shown in Table 2. It was found that when
Nuclear magnetic resonance (NMR) spectroscopy analysis
and binding sites
Nuclear magnetic resonance titration experiments were conducted to understand the binding sites of the interaction with
the G-quadruplex structures. Complex 1 was added into
a G4 A3-quadruplex prepared from a shorter oligonucleotide as
telomere sequence (G4 A3: 5’-TAGGGTTA-3’).[28] The numbering
of the residues, starting at the 5’ end of the DNA sequence, is
T1, A2, G3, G4, G5, T6, T7, and A8. The imino group resonances
of G3, G4, and G5 were at d = 11.5, 11.3, and 10.8 ppm, respectively (Figure 6), and are in agreement with a previous
report.[29] Upon the titration of complex 1, the proton signal
corresponding to the G5-imino unit first exhibited broadening
compared with the others, thus suggesting that complex 1
first binds close to the 3’-terminal face of the G-quadruplex
DNA. At a [1]/[G4 A3-quadruplex] ratio of 0.5, all the imino resonances at d = 10.8–12 ppm experienced significant broadening consistent with the multiple stacking configurations on the
top of the G-tetrad. Such differential broadening are nearly
ChemPlusChem 2012, 77, 551 – 562
Figure 7. Energy-minimized structures for possible binding sites of the complex 1 with (A) 5’-TAGGG*TTA-3’, (B) 5’-TAGG*GTTA-3’, (C) 5’-TAG*GGTTA-3’,
and (D) 5’-TA*GGGTTA-3’, where the asterisks represent binding sites. The
RuII complex is shown in yellow.
2012 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chempluschem.org
555
J. Liu et al.
Table 2. Estimated interaction energy (DE) of possible binding sites of
complex 1 with d(T2AG3T)4 by molecular modeling studies.
Binding sites
DE [kcal mol1]
5’-TAGGG*TTA-3’
5’-TAGG*GTTA-3’
5’-TAG*GGTTA-3’
5’-TA*GGGTTA-3’
52.2
43.6
46.3
44.7
the planar ligand intercalates into the G-quadruplex, the
square p-aromatic plane stacks on the 3’-terminal G-tetrads
(Figure 7 A), exhibiting a smaller binding energy (52.20 kcal
mol1) than the other three possible binding models (Figure 7 B,D). This notable result indicates that 1 preferred intercalating into the exterior 3’ surface of G-quadruplex with 1:1 stoichiometry, which is consistent with the model derived from
the results of continuous variation analysis and NMR experiments, as described in the previous section. Notably, more
energy will be required to intercalate into the G-quadruplex
more effectively for the whole complex. This result validates
the hypothesis that it is difficult for the octahedral metal complex itself to be embedded in a p-stacking unit or even in
direct proximity to the G-tetrads, but the charged molecule as
a whole interacts with grooves/loops and the phosphate backbone of quadruplexes.[31] In view of the most favorable biding
mode (Figure 7 A), we can figure out that 1,10-phenanthroline,
as a larger planar ancillary ligand, provides more possibilities
to interact with grooves/loops and the phosphate backbone of
quadruplexes in addition to the intercalative interaction and/or
multiple stacking on the G-tetrad binding. Such results may explain why complex 1, with the same positively charged center
as in complex 2, is a more efficient G-quadruplex binder.
Inducing/stabilizing the G-quadruplex structure by circular
dichroism (CD) spectroscopy
The G-rich telomeric sequence forms intra- and intermolecular
G-quadruplexes in monomeric (M), dimeric (D), and tetrameric
(T) structures through multiple methods (Scheme 2).[32] Circular
dichroism spectroscopy is one of the well-established methods
for determining the presence and to some degree the folding
of G-quadruplex structures. All guanine units in the parallel-
stranded G-quadruplex have the same anti glycosidic conformation, exhibiting a large positive band at 295 nm and a negative band at 240 nm.[33] By contrast, guanine units in the antiparallel-stranded G-quadruplex have alternating anti and syn
glycosidic conformations along each DNA strand, exhibiting
a characteristic positive band at 295 nm, a smaller negative
band at 265 nm, and a smaller positive band at 245 nm in the
CD spectra.[34] Here, complex binding studies were conducted
in the absence and presence of a stabilizing salt by CD spectroscopy to investigate the induction and conversion among
various kinds of human telomeric quadruplexes. In addition, to
measure the real CD signal, we deducted background noise of
complexes (Figure S2) and the results in Figure 8 and Figure S3
show just the CD signal from the HTG21.
In the absence of any salt, the HTG21 oligonucleotide was
dissociated partially to single-stranded molecules with a negative band centered at 238 nm, a major positive band at
257 nm, a minor negative band at 280 nm, and a positive band
near 295 nm (Figure 8 A,B, black line). The bands at 238 and
257 nm disappeared gradually upon the addition of complex 2
(0–12 mm), thereby revealing a major negative band at 260 nm
and significantly increasing the intensity of the band centered
at 295 nm (Figure 8 B). These changes are consistent with the
induction of the guanine-rich DNA in forming the antiparallel
G-quadruplex structure by complex 2. Aside from the similar
changes as for the addition of complex 2, a new and positive
band at 270 nm was observed in the CD signal upon the addition of complex 1 to the same solution (Figure 8 A). This result
indicates that complex 1 can induce HTG21 to form a hybridtype quadruplex. In addition, this striking trait suggests that
the binding of the Ru complexes to DNA causes their remarkable ability to promote different quadruplex formations (Figure 8 C).
Furthermore, CD experiments were conducted in a solution
of Na + or K + ions to determine whether the compounds could
lead to structural conversion. Figure 9 A,B show that the quadruplex molecules exist as typical parallel G-quadruplexes conformations in the presence of K + , revealing a large positive
band at 295 nm and a negative band at 240 nm.[33]Although
no significant change was found, the intensity of CD signals
corresponding to DNA was altered after the addition of complex 2 to HTG21. However, the intensity of the negative band
at 260 nm increased significantly with an increasing amount of
Scheme 2. Schematic representation of the G-quadruplex structures that can be adopted by telomeric G-strand sequences.
556
www.chempluschem.org
2012 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemPlusChem 2012, 77, 551 – 562
Bioinorganic Chemistry
Figure 8. The CD titration spectra of HTG21 (2 mm) at increasing concentrations of complexes 1 (A) and 2 (B) in 10 mm Tris-HCl buffer, pH 7.40, 25 8C.
Arrows indicate the increasing amounts of complexes. C) Representative illustration of the two RuII complexes inducing the single-stranded human telomeric DNA into different G-quadruplexes.
complex 1, and a positive band at 270 nm started to appear.
This result indicates that complex 1 can convert the parallel
quadruplex into an obvious hybrid-type G-quadruplexes, thus
signifying that this complex interacts more preferentially with
the hybrid-type quadruplexes than does complex 2. Both complexes could increase the intensity of the CD signals corresponding to HTG21 in a 100 mm solution of K + , probably
caused by the structural conversion between the intra- and intermolecular G-quadruplexes,[35] with reference to the following
gel mobility shift assay (EMSA).
By contrast, the addition of increasing amounts of complexes 1 and 2 to HTG21 in 100 mm NaCl buffer resulted in no
significant changes in the CD spectra (Figure S3). This result
implies that the conformation of the G-quadruplex is stabilized
by Na + , and neither of the two Ru complexes could change
the conformation of the G-quadruplex at high ionic
strength.[36]
ChemPlusChem 2012, 77, 551 – 562
Figure 9. The CD titration spectra of HTG21 (2 mm) at increasing concentrations of complexes 1 (A) and 2 (B) in 10 mm Tris-HCl buffer, 100 mm KCl,
pH 7.40, 25.0 8C. Arrows indicate the increasing amounts of complexes. Effects of complexes 1 (C) and 2 (D) on the assembly of the telo21 structure illustrated by PAGE analysis in 10 mm Tris-HCl (pH 7.40) containing 100 mm
KCl. Major bands were identified as monomer (M) and dimer (D). E) Representative illustration of complexes converting the formation between the
intra- and intermolecular structures in the presence of 100 mm KCl.
Specificity for G-quadruplex DNA by the electrophoretic mobility shift assay (EMSA)
The EMSA was performed to further identify whether the complexes can facilitate the formation of G-quadruplexes from the
oligonucleotide HTG21 or converted the formation between
the intra- and intermolecular structures. In the absence of Ru
2012 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chempluschem.org
557
J. Liu et al.
complexes, only the band corresponding to the monomer
could be observed in the electrophoretic mobility shift assays
(Figure 9 C,D), in accordance with the previous gel-shift data.[37]
The increasing amounts of Ru complexes to HTG21 oligonucleotide resulted in the progressive appearance of new bands
with reduced mobility, corresponding to the dimer forms. According to the quantification of the gels in the Figure S4, complex 1 gave the dimer band at 5 mm, whereas complex 2 did at
a higher concentration (10 mm). In addition, complex 1 gave
more of the dimer band (35 %) at 30 mm than complex 2
(25 %). These result reveal that both Ru complexes can efficiently promote different intermolecular quadruplex formation
at high K + (Figure 9 E), and complex 1 has a stronger quadruplex affinity and more preference for the hybrid-type quadruplex structure than complex 2. This observation is in good
agreement with the CD results in a solution of K + ions and the
G-quadruplex stabilizing effects shown in other methods.
In vitro inhibition of the telomerase activity by the telomerase repeat amplification protocol (TRAP) assay
The TRAP assay is commonly used in evaluating telomerase activity in tissues or cell extracts and in determining the inhibitory properties of small molecules against telomerase.[38] The
complexes exhibited high binding affinity and stabilizing ability
to telomeric G-quadruplexes. Therefore, the telomerase inhibition of the complexes was investigated using the TRAP assay.
In this experiment, solutions of complex 1 or 2 were added to
the telomerase reaction mixture containing an extract from
cracked HepG2 cell lines. The results of the telomerase activity
are listed in Figure 11. At a concentration of 5–10 mm for com-
Stabilizing the G-quadruplex structure by the polymerase
chain reaction (PCR)-stop assay
A PCR-stop assay was used to ascertain whether the Ru complexes could bind to a test oligomer HTG21 and therefore stabilize the G-quadruplex structure. The sequences of HTG21
and its corresponding complementary sequence (Rev 21G) can
hybridize a final double-stranded DNA PCR product when used
with Taq DNA polymerase as the catalyst. However, in the presence of some G-quadruplex stabilizers, the template sequence
HTG21 was induced into a G-quadruplex structure that blocked
the hybridization and the detection of the final PCR product.
Figure 10 illustrates the inhibitory properties of the complexes
Figure 10. Effect of complexes 1 and 2 on the hybridization of HTG21 by
the PCR-stop assay.
to a PCR process with similar concentration gradients. The inhibitory effect increased with increasing concentration. Complex 1 showed an inhibitory effect on telo21 at only 15 mm,
whereas the effect of complex 2 was evident at 25 mm. The
IC50 values (the concentrations that inhibited hybridization by
50 %) of complexes 1 and 2 are 7.77 and 13.28 mm, respectively.
A correlation between the PCR-stop assay results and FRET
data could be drawn. Complexes with greater G-quadruplex
structure-stabilizing power are generally better inhibitors of
amplification in HTG21. Thus, complex 1 is a better G-quadruplex binder.
Figure 11. The influence of complexes 1 and 2 to the telomere activity of
HepG2.
plex 1, the TRAP assay revealed a dose-dependent inhibition of
the telomere, and the number of bands clearly decreased with
respect to the control. By contrast, no complete inhibition was
observed in the presence of complex 2, even at 12.5 mm concentration. Complex 1 (TelIC50 = 5.8 mm) exhibited a better inhibitory effect on the telomerase activity compared with complex 2 (TelIC50 > 12.5 mm). This result not only agrees with the
experimental data from the aforementioned studies, but also
indicates that complex 1 might be a potential human telomerase inhibitor.
Cytotoxicity test by the 3-(4,5-dimethylthiazol-2-yl)-2, 5-diphenyltetrazolium bromide (MTT) assay
The cytotoxicity of the Ru complexes against HepG2, SW620,
A549, MDA-MB-231, and NIH/3T3 cell lines were determined by
the MTT assay. Details of the results are given in Table 3 and
Figure 12. Complexes exhibited a differential cytotoxicity
Table 3. Cytotoxic effects of Ru complexes and Cisplatin on human cancer
cell lines.
Complex
HepG2
SW620
IC50 values [mm][a]
A549
MDA-MB231
NIH/3T3
1
11.21 2.02 27.43 3.23 25.56 3.52 19.90 1.42 > 100
2
15.56 2.83 54.38 2.26 34.34 3.74 40.37 1.21 59.31 2.36
Cisplatin 13.61 1.13 30.02 1.81 3.23 0.27 –
19.72 1.59
[a] The values are expressed as the mean standard deviation (triplicates).
558
www.chempluschem.org
2012 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemPlusChem 2012, 77, 551 – 562
Bioinorganic Chemistry
a better telomeric quadruplex stabilizer and telomerase inhibitor than complex 2. These notable differences in the two complexes prove that planarity and p-delocalized system are vital
for the G-quadruplex recognition in binding. Furthermore,
complex 1 exhibits an antitumor function by locking a telomeric DNA into a G-quadruplex conformation that cannot be extended by telomerase. Ruthenium complexes definitely represent an exciting new strategy in designing new antitumor
agents for targeting DNA quadruplexes.
Figure 12. IC50 values of Ru complexes and Cisplatin on human cancer cell
lines.
against cancer cells after 2 days. In particular, we found the
IC50 values of the Ru complexes on NIH/3T3 from 59.31 mm to
greater 100 mm were significantly higher than those of Cisplatin (19.72 mm). This finding suggests that this series of complexes inhibited the growth of cancer cell lines better than
normal cells and the RuII complexes were much less toxic
toward normal cells. Complex 1 exhibited a broad spectrum of
inhibition on human cancer cells, with IC50 values ranging from
11.21 to 27.43 mm. More importantly, as evidenced by the
lower IC50 values, complex 1 exhibited much higher antiproliferative activities on the cancer cell lines than complex 2. Notably, complex 1 showed a distinct preference for the HepG2
cells (IC50 = 11.21 mm, lower than Cisplatin in killing HepG2
cells).This result, combined with the TRAP assay, suggests that
the cytotoxicity of this metal complex may be attributed to its
role in telomerase inhibition.
Conclusion
Two RuII polypyridyl complexes as telomeric quadruplex stabilizers have been synthesized and evaluated using biophysical
and biochemical studies. The CD data reveal that complex 1
selectively induces the formation of hybrid-type quadruplexes,
whereas complex 2 could induce antiparallel G-quadruplexes
in the absence of any salt and keep parallel G-quadruplexes in
a solution of K + ions. Absorption and emission studies suggest
that complex 1 can bind to the G-quadruplex with larger binding constants. The FRET and thermal denaturation studies
imply that complex 1, increasing the DTm value by 18.8 8C at
2 mm, is a more effective quadruplex-DNA stabilizer. In addition, continuous variation analyses, NMR spectroscopy, and
molecular modeling were used to investigate the multiple interaction (mainly intercalating) of complex 1 at the 3’-terminal
face through a 1:1 [quadruplex]/[complex] binding mode ratio.
The biochemical studies, including the PCR-stop and EMSA
assays, further demonstrate that both complexes can promote
HTG21 into the dimer forms. Complex 1 inhibits the telomerase activity in a cell-free system by the TRAP assay. The MTT
assay determines that complex 1 is moderately cytotoxic
(IC50 = 11.21 mm) in the HepG2 cell lines.
Hence, complex 1 containing a larger planar ancillary ligand
such as 1,10-phenanthroline, which can interact with grooves/
loops and the phosphate backbone of quadruplexes, is
ChemPlusChem 2012, 77, 551 – 562
Experimental Section
CAUTION! Metal perchlorates are potentially explosive and
should therefore be handled with great care.
Reagents and materials
The oligomers or primers used in this study were purchased from
Sangon (Shanghai, China) and used without further purification.
The DNA oligomers include HTG21 = 5’-G3(T2AG3)3-3’, G4 A3 = 5’TAGGGTTA-3’, double-stranded competitor (ds26 = 5’-CAATCGGATCGAATTCGATCCGATTG-3’, and F21T = 5’-FAM-d(G3[T2AG3]3)TAMRA-3’, of which the donor fluorophore FAM is 6-carboxy-fluorescein and the acceptor fluorophore TAMRA is 6-carboxytetramethylrhodamine. Concentrations of these oligomers were determined by measuring the absorbance at 260 nm after melting.
Single-strand extinction coefficients were calculated from mononucleotide data using a nearest-neighbor approximation.[39] The formations of intramolecular G-quadruplexes were carried out as follows: the oligonucleotide samples were annealed in different buffers at 95 8C for 5 min, slowly cooled to RT, and then incubated at
4 8C for 12 h. Buffer A: 10 mm tris(hydroxymethyl)aminomethaneHCl (Tris-HCl), pH 7.40; buffer B: 10 mm Tris-HCl, 100 mm NaCl,
pH 7.40; buffer C: 10 mm Tris-HCl, 100 mm KCl, pH 7.40. Other reagents and solvents were purchased commercial sources unless
otherwise specified. Doubly distilled water was used to prepare
buffer solutions.
Physical measurement
Electrospray ionization mass spectra (ESI-MS) were acquired by
using a Thermo Finnigan LCQ DECA XP ion-trap mass spectrometer, equipped with an ESI source. The 1H NMR spectra were acquired on a Varian 400 MHz spectrometer with (CD3)2SO as solvent
at RT, and all chemical shift values were given relative to tetramethylsilane (TMS). Elemental analyses for C, H, and N were carried
out with a PerkinElmer 240 8C elemental analyzer and purity of all
target compounds used in the biophysical and biological studies
was > 95 %. The absorption spectra in the UV/Vis region were recorded on a Varian Cary 300 spectrophotometer. Emission spectra
of the synthetic Ru complexes were measured on a Shimadzu RF5000 fluorescence spectrophotometer with excitation at 468 nm.
The CD spectra were measured on a JASCO J-810 spectropolarimeter and high-frequency noise was filtered out using JASCO J600 software.
Synthesis and characterization
1,10-phenanthroline-5,6-dione, [Ru(phen)2Cl2]·2H2O, and [Ru(bpy)2Cl2]·2H2O were prepared and characterized according to the
literature.[40]
2012 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chempluschem.org
559
J. Liu et al.
Synthesis of 2-thiophenimidazo[4,5-f][1,10]phenanthroline
(TIP)
The ligand was prepared by using modified literature procedures.[41] A mixture of 1,10-phenanthroline-5,6-dione (0.21 g,
1 mmol), 2-thiophene-carboxaldehyde (0.14 g, 1.2 mmol), ammonium acetate (0.15 g, 20 mmol), and glacial acetic acid (20 mL) was
heated at reflux for 4 h. The cooled deep-red solution was diluted
with water (40 mL) and neutralized with ammonium hydroxide to
give a yellow precipitate. The precipitate was collected and purified by column chromatography on silica gel (60–100 mesh) with
ethanol as eluent to give the ligand as yellow powder (yield of
0.25 g, 85 %). ESI-MS (MeOH) m/z: 302.0 (100, [M + H] + ); elemental
analysis (%) calcd for C17H10N4S: C 67.53, H 3.33, N 18.53; found:
C 67.52, H 3.41, N 18.60.
Synthesis of [Ru(phen)2(TIP)] (ClO4)2 (1)
Complex 1 was synthesized by using a modified literature procedure.[42] [Ru(phen)2Cl2]·2H2O (0.29 g, 0.5 mmol) was dissolved in glacial acetic acid (20 mL) and tip (0.18 g, 0.6 mmol) was added. The
mixture was heated at reflux for 8 h under argon. Upon cooling,
a red precipitate was obtained by dropwise addition of saturated
aqueous NaClO4 solution. The precipitated complex was isolated
by filtration and air-dried, then purified by chromatography on alumina (200 mesh) with acetonitrile (ACN)/toluene (4:1, v/v) as an
eluent to give the complex as red (yield of 0.27 g, 56 %). The
sample shows good solubility in solvents such as ACN, dimethyl
sulfoxide (DMSO), and acetone. 1H NMR ([D6]DMSO): d = 8.94 (d,
2 H), 8.76 (d, 4 H), 8.39 (s, 4 H), 8.10–8.12 (m, 4 H), 7.89 (d, 3 H), 7.75–
7.79 (m, 4 H), 7.70 (t, 2 H), 7.65 (d, 1 H), 7.24 ppm (t, 1 H); UV/Vis
(ACN; l (e)): 457 (32400), 288 (84700), 262 nm (149500 m1 cm1);
ESI-MS (ACN) m/z: 864.9 ([MClO4] + ), 763.2 ([M2ClO4H] + ), 382.3
([M2ClO4]2 + ); elemental analysis (%) calcd for C41H26N8Cl2O8RuS:
C 51.15, H 2.72, N 11.64; found: C 51.13, H 2.81, N 11.56. See NMR
and ESI-MS spectra in the Supporting Information.
Complex 2 was prepared by the same method as above but with
[Ru(bpy)2Cl2]·2H2O (0.242 g, 0.5 mmol) to give a yield of 0.228 g,
50 %. 1H NMR ([D6]DMSO): d = 8.98 (d, 2 H), 8.87 (d, 2 H), 8.83 (d,
2 H), 8.22 (t, 2 H), 8.11 (t, 2 H), 7.93 (d, 2 H), 7.82–7.87 (m, 5 H), 7.64
(d, 1 H), 7.59 (t, 4 H), 7.37 (t, 2 H), 7.24 ppm (t, 1 H); UV/Vis (ACN;
l (e)): 462 (17100), 286 (66800), 244 nm (40500 m1 cm1); ESI-MS
(ACN): m/z = 814.7 ([MClO4] + ), 715.1 ([M2ClO4-H] + ), 358.3
([M2ClO4]2 + ); elemental analysis (%) calcd for C37H26N8Cl2O8RuS:
C 48.58, H 2.87, N 12.25; found: C 48.62, H 2.77, N 12.18. See NMR
and ESI-MS spectra in the Supporting Information.
Absorption and emission spectra
Analysis of absorption and emission spectra are the most common
ways to investigate the interactions of complexes with DNA. The titration was performed by using a fixed complex concentration
(5 mm in buffer C) to which increments of the DNA stock solution
were added at RT. The volume of the complex was 3000 mL. Solutions of complex DNA were incubated for 5 min before absorption
spectra were recorded. The titration processes were repeated several times until no change was observed in the spectra, indicating
that binding saturation was achieved. Changes in the concentration of metal complexes as a result of dilution at the end of each
www.chempluschem.org
Continuous variation analysis
Continuous variation analysis was performed by using a Shimadzu
RF-5000 fluorescence spectrophotometer following previously
published procedures.[25] Each complex and HTG21 were prepared
as 10 mm solution in Tris-HCl buffer containing 100 mm KCl. Two
series of solutions were used for the experiments: one with varying
mole fractions of complex and HTG21, and another one with varying concentrations of complex. The sum of the complex and
HTG21 concentrations was always 10 mm. Their emission spectrum
was collected from 500 to 750 nm using a quartz cell with a path
length of 1 cm at 25 8C. The DI values were calculated by subtracting the fluorescence intensity of complex solution without HTG21
from the fluorescence intensity of corresponding complex solution
with HTG21 at lmax. This value was plotted versus the complex
mole fraction to generate a Job plot. Binding stoichiometries were
obtained from the intercepts of the linear plot obtained by linear
least-squares fits to the left- and right-hand portions of the Job
plots. Final analysis of the data was carried out using Origin 7.5
software (Origin Lab Corp.).
Thermal denaturation study
The thermal denaturation study was carried by using a JASCO J810 spectropolarimeter equipped with a Peltier temperature-controlling programmer PTP-6. With the use of the thermal melting
program, the temperature of the cell containing the cuvette was
ramped from 40 to 100 8C. Thermal melting curves and calculation
of DTm values were performed in Tris-KCl buffer with 2 mm HTG21
in the presence or absence of 20 mm Ru complexes. The CD spectrum at 295 nm was monitored at a rate of 1 8C min1, and measured every 1 8C.
FRET assay
Synthesis of [Ru(bpy)2(TIP)] (ClO4)2 (2)
560
titration were negligible. The titrations for each sample were repeated at least three times.
The fluorescent-labeled oligonucleotide F21T, was prepared as
a 100 mm solution in buffer C and then annealed by heating to
90 8C for 5 min, and subsequently cooled to RT overnight. Fluorescence melting curves were determined by using a Bio-Rad iQ5
real-time PCR detection system, using a total reaction volume of
20 mL, with 0.2 mm of labeled oligonucleotide and different concentrations of complexes. Fluorescence readings with excitation at
470 nm and detection at 530 nm were taken at intervals of 1 8C
over the range 37–99 8C, with a constant temperature being maintained for 30 s prior to each reading to ensure a stable value. The
melting of the G-quadruplex was monitored alone or in the presence of various concentrations of complexes and/or of doublestranded competitor ds26. Final analysis of the data was carried
out using Origin 7.5 software (Origin Lab Corp.).
NMR experiments
The 1H NMR experiments were performed by using a Bruker
AVANCE 400 spectrometer with a typical acquisition condition of
45 pulse length, 2.0 s relaxation delay, 16 K data points, 16–32 transients. The stock solutions of G4 A3 G-quadruplex were prepared
by dissolving the sample in 90 % H2O/10 % D2O with 150 mm KCl,
25 mm KH2PO4, and 1 mm ethylenediaminetetraacetic acid (EDTA),
pH 7.4. Aliquots of stock solutions containing complex were titrat-
2012 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemPlusChem 2012, 77, 551 – 562
Bioinorganic Chemistry
ed directly into the DNA solution inside an NMR tube. The 1H NMR
spectra were recorded at 300 K by the pulse program with gradients for water suppression.
Molecular docking calculations
A model study on the interaction between complex 1 and G-quadruplex DNA was performed by using Accelrys Discovery Studio 2.1
Software. The CHARMm force-field was used in the input of the
calculations. The correction of the partial charge distribution for all
atoms in the RuII complexes was made following the output from
Gaussian 03.[43] Model building and optimization were performed
using the model building and energy-minimization modules of the
Accelrys Software Package. The DNA crystal structures for
d(T2AG3T)4 (PDB code 1NP9)[44] was downloaded from the Protein
Data Bank. The docking options were then performed to find the
most stable and favorable orientation. The binding site was assigned across the all of the minor and major grooves of the DNA
molecule. Interaction energies of Ru–DNA complexes can be estimated by calculating the difference between their total energies
and the sum of lowest energies found for the optimized structures
of free DNA and binuclear complex. The negative of the interaction
energy is the binding energy, DE = TEsum of the individual
energy (BE), where DE is the interaction energy, TE is the total
energy of DNA/complex, and BE is the binding energy.[45]
Circular dichroism study
All CD experiments were measured at RT using a quartz cell with
a path length of 1 cm. The spectra were collected from 220 to
400 nm and with a scanning speed of 200 nm min1. The bandwidth was 5 nm, and the response time was 2 s. The CD titration
was performed at a fixed HTG21 concentration (2 mm) with various
concentrations (0–5 mol equiv) of the complexes in different buffers; (a) 10 mm Tris-HCl, pH 7.4; (b) 100 mm NaCl, 10 mm Tris-HCl,
pH 7.4; (c) 100 mm KCl,10 mm Tris-HCl, pH 7.4. After each addition
of Ru complex, the reaction was stirred and allowed to equilibrate
for at least 5 min (until no elliptic changes were observed) and
a CD spectrum were measured. During the experiment, the instrument was flushed continuously with pure nitrogen.
Polymerase chain reaction (PCR)-stop assay
The PCR-stop assay was carried out according to a modified protocol of a previous study.[46] The test oligomers HTG21 and the corresponding
complementary
sequences
Rev 21 G
(ATCGCT2CTCGTC3TA2C2) were used in the current study. The reactions were performed in 1 PCR buffer, containing 10 pmol of each
oligonucleotide, 0.16 mm dNTP, 2.5 U Taq polymerase, and different
concentrations of complexes. Reaction mixtures were incubated in
a thermocycler with the following cycling conditions: 94 8C for
3 min, followed by 30 cycles of 94 8C for 30 s, 58 8C for 30 s, and
72 8C for 30 s. The products generated from PCR were then analyzed on 15 % nondenaturing polyacrylamide gels in 1 TBE and
silver stained. The photos were taken and IC50 values were calculated.
KCl. After the DNA was cooled to RT, a 10 mL stock solution of
metal complex was added to each sample to produce the specified
concentrations at a total volume of 20 mL. The reaction mixture
was incubated for 12 h at RT. Each mixture was added 4 mL of loading buffer (50 % glycerol, 0.25 % bromophenol blue, and 0.25 %
xylene cyanol) and analyzed on a 20 % native polyacrylamide gel
electrophoresis (PAGE; the gel was pre run for 30 min). Electrophoresis proceeded for 2 h in TBE (Tris-borate EDTA) running buffer.
The gels were silver-stained according to a previously reported
protocol.[47]
TRAP assay
The TRAP assay was carried out according to a modified protocol
of a previous study.[48] Telomerase extraction from HepG2 cells. The
TRAP assay was performed in two steps: 1) telomerase-mediated
extension of the forward primer (TS: 5’-AATCCGTCGAGCAGAGTT-3’)
contained in a 20 mL reaction mixture comprising TRAP buffer
(20 mm, pH 8.3), 68 mm KCl, 1.5 mm MgCl2, 1 mm EGTA, 0.05 % v/v
Tween-20, 0.05 mg of bovine serum albumin, 50 mm of each deoxynucleotide triphosphate, 0.1 mg of TS primer, and 3 mCi of [R32P]dCTP. The protein (0.04 mg) was incubated with the reaction
mixture agent (acid addition and quaternary dimethiodide salts)
at final concentrations of up to 50 mm for 20 min at 25 8C. Analysis
buffer (no protein) control, heat-inactivated protein control, and
50 % protein (0.02 mg) control were included in each assay. 2) While
the mixture was being heated at 80 8C in a polymerase chain reaction (PCR) block of a thermal cycler for 5 min to inactivate telomerase activity, 0.1 mg of reverse CX primer (3’-AATCCCATTCCCATTCCCATTCCC-5’) and two units of Taq DNA polymerase were added. A
three-step PCR was then performed: 94 8C for 30 s, 50 8C for 30 s,
and 72 8C for 1 min for 31 cycles. Telomerase-extended PCR products in the presence or absence of complexes were determined by
electrophoretic separation using 8 % w/w acrylamide denaturing
gels.
MTT assay
HepG2, SW620, A549, MDA-MB-231, and NIH/3T3 were seeded on
96-well plates (1.0 103 per well) and exposed to various concentrations of complexes. The microplate was incubated for 48 h at
37 8C, 5 % CO2, and 95 % air in a humidified incubator. After incubation, 10 mL of MTT reagent (5 mg mL1) was added to each well
and further incubated for 2 h. The cells in each well were then
treated with DMSO (150 mL for each well) and the optical density
(OD) was recorded at 570 nm. All drug doses were parallel tested
in triplicate, and the IC50 values were derived from the mean OD
values of the triplicate tests versus drug concentration curves.[49]
Acknowledgements
This study was supported by the National Natural Science Foundation of China (20871056, 21171070), the Planned Item of Science and Technology of Guangdong Province (c1011220800060),
the Fundamental Research Funds for the Central Universities, the
Planned Item of Science, and the Fundamental Research Foundation of Shenzhen City (Major Project; JC201005250058A).
Gel mobility shift assay
The formation of G-quadruplex DNA was carried out as described
in the literature.[47] Briefly,10 mm the oligomer (HTG21) was heated
at 95 8C for 10 min in 10 mm Tris-HCl (pH 7.40) containing 100 mm
ChemPlusChem 2012, 77, 551 – 562
Keywords: antitumor agents · bioinorganic chemistry · DNA ·
G-quadruplex DNA · ruthenium complexes · telomerase
2012 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chempluschem.org
561
J. Liu et al.
[1] a) J. T. Davis, Angew. Chem. 2004, 116, 684 – 716; Angew. Chem. Int. Ed.
2004, 43, 668 – 698; b) J. L. Huppert, Chem. Soc. Rev. 2008, 37, 1375 –
1384; c) J. T. Wang, Y. Li, J. H. Tan, L. N. Ji, Z. W. Mao, Dalton Trans. 2011,
40, 564 – 566.
[2] a) D. Monchaud, M. P. Teulade-Fichou, Org. Biomol. Chem. 2008, 6, 627 –
636; b) S. Burge, G. N. Parkinson, P. Hazel, A. K. Todd, S. Neidle, Nucleic
Acids Res. 2006, 34, 5402.
[3] E. H. Blackburn, Nature 1991, 350, 569 – 573.
[4] a) C. B. Harley, A. B. Futcher, C. W. Greider, Nature 1990, 345, 458 – 460;
b) M. Y. Kim, H. Vankayalapati, K. Shin-ya, K. Wierzba, L. H. Hurley, J. Am.
Chem. Soc. 2002, 124, 2098 – 2099.
[5] L. R. Kelland, Anti-Cancer Drugs 2000, 11, 503 – 513.
[6] N. W. Kim, M. A. Piatyszek, K. R. Prowse, C. B. Harley, M. D. West, P. L. Ho,
G. M. Coviello, W. E. Wright, S. L. Weinrich, J. W. Shay, Science 1994, 266,
2011 – 2015.
[7] a) C. I. Nugent, V. Lundblad, Genes Dev. 1998, 12, 1073 – 1085; b) A.
De Cian, L. Lacroix, C. Douarre, N. Temime-Smaali, C. Trentesaux, J. F.
Riou, J. L. Mergny, Biochimie 2008, 90, 131 – 155.
[8] a) T. Ou, Y. Lu, J. Tan, Z. Huang, K. Y. Wong, L. Gu, ChemMedChem 2008,
3, 690 – 713; b) J. L. Zhou, Y. J. Lu, T. M. Ou, J. M. Zhou, Z. S. Huang, X. F.
Zhu, C. J. Du, X. Z. Bu, L. Ma, L. Q. Gu, J. Med. Chem. 2005, 48, 7315 –
7321; c) E. M. Rezler, J. Seenisamy, S. Bashyam, M. Y. Kim, E. White, W. D.
Wilson, L. H. Hurley, J. Am. Chem. Soc. 2005, 127, 9439 – 9447.
[9] a) S. T. D. Hsu, P. Varnai, A. Bugaut, A. P. Reszka, S. Neidle, S. Balasubramanian, J. Am. Chem. Soc. 2009, 131, 13399 – 13409; b) B. Rubis, M.
Kaczmarek, N. Szymanowska, E. Galezowska, A. Czyrski, B. Juskowiak, T.
Hermann, M. Rybczynska, Invest. New Drugs 2009, 27, 289 – 296.
[10] K. L. Haas, K. J. Franz, Chem. Rev. 2009, 109, 4921 – 4960.
[11] a) R. T. Wheelhouse, D. Sun, H. Han, F. X. Han, L. H. Hurley, J. Am. Chem.
Soc. 1998, 120, 3261 – 3262; b) F. Gao, H. Chao, L. N. Ji, Chem. Biodiversity
2008, 5, 1962 – 1979.
[12] L. Xu, G. L. Liao, X. Chen, C. Y. Zhao, H. Chao, L. N. Ji, Inorg. Chem.
Commun. 2010, 13, 1050 – 1053.
[13] J. E. Reed, A. A. Arnal, S. Neidle, R. Vilar, J. Am. Chem. Soc. 2006, 128,
5992 – 5993.
[14] S. Shi, X. Geng, J. Zhao, T. Yao, C. Wang, D. Yang, L. Zheng, L. Ji, Biochimie 2010, 92, 370 – 377.
[15] S. Shi, J. Liu, T. Yao, X. Geng, L. Jiang, Q. Yang, L. Cheng, L. Ji, Inorg.
Chem. 2008, 47, 2910 – 2912.
[16] T. Wilson, M. P. Williamson, J. A. Thomas, Org. Biomol. Chem. 2010, 8,
2617 – 2621.
[17] A. Ghosh, P. Das, M. R. Gill, P. Kar, M. G. Walker, J. A. Thomas, A. Das,
Chem. Eur. J. 2011, 17, 2089 – 2098.
[18] L. Xu, D. Zhang, J. Huang, M. Deng, M. Zhang, X. Zhou, Chem. Commun.
2010, 46, 743 – 745.
[19] C. Rajput, R. Rutkaite, L. Swanson, I. Haq, J. A. Thomas, Chem. Eur. J.
2006, 12, 4611 – 4619.
[20] C. Wei, G. Jia, J. Yuan, Z. Feng, C. Li, Biochemistry 2006, 45, 6681 – 6691.
[21] J. Sun, Y. An, L. Zhang, H. Y. Chen, Y. Han, Y. J. Wang, Z. W. Mao, L. N. Ji,
J. Inorg. Biochem. 2011, 105, 149 – 154.
[22] S. Satyanarayana, J. C. Dabrowiak, J. B. Chaires, Biochemistry 1992, 31,
9319 – 9324.
[23] I. Haq, J. O. Trent, B. Z. Chowdhry, T. C. Jenkins, J. Am. Chem. Soc. 1999,
121, 1768 – 1779.
[24] a) M. R. Gill, J. A. Thomas, Chem. Soc. Rev., 2012, 41, 3179 – 3192; b) Y. Li,
Z. Y. Yang, J. C. Wu, Eur. J. Med. Chem. 2010, 45, 5692 – 5701; c) S. N.
562
www.chempluschem.org
Georgiades, N. H. Abd Karim, K. Suntharalingam, R. Vilar, Angew. Chem.
2010, 122, 4114 – 4128; Angew. Chem. Int. Ed. 2010, 49, 4020 – 4034.
[25] J. Dash, Z. A. E. Waller, G. D. Panto, S. Balasubramanian, Chem. Eur. J.
2011, 17, 4571 – 4581.
[26] A. D. Moorhouse, A. M. Santos, M. Gunaratnam, M. Moore, S. Neidle,
J. E. Moses, J. Am. Chem. Soc. 2006, 128, 15972 – 15973.
[27] J. L. Mergny, L. Lacroix, M. P. Teulade-Fichou, C. Hounsou, L. Guittat, M.
Hoarau, P. B. Arimondo, J. P. Vigneron, J. M. Lehn, J. F. Riou, Proc. Natl.
Acad. Sci. USA 2001, 98, 3062 – 3067.
[28] D. L. Ma, C. M. Che, S. C. Yan, J. Am. Chem. Soc. 2008, 130, 1835 – 1846.
[29] E. Gavathiotis, R. A. Heald, M. F. G. Stevens, M. S. Searle, Angew. Chem.
2001, 113, 4885 – 4887; Angew. Chem. Int. Ed. 2001, 40, 4749 – 4751.
[30] A. T. Phan, V. Kuryavyi, H. Y. Gaw, D. J. Patel, Nat. Chem. Biol. 2005, 1,
167 – 173.
[31] P. Murat, Y. Singh, E. Defrancq, Chem. Soc. Rev., 2011, 40, 5293 – 5307.
[32] W. Peti, J. Meiler, R. Brschweiler, C. Griesinger, J. Am. Chem. Soc. 2002,
124, 5822 – 5833.
[33] R. Jin, B. L. Gaffney, C. Wang, R. A. Jones, K. J. Breslauer, Proc. Natl. Acad.
Sci. USA 1992, 89, 8832 – 8836.
[34] V. Dapic, V. Abdomerovi, R. Marrington, J. Peberdy, A. Rodger, J. O.
Trent, P. J. Bates, Nucleic Acids Res. 2003, 31, 2097 – 2107.
[35] C. T. Lin, T. Y. Tseng, Z. F. Wang, T. C. Chang, J. Phys. Chem. B. 2011, 115,
2360 – 2370.
[36] A. Ambrus, D. Chen, J. Dai, T. Bialis, R. A. Jones, D. Yang, Nucleic Acids
Res. 2006, 34, 2723 – 2735.
[37] H. Han, D. R. Langley, A. Rangan, L. H. Hurley, J. Am. Chem. Soc. 2001,
123, 8902 – 8913.
[38] D. Gomez, J. L. Mergny, J. F. Riou, Cancer Res. 2002, 62, 3365 – 3368.
[39] M. E. Reichmann, S. A. Rice, C. A. Thomas, P. Doty, J. Am. Chem. Soc.
1954, 76, 3047 – 3053.
[40] S. Shi, J. Liu, J. Li, K. C. Zheng, X. M. Huang, C. P. Tan, L. M. Chen, L. N. Ji,
J. Inorg. Biochem. 2006, 100, 385 – 395.
[41] J. Liu, W. Zheng, S. Shi, C. Tan, J. Chen, K. Zheng, L. Ji, J. Inorg. Biochem.
2008, 102, 193 – 202.
[42] W. J. Mei, J. Liu, K. C. Zheng, L. J. Lin, H. Chao, A. X. Li, F. C. Yun, L. N. Ji,
Dalton Trans. 2003, 1352 – 1359.
[43] A. K. Patra, T. Bhowmick, S. Roy, S. Ramakumar, A. R. Chakravarty, Inorg.
Chem. 2009, 48, 2932 – 2943.
[44] E. Gavathiotis, M. S. Searle, Org. Biomol. Chem. 2003, 1, 1650 – 1656.
[45] R. Bhowmik, K. S. Katti, D. Katti, Polymer 2007, 48, 664 – 674.
[46] B. Fu, D. Zhang, X. Weng, M. Zhang, H. Ma, Y. Ma, X. Zhou, Chem. Eur. J.
2008, 14, 9431 – 9441.
[47] a) J. L. Zhou, Y. J. Lu, T. M. Ou, J. M. Zhou, Z. S. Huang, X. F. Zhu, C. J. Du,
X. Z. Bu, L. Ma, L. Q. Gu, J. Med. Chem. 2005, 48, 7315 – 7321; b) D. Sun,
R. Zhang, F. Yuan, D. Liu, Y. Zhou, J. Liu, Dalton Trans., 2012, 41, 1734 –
1741.
[48] a) H. L. Huang, Y. J. Liu, C. H. Zeng, J. H. Yao, Z. H. Liang, Z. Z. Li, F. H.
Wu, J. Mol. Struct. 2010, 966, 136 – 143; b) D. Sun, Y. N. Liu, D. Liu, R.
Zhang, X. Yang, J. Liu, Chem. Eur. J. 2012, 18, 4285 – 4295.
[49] Y. Ma, T. M. Ou, J. H. Tan, J. Q. Hou, S. L. Huang, L. Q. Gu, Z. S. Huang,
Eur. J. Med. Chem. 2011, 46, 1906 – 1913.
Received: February 29, 2012
Revised: April 17, 2012
Published online on May 21, 2012
2012 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemPlusChem 2012, 77, 551 – 562