Surface confined heteroleptic copper(ii)-polypyridyl complexes for photonuclease activity

Article abstract of DOI:10.1039/c4cc05063k

Heteroleptic copper(II)-polypyridyl complexes with extended π-conjugated, aromatic terminal units were immobilized on glass/Si substrates to intercalate DNA and cleave it upon photoexposure. Photonuclease activity is shown to be high, well reproducible and non-destructible towards the assembled complexes.

Full text of DOI:10.1039/c4cc05063k

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Surface confined heteroleptic copper(II)–polypyridyl
complexes for photonuclease activity†‡
Cite this: Chem. Commun., 2014,
0, 11484
5
a
ab
a
c
Vikram Singh, Prakash C. Mondal, Anup Kumar, Yekkoni L. Jeyachandran,
a
d
c
Received 2nd July 2014,
Accepted 3rd August 2014
Satish K. Awasthi, Rinkoo D. Gupta* and Michael Zharnikov*
DOI: 10.1039/c4cc05063k
www.rsc.org/chemcomm
Heteroleptic copper(II)–polypyridyl complexes with extended
p-conjugated, aromatic terminal units were immobilized on glass/Si
substrates to intercalate DNA and cleave it upon photoexposure.
Photonuclease activity is shown to be high, well reproducible and
non-destructible towards the assembled complexes.
Cu(II)–polypyridyl complexes are frequently used for photoinduced
DNA cleavage due to suitable redox, spectral, and structural
1
properties. The respective experiments are, however, usually
performed in solution, whereas immobilization of these complexes
2
on solid substrates for nuclease activity is largely unexplored.
3
Analogous studies have only been reported for fullerene or
lomefloxacin based assemblies, whereas no supported systems
4
containing metal–Cu(II) complexes were addressed in context of
photonuclease activity. Moreover, the available DNA binding and Fig. 1 Schematic of complexes 1 and 2 immobilized on Si/glass substrates via
photocleavage results are mostly based on optical (UV/Vis) siloxane based attachment chemistry and forming monolayers M1 and M2.
experiments and lack most conclusive gel electrophoresis evidence
3,4
for formation of DNA fragments–oligonucleotides. In this context,
we describe here two novel heteroleptic Cu(II)–polypyridyl complexes designed for efficient intercalation into a DNA strand. DNA
confined to glass/Si supports via robust siloxane based attachment binding studies in solution divulged that 1 and 2 are avid DNA
5
chemistry. These assemblies are utilized for DNA binding and binders as suggested by the high intrinsic binding constant, K
b
À1
=
5
À1
6
subsequent photo-induced cleavage, with the process monitored B10 M and the apparent binding constant, Kapp = B10 M
optically via UV/Vis spectroscopy and gel electrophoresis. for both 1 and 2 (see Fig. S1–S6 in the ESI‡ for details). This binding
The supported complexes 1 and 2 are shown in Fig. 1. They affinity is comparable with those reported for other Cu(II)–polypyridyl
have a conduit imidazole spine and are terminated with 9-anthryl complexes. Gel electrophoresis studies implied formation of
and 1-pyrenyl aromatic groups, respectively, which are basically more than 90% nicked circles from supercoiled DNA for as little
as 10 mM of the complexes upon UV-A light exposure for B0.5 h
1
,6
(
365 nm, 12 W). This indicates the involvement of metal-to-
a
b
c
Department of Chemistry, University of Delhi, Delhi-110 007, India
Weizmann Institute of Science, Rehovot-76 100, Israel
Applied Physical Chemistry, University of Heidelberg, D-69120 Heidelberg,
Germany. E-mail: Michael.Zharnikov@urz.uni-heidelberg.de
Faculty of Life Sciences and Biotechnology, South Asian University,
Delhi-110 021, India
ligand charge transfer (MLCT) bands (350 nm to 400 nm for
both 1 and 2), associated with the Cu(II)–polypyridyl core, in the
photoexcitation process. Photonuclease activity of 1 and 2
reduced drastically under an N atmosphere, which suggested
2
an important role of the reactive oxygen species (see Fig. S7 in the
d
†
‡
Dedicated to Late Dr Tarkeshwar Gupta.
ESI‡ for details). Mechanistic analysis corroborated participation of
Electronic supplementary information (ESI) available: Materials and methods,
1
singlet oxygen ( O
inhibit the DNA cleavage activity, while the presence of D O
2
increased the amount of cleaved DNA. Note that involvement of
2
) in the cleavage reactions as NaN
3
was found to
experimental details, UV-Vis data, DNA binding activity of 1 and 2, the gel
electrophoresis diagram for DNA cleavage in solution, XPS and NEXAFS data,
and the FT-IR spectrum of ct-DNA in the buffer solution. See DOI: 10.1039/
c4cc05063k
7
8
ꢀ
2À
hydroxyl radicals (OH ) or peroxide anion (O
2
) was neglected
1
1484 | Chem. Commun., 2014, 50, 11484--11487
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as DMSO–KI could not quench DNA cleavage (see Fig. S8 in the covalent attachment and efficient assembly of 1 and 2 on the
9
ESI‡ for the probable mechanism).
substrates.
Unlike DNA binding in solution, where different possibilities of
Robust siloxane-based monolayers of 1 and 2 (abbreviated as
M1 and M2, respectively; see Fig. 1) were fabricated on glass/Si aggregation of the complexes with DNA exist, only the terminal
10
supports following the established procedures. In brief, at first, a aromatic groups of 1 and 2 are exposed to DNA for the supported
silane (3-iodo-n-propyl-trimethoxysilane) coupling layer (CL) was complexes. To prove the activity of these functional groups towards
formed on the freshly cleaned substrate. The CL covered substrates DNA, M1 and M2 were immersed in 50 mM tris-HCl–NaCl buffer
were then immersed in a dry acetonitrile–toluene (3 : 7, v/v) solution solution of ct-DNA (pH = 7.2) for 4 h at room temperature inside an
of the complex 1 or 2 (0.5 mM) under an N
2
atmosphere and kept
N
2
filled glovebox (O
2
o 0.5 ppm). The monolayers were then
at 85 1C for 52 h. Subsequently, the functionalized substrates washed several times with fresh buffer solution to remove free/
were rinsed thoroughly and sonicated gently in acetonitrile and physisorbed ct-DNA and subsequently dried at room temperature
isopropanol to remove any physisorbed material. UV/Vis measure- under an N2 stream. UV-Vis spectra of M1 and M2 showed a
ments on M1 and M2 showed that the complexes adhered strongly bathochromic shift of B6 and 12 nm, respectively, upon DNA
to the substrate surface as they could be removed neither by the binding, along with a hypochromism of B13 and B28% with
‘
‘scotch tape decohesion’’ test nor by a stream of critical carbon respect to their MLCT bands (see Fig. 3). This suggests the inter-
11
dioxide (snow-jet). The assembly of 1 and 2 on the substrate was calation as the preferred binding mode of DNA to supported 1 and 2,
also evidenced by X-ray photoelectron spectroscopy (XPS) and near- similar to the analogous interaction in solution (see Fig. S4, ESI‡). As
edge X-ray absorption fine structure (NEXAFS) spectroscopy. The a negative control, M1 and M2 were kept in DNA-free tris-HCl–NaCl
respective spectra exhibited emissions and absorption resonances buffer solution. After prolonged storage, no changes in the UV-Vis
characteristic of the building blocks and functional units of 1 and 2 spectra were observed.
(see Fig. S9 and S10 in the ESI‡ for details).
The intercalation process was additionally monitored by FT-IR
UV/Vis spectra of the monolayers showed deviations from spectroscopy which has already been successfully used for detection
15
the respective spectra of 1 and 2 in CH
3
CN. For M1 and M2, of DNA immobilized on surfaces like diamond. At first, the FT-IR
MLCT bands appeared at lmax = 372 and 379 nm, respectively, spectrum of ct-DNA in tris-HCl–NaCl buffer solution was recorded
1
6
exhibiting a red shift of B5 and B12 nm with respect to the after preliminary baseline adjustments (see Fig. S11, ESI‡). The
spectra of 1 and 2 in CH CN (see Fig. 3 and ESI‡). This could be characteristic bands of the DNA moieties were observed at
a consequence of quaternization of pendent pyridyl N-atoms
and/or related to cofacial orientation of the molecules within and C–O backbone stretching vibration of the ribose–phosphate
3
1
0c
À1
À
B1100 cm , assigned to the symmetric PO2 stretching mode
1
2a
À1
À
the purview of exciton theory. Other factors can be involved as bond, and at B1210 cm , assigned to asymmetric PO stretching.
2
12b,c
À1
well.
For both M1 and M2, the average surface density was Bands in the range of 1300–1800 cm are ascribed to the ring
2
estimated at B2 molecules per 100 Å of the substrate area, which is stretching modes in the base residues, viz. guanine (G), cytosine (C),
1
3
reasonable for such species. Fourier-transform infrared (FT-IR) adenine (A) and thymine (T); they are comprised of CQO, CQC, and
À1
spectra of M1 and M2 (Fig. 2) revealed the characteristic N–H out- CQN stretching vibrations in the base planes (1746 cm for G,
À1
À1
À1
13
of-plane stretch at B820 cm , the out-of-plane aromatic C–H 1691 cm for T, 1655 cm for A).
À1
À1
stretch at B900 cm , the Sisurface–O–Si stretch at B1110 cm
aromatic C–H wagging at B1450 cm , as well as Si –CH -stretches assemblies were measured and compared to the spectra of M1
,
In the second step, FT-IR spectra of the monolayer–DNA
CL
À1
CL
2
À1 14
at B1300 and 970 cm
. Along with the UV/Vis, XPS, and NEXAFS and M2. The latter spectra served as a baseline, so that only the
data, characteristic stretches in the FT-IR spectra are indicative of the stretches corresponding to DNA could be expected in the spectra
Fig. 3 Representative UV/Vis spectra of M2 (green curve) and M2 + DNA
Fig. 2 FT-IR spectra of M1 (red curve) and M2 (green curve) assembled
on silicon supports. A suitable baseline was recorded using a bare, freshly
cleaned silicon substrate. Vertical dotted lines indicate the positions of the
characteristic IR stretches. The respective assignments are given.
(magenta curve) on glass supports. Suitable baselines were recorded using
freshly cleaned bare glass substrates (thin black curves). Black vertical lines
indicate absorption maxima while double headed arrows emphasize the
extent of bathochromic/hypochromic shifts upon DNA binding.
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Fig. 5 UV/Vis spectra of tris-HCl–NaCl buffer solution recorded after irradiating
either the M1 + DNA (a; olive curve) or the M2 + DNA (b; magenta curve)
assembly immersed in buffer for 0.5 h. The horizontal black line represents
UV/Vis spectra of the buffer solution when the samples were kept in the dark.
The vertical dotted line indicates the peak position of the spectra.
Fig. 4 FT-IR spectra of M1 + DNA (olive curve) and M2 + DNA (magenta
curve) assemblies on the silicon substrate recorded after preliminary baseline
adjustments. The spectra exhibit IR stretches associated with the immobilized
DNA only. Significant variations are observed as compared to the spectra of
ct-DNA in solution. Vertical dotted lines indicate the positions of characteristic
IR stretches. The respective assignments are given.
were cleaved into shorter fragments such as oligonucleotides
under UV-A light irradiation, with subsequent diffusion into the
of the monolayer–DNA assemblies (Fig. 4). In these spectra, shifts bulk phase. Note that the amount of DNA fragmented is higher
À1
À1
À1
of B10–12 cm for the 1746 cm peak, B4–10 cm for the for M2 as compared to M1, because the absorbance at lmax
1
=
À1
À1
À1
691 cm peak, and B0–2 cm for the 1602 cm peak were B260 nm in the M2 case is almost twice as large as that for M1
observed in the ring stretching modes of the base residues as (see Fig. 5). This can be tentatively explained by the superior
compared to the spectrum of ct-DNA in solution, which is indicative p-conjugation, aromaticity, and planarity of the pyrenyl moiety of
of monolayer intercalation mainly into the G–C base pairs. In M2, which eventually makes this system a better monolayer-based
À
À1
addition, the shift of the asymmetric as well as symmetric PO
modes (B1222 and B1064 cm , respectively) by B2–4 cm
2
photonuclease. In this context, the most plausible role of the
assembled 1 and 2 appears to be that of a photosensitizer, where
À1
indicates electrostatic interactions between the complexes on the the photosensitized Cu(II)–polypyridyl cores of 1 and 2 transfer
Si substrate and the phosphate backbone of the ct-DNA. Thus, FT-IR energy to activate oxygen available in the buffer solution from
3
3
À
g
) to a highly reactive singlet state
spectroscopy data provide clear and plausible evidence for the DNA its stable triplet state ( O
attachment to the monolayers, predominantly via the intercalation ( O , D
2 g
binding mode.
2
, S
) in the near vicinity of DNA leading to its rupture.
In addition to the UV/Vis studies, agarose gel electrophoresis
1
1
1a,17
In further experiments, a set of 8 identical glass slides experiments were conducted using the same buffer extraction
(
3.0 cm  1.0 cm  1.25 mm), grafted with either M1 or M2, procedure, following the exposure of the monolayer–DNA
was dipped in ct-DNA solution in 50 mM tris-HCl–NaCl buffer assemblies to UV-A light (see Fig. 6). 20 mL solution of the
(
pH = 7.2) for B4 h using a Teflon slide holder. Afterwards, the buffer containing DNA fragments and 2 mL of the loading buffer
slides were thoroughly washed with buffer solution to remove containing 25% bromophenol blue, 0.25% xylene cyanol, and
free/physisorbed ct-DNA and subsequently dried under an N 30% glycerol, were used for running the 0.8% agarose gel in
2
stream. The slides were then kept in fresh buffer solution and 1Â tris-acetate–EDTA buffer at a potential of 20 V for 15 minutes.
irradiated with UV-A light (365 nm, 12 W) for either 0.5 or 1 h. As seen in the gel pictures, DNA fragmentation has indeed
Finally, the container was shaken thoroughly and 2 mL of occurred as supercoiled DNA (SC) has moved only slightly while
the solution were extracted and tested by UV-Vis spectroscopy the fragmented DNA has moved much farther. It is also evident
(200–400 nm range) in transmission mode. As a negative control, that M2 cleaves DNA with higher efficiency as the intensity of
similar experiments were performed under dark conditions and DNA fragments for M2 is higher than that for M1. These findings
the UV/Vis spectra of the extracted solution were recorded agree with the results of the UV/Vis measurements (vide supra).
thereafter. In the case of exposure of the monolayer–DNA Moreover, as can be expected, an increase of the irradiation time
assembly to UV-A light, UV-Vis spectra of the solution showed to 1 h resulted in an increase of the intensity of the bands
a strong signal with a maximum at B260 nm indicating the associated with the DNA fragments, as shown in Fig. 6.
presence of DNA fragments (see Fig. 5). In contrast, analogous
We demonstrated that the Cu(II)–polypyridyl complexes 1
UV-Vis spectra of the samples kept in the dark did not show any and 2 immobilized on silicon/glass supports in a monolayer
band characteristic of DNA (see Fig. 5). Additional control fashion (M1 and M2) are able to intercalate into DNA and cleave
experiments, in which M1 and M2 immersed into DNA-free it efficiently upon exposure to UV-A light without the need of any
buffer solution were subjected to UV-A light for 0.5 h, showed additional reducing or oxidizing chemicals. No photo-degradation of
no effect of the irradiation on the monolayers themselves, with the monolayer assemblies was observed. The DNA cleavage activity
no evidence for their damage or fragmentation. The above was monitored by UV/Vis spectroscopy and gel-electrophoresis
results suggest that the DNA chains bound to the monolayers experiments. The terminal aromatic moieties in M1 and M2
1
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2
3
(a) E. Ostuni, L. Yan and G. M. Whitesides, Colloids Surf., B, 1999, 15, 3;
(b) S. Zhang, Nat. Biotechnol., 2003, 21, 1171; (c) V. Singh, M. Zharnikov,
A. Gulino and T. Gupta, J. Mater. Chem., 2011, 21, 10602.
(a) N. Higashi, T. Inoue and M. Niwa, Chem. Commun., 1997, 1507;
(
b) Y. Zhang, S. Yang, C. Liu, X. Dai, W. Cao, J. Xu and Y. Li, New
J. Chem., 2002, 26, 617–620.
S. Sortino, S. Petralia, G. G. Condorelli, S. Conoci and G. Condorelli,
Langmuir, 2003, 19, 536.
4
5
6
C. Haensch, S. Hoeppener and U. S. Schubert, Chem. Soc. Rev., 2010,
3
9, 2323.
Fig. 6 Gel electrophoresis diagram of photonuclease activity of M1 and
M2. NC and SC refer to nicked circular (form II) and supercoiled (form I)
forms of ct-DNA, respectively. ‘A’ and ‘B’ correspond to a UV-A irradiation
time of 0.5 and 1 h, respectively.
(a) V. Uma, M. Kanthimathi, T. Weyhermuller and B. U. Nair,
J. Inorg. Biochem., 2005, 99, 2299; (b) M. Scarpellini, A. Neves,
R. Horner, A. J. Bortoluzzi, B. Szpoganics, C. Zucco, R. A. N. Silva,
V. Drago, A. S. Mangrich, W. A. Ortiz, W. A. C. Passos, M. C. B. de
Oliveira and H. Terenzi, Inorg. Chem., 2003, 42, 8353; (c) Y. Jin and
J. A. Cowan, J. Am. Chem. Soc., 2005, 127, 8408.
7
8
B. W. Henderson and T. Dougherty, J. Photochem. Photobiol., 1992,
are assumed to play a key role in DNA binding and cleavage
activities, with the pyrenyl group (M2) performing better than
55, 145.
J. R. Harbour and S. L. Issler, J. Am. Chem. Soc., 1982, 104, 903.
9-anthryl (M1). The fabricated monolayers as well as other analogous
9 (a) P. B. Merkel, R. Nilsson and D. R. Kearns, J. Am. Chem. Soc., 1972,
4, 1030; (b) J. E. Repine, J. W. Eaton, M. W. Anders, J. R. Hoidal and
9
systems represent efficient model systems for controlled DNA
cleavage. In addition, they can be potentially utilized for fibre-
optics based DNA cleavage in vivo.
This work was financially supported by the Department of
Science and Technology-Nano Mission (SR/NM/NS-12/2010),
New Delhi. VS and AK thanks the University Grants Commission
for a senior research fellowship. PCM thanks the Council of
Scientific and Industrial Research for a senior research fellowship.
RDG thanks the Department of Science and Technology (SERB/F/
R. B. Fox, J. Clin. Invest., 1979, 64, 1642.
1
0 (a) T. Gupta, P. C. Mondal, A. Kumar, Y. L. Jeyachandran and
M. Zharnikov, Adv. Funct. Mater., 2013, 23, 4227; (b) A. Kumar,
M. Chhatwal, P. C. Mondal, V. Singh, A. K. Singh, D. A. Cristaldi,
R. D. Gupta and A. Gulino, Chem. Commun., 2014, 50, 3783;
(
c) V. Singh, P. C. Mondal, M. Chhtawal, Y. L. Jeyachandran and
M. Zharnikov, RSC Adv., 2014, 4, 23168.
1
1
1 B. Y. Chow, D. W. Mosley and J. M. Jacobson, Langmuir, 2005, 21, 4782.
2 (a) K. Oberg, U. Edlund, B. Eliasson, A. Shchukarev, K. Sheshadri
and D. Allara, J. Phys. Chem., 2000, 104, 10627; (b) X. Yan and
D. Holten, J. Phys. Chem., 1988, 92, 409; (c) B. A. Gregg, M. A. Fox and
A. J. Bard, J. Phys. Chem., 1989, 93, 4227.
1
424/2013–14) for financial assistance. MZ and YLJ thank Ch. W ¨o ll
13 V. Singh, P. C. Mondal, Y. L. Jeyachandran, M. Zharnikov and
and A. Nefedov for the technical cooperation at BESSY II and the
BESSY II staff for the assistance.
T. Gupta, Analyst, 2012, 137, 3216.
14 M. Li, Modification of Silicon by Self-Assembled Monolayers for Application
in Nano-Electronics and Biology, PhD thesis, Rutgers University, 2007.
1
5 (a) K. Ushizawa, Y. Sato, T. Mitsumori, T. Machinami, Y. Ueda and
T. Ando, Chem. Phys. Lett., 2002, 351, 105; (b) V. Vamvakaki and
N. A. Chaniotakis, Electroanalysis, 2008, 20, 1845.
Notes and references
1
(a) S. Dhar, D. Senapati, P. K. Das, P. Chattopadhyay, M. Nethaji and 16 (a) E. Taillandier and J. Liquier, Methods Enzymol., 1992, 211, 307;
A. R. Chakravarty, J. Am. Chem. Soc., 2003, 125, 12118; (b) S. Dhar and
A. R. Chakravarty, Inorg. Chem., 2005, 44, 2582; (c) U. Schatzschneider,
(b) D. K. Jangir, G. Tyagi, R. Mehrotra and S. Kundu, J. Mol. Struct.,
2010, 969, 126.
Eur. J. Inorg. Chem., 2010, 1451; (d) K. Suntharalingam, D. J. Hunt, 17 (a) M. B. Fleisher, K. C. Waterman, N. J. Turro and J. K. Barton, Inorg.
A. A. Duarte, A. J. P. White, D. J. Mann and R. Vilar, Chem. – Eur. J.,
012, 18, 15133; (e) B. Zheng, C. Wang, X. Xin, F. Liu, X. Zhou,
J. Zhang and S. Guo, J. Phys. Chem. C, 2014, 14, 7637.
Chem., 1986, 25, 3549; (b) J. Piette, J. Photochem. Photobiol., B, 1991,
11, 241; (c) T. Torring, S. Helmig, P. R. Ogilby and K. V. Gothelf, Acc.
Chem. Res., 2014, 47, 1799.
2
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