Correlation Between Cellular Localization and Binding Preference to

Feb 2, 2011 - to RNA, DNA, and Phospholipid Membrane for Luminescent. Ruthenium(II) Complexes. Maria Matson, Frida R. Svensson,* Bengt Nordén, and ...
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Correlation Between Cellular Localization and Binding Preference to RNA, DNA, and Phospholipid Membrane for Luminescent Ruthenium(II) Complexes Maria Matson, Frida R. Svensson,* Bengt Norden, and Per Lincoln

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Department of Chemical and Biological Engineering, Chalmers University of Technology, Kemiv€agen 10, SE-41296, Gothenburg, Sweden ABSTRACT: Because of their unique photophysical properties, sensitively depending on environment, ruthenium dipyridophenazine (dppz) complexes are interesting as probes for cellular imaging with fluorescence microscopy. Here three complexes derivatized with alkyl ether chains of varied length, which exhibit distinctly different cellular staining patterns by confocal laser scanning microscopy, are studied regarding their binding preference for rRNA compared with calf thymus DNA (ct-DNA) and phospholipid membranes. Co-staining with commercial RNA and membrane-specific dyes shows that whereas the least lipophilic complex exclusively stains DNA inside the nucleus, the most lipophilic complex preferentially stains membrane-rich parts of the cell. Interestingly, only the intermediate lipophilic complex shows intense staining of the RNA-rich nucleoli. The intracellular localizations of the probes correlate with their binding preferences concluded from spectroscopy measurements.

’ INTRODUCTION Optical probes for cellular imaging are important in the research aiming for understanding structure and function of biological systems. When new probes are designed and synthesized, detailed knowledge of their cellular distribution and binding preference for intracellular molecules is essential for how they may be applied. Ruthenium dipyridophenazine (dppz) complexes are known to show environment-dependent emission, a “light-switch effect” making them highly luminescent in hydrophobic environments, such as inside stacks of DNA bases or lipid membrane bilayers, whereas in aqueous solution, the emission is strongly quenched.1 Their low background emission and other characteristic photophysical properties, including longlived excited metal-to-ligand charge-transfer state (MLCT) and the red-shifted emission, make them interesting candidates as molecular probes for cellular imaging using fluorescence microscopy. Recently, polypyridyl complexes of ruthenium(II), rhenium(I), and iridium(III) have been studied for such applications,2-12 and their cellular localization and uptake have been explored.13-23 The DNA binding properties of monoand binuclear ruthenium complexes have been extensively investigated;24-32 however, only a few studies have been reported on RNA binding properties. O’Connor et al. studied a phenanthridine-ruthenium(II) complex, which is promising as an RNA probe because of its altered lifetime and fluorescence intensity upon binding to RNA.33 Fluorescence microscopy showed that the complex indeed stains RNA-rich areas, such as the nucleoli and regions outside the nucleus, but no studies of its DNA or membrane-binding properties were reported. The nucleic r 2011 American Chemical Society

acid structure has been concluded to have a significant effect on the binding of ruthenium complexes. Xu et al. reported a ruthenium polypyridyl complex that binds more strongly to tRNA than to ct-DNA,34 and Spillane et al. discovered that their binuclear complexes bind only weakly to duplex RNA, although they were able to detect the presence of bulges (stretches of unpaired RNA bases) by a 50-fold increase in binding affinity.35 Other studies have focused on the enantioselectivity of ruthenium complexes with respect to RNA, and significant variations in binding affinity depending on enantiomeric form have been concluded.36-39 In addition to their nucleic acid binding, ruthenium dppz complexes have also been investigated as membrane probes. We have previously shown that nitrile- and amide-substituted ruthenium dppz complexes tend to align the dppz ligand parallel with the membrane surface, whereas for the unsubstituted Ru(phen)2dppz2þ complex, the dppz moiety is oriented parallel to the lipid hydrocarbon chains.40 This is also the preferred orientation for dppz derivates substituted with alkyl ether chains that insert themselves deep into the membrane.41 We recently reported that the binding preference for DNA and negatively charged phospholipid membranes can be tuned by small changes in lipophilicity of alkyl-ether-substituted ruthenium dppz complexes. These complexes were also shown to traverse the cell membrane as a result of photoactivated uptake Received: October 4, 2010 Revised: December 10, 2010 Published: February 2, 2011 1706

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The Journal of Physical Chemistry B Scheme 1. Molecular Structure of the Three Ruthenium(II) Dppz Complexes, Denoted D2, D4, and D6 Depending on the Length of the Alkyl Ether Chains

and accumulate in internal structures upon illumination.42 Here the cellular localization of these complexes compared with RNAand membrane-selective dyes is studied in methanol-fixed cells by confocal laser scanning microscopy (CLSM). Their binding preferences to rRNA, calf thymus DNA (ct-DNA), and negatively charged, as well as neutral zwitterionic, phospholipid membranes are explored by steady-state emission spectroscopy. Furthermore, the DNA-binding geometries of the complexes are studied with linear dichroism spectroscopy on flow-oriented DNA. The cellular biomacromolecular binding of the complexes is compared with their preferential affinity for the pure microenvironments in vitro.

’ EXPERIMENTAL METHODS Materials. Ribosomal RNA from bovine liver and calf thymus DNA was purchased from Sigma. DOPC (1,2-dioleoyl-sn-glycero-3-phosphocholine) and DOPG (1,2-dioleoyl-sn-glycero3-phosphoglycerol) were from Larodan (Malm€o, Sweden). The buffer used for emission spectroscopy and linear dichroism was 150 mM NaCl, 10 mM HEPES, and 1 mM EDTA dissolved in mQ-H2O, pH 7.4. Chinese ::hamster ovarian (CHO-K1) cells were a kind gift from Prof. Ulo Langel, Stockholm University. Cell culture reagents, calf bovine serum, HAM’s F12 medium, trypsin, and L-glutamine were purchased from GTF. The commercial RNA probe “SytoRNA select” (structure not disclosed by manufacturer) and the membrane probe N-(3-triethylammoniumpropyl)-4-(6-(4-(diethylamino)phenyl)hexatrienyl) pyridinium dibromide (FM 4-64) were from Molecular Probes. Synthesis. The complexes used in this study are shown in Scheme 1, where the dppz ligand of the complex Ru(phen)2dppz2þ has been substituted with alkyl ether chains of varying lengths. The complexes were synthesized as described elsewhere.41,42 Cell Culturing. CHO-K1 cells were grown in HAM’s F12 medium with 10% calf bovine serum and L-glutamine (2 mM) at 37 °C and 5% CO2. The cells were seeded on round coverslips at a density of ∼80 000 cells/coverslip and cultured for 2 days. Prior to imaging, cells were fixed by incubation in cold methanol (-20 °C) for 10 min, rinsed once with serum-free medium, and mounted in a solution chamber. Ruthenium complexes were dissolved in DMSO and diluted in serum-free medium to the final concentration of 10 μM, “SytoRNA select” to 1 μM, and FM 4-64 to 6.6 μM before the solutions were added to the coverslips. Confocal Laser Scanning Microscopy. For confocal fluorescence imaging, Leica TCS SP2 RS (Wetzlar, Germany) with a PL APO 63/1.32 objective was used. An Ar laser (488 nm) was used for excitation of ruthenium complexes, RNA probe, and FM 4-64, and the emission was acquired at 650-700 nm for the

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ruthenium complexes and FM 4-64 and at 520-540 nm for the RNA probe. Co-staining of ruthenium complexes and the RNA probe as well as with the RNA probe and FM 4-64, was performed. The PMT and gain were optimized for each image. The intensity in a selected line was plotted for each dye to compare the intensity profiles through the cell. Preparation of Large Unilamellar Lipid Vesicles (LUVs). DOPC and DOPG dissolved in chloroform were mixed at a ratio of 4:1 to yield 20% negatively charged LUVs, and for zwitterionic LUVs, pure DOPC was used. The lipids were dried by rotary evaporation and then kept under vacuum for at least 2 h. The lipid film was then dispersed in buffer while vortexing, whereupon five freeze-thaw cycles (liquid nitrogen/37 °C) were performed. The vesicles were extruded 21 times through two polycarbonate filters with pore size 100 nm using a handheld syringe LiposoFast-Pneumatic extruder (Avestin, Canada) to obtain LUVs of uniform size. Emission Spectroscopy. Steady-state emission measurements were performed on a Cary Eclipse fluorescence spectrophotometer (Varian) at room temperature. Emission of ruthenium complex hexafluorophosphates (2 μM) and the RNA probe “SytoRNA select” (4 μM) were measured in DOPC/DOPG-LUVs (200 μM), ct-DNA (40 μM bases), and rRNA (40 μM bases). The maximal emission wavelengths of the complexes were measured in zwitterionic DOPC-LUVs (200 μM) upon the addition of ct-DNA or rRNA. For binding preference measurements, samples with complex bound to rRNA or DNA were gradually mixed with each other until the RNA and DNA concentrations were the same in both samples, and for each mixing step, an emission spectrum was recorded. Ruthenium complexes and the RNA probe were excited at 440 and 490 nm, and emission was recorded between 550 and 800 nm and 500 and 700 nm, respectively. Flow Linear Dichroism Spectroscopy. Linear dichroism (LD) is defined as the difference in absorption of linearly polarized light parallel and perpendicular to an orientation axis. To obtain an LD signal, oriented molecules are required, which here are achieved by a shear flow in a Couette cell. The sample is applied in the narrow gap between two cylinders, where the outer one is rotating at 1000 rpm. The transition moments of the bases in flow-oriented DNA are perpendicular to the orientation axis and will hence absorb light, resulting in a negative LD peak around 260 nm.43 Molecules that are too small to orient in the flow can be detected by LD when bound to DNA if they absorb in the visible region, and from the sign and magnitude of the signal, the binding orientation can be assessed.29 LD was measured with a Chirascan spectrometer (Applied Photophysics, U.K.) equipped for linear dichroism detection. Spectra were recorded between 200 and 600 nm, and the concentrations used were 100 μM DNA bases and 10 μM complex.

’ RESULTS The cellular localization of the three complexes was studied by CLSM, characteristic cellular staining patterns shown in Figure 1. The least lipophilic complex D2 is found to only stain DNA in the nucleus, however, excluding the nucleoli. By contrast, D4 shows intense staining of cytoplasmic and nucleoli territories and also some weaker staining of the nucleus, whereas the most lipophilic complex, D6, shows weak emission in the nucleus but stains the cytoplasm and to some extent also the nucleoli. To compare the accumulation of the ruthenium complexes 1707

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Figure 1. CLSM images of methanol-fixed cells costained with complex (10 μM, red) or FM 4-64 (6.6 μM, purple) and “SytoRNA select” (1 μM, green). The graphs show intensity profiles through a cell where red curves correspond to the ruthenium complexes, green curves correspond to the RNA probe, and purple curves correspond to the membrane probe. The black horizontal bars indicate the location of the nucleoli. Because the voltage over the photomultiplier tube of the microscope was optimized for each image, the intensity profiles, and not the absolute intensities, of the complexes and the probes should be compared.

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with the cellular localization of RNA, we performed costaining experiments with the probe “SytoRNA select”. The RNA probe stains the cytoplasm and the nucleoli, areas known to be rich in RNA.44 The CLSM images and the intensity profiles through the cells clearly show that the two most lipophilic complexes have staining patterns closely resembling those of the commercial RNA probe. The differences are mainly that D4, in contrast with the RNA probe, also shows some emission in the nucleus and that the nucleoli are not as intensely stained by D6. In contrast with D4 and D6, the least lipophilic complex D2 shows a staining pattern completely opposite to that of the RNA probe. This is clearly shown in the intensity profile plot of D2, where the emission is much less intense in the nucleoli compared with the rest of the nucleus. The cytoplasm is also rich in membrane structures; therefore, the staining pattern for the RNA probe was compared with that of the membrane probe FM 4-64. As seen in Figure 1, the staining by the two probes is similar, but in comparison with the RNA probe, FM 4-64 more distinctly stains some parts outside the nucleus, whereas the nucleoli staining is less intense. Because the luminescence intensity and emission wavelength of the ruthenium dppz chromophore depend sensitively on the polarity and water accessibility of its microenvironment, the biomolecular binding preferences of the complexes are readily studied with emission spectroscopy. Figure 2 shows emission spectra of D4 and the RNA probe in the presence of DNA, rRNA, and LUVs. For the ruthenium complex, we observe strong emission in DNA, somewhat smaller and red-shifted emission in phospholipid membrane, and rather weak emission in rRNA. The RNA probe shows about four times more intense emission in rRNA compared with DNA, whereas the emission in LUVs is very low. The finding that D4 stains the cytoplasm and rRNA-rich nucleoli more effectively than the nucleus, even though the emission in pure systems shows that DNA-bound complex has

Figure 2. Emission spectra of D4 (2 μM) in rRNA (40 μM bases, red), DNA (40 μM bases, black), DOPC/DOPG-LUVs (200 μM, green), and pure buffer (blue) and the corresponding emission spectra of the RNA probe “SytoRNA select” (4 μM). 1708

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The Journal of Physical Chemistry B the highest emission intensity, suggests that the binding preference of the complex has an important role for the emission intensity pattern in the cellular environment. Competition experiments of all three complexes in mixtures of zwitterionic LUVs and RNA or DNA were therefore studied, and the maximum emission wavelengths for D4 and D6 are displayed in Figure 3. For D4, a clear wavelength shift is observed when rRNA is added to LUV-bound complex, and the maximum emission wavelength in this mixture is between that for D4 in pure rRNA and LUV. In a previous study, with negatively charged LUVs, we showed that both D4 and D6 exclusively prefer membrane binding over binding to DNA,42 and the same membrane-binding preference is observed in a mixture of RNA and negatively charged membranes (not shown). With the zwitterionic membranes studied here, the most lipophilic complex, D6, is found to

Figure 3. Emission wavelength maximum of D4 (black) and D6 (red) in DNA, LUV, and a mixture thereof (solid lines) and in rRNA, LUV, and a mixture thereof (dashed lines).

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still prefer membrane binding because the emission obviously does not shift in wavelength upon the addition of either rRNA or DNA. By contrast, the least lipophilic complex, D2, shows no significant emission in LUVs and only weak emission when bound to rRNA, whereas the emission when bound to DNA is strong (six-fold increase compared with rRNA). Upon addition of DNA or rRNA to a LUV solution containing D2, the emission increases to the value in pure nucleic acid (not included in Figure 3). To get further insight into the binding preferences of the complexes, rRNA and ct-DNA binding were compared. In Figure 4, emission spectra of complexes bound to ct-DNA (thick black lines) and rRNA (thick red lines) for the ratio nucleic acid phosphate (P) to ruthenium complex of P/Ru 10:1 are shown. Samples with DNA and RNA solutions were gradually mixed, and emission spectra were recorded for each mixing step (thin lines). For all complexes, a decrease in intensity is observed with increasing RNA concentration, and D2 and D4 show stronger emission upon the addition of DNA. Although the most lipophilic complex, D6, has low emission intensity when bound to RNA, no spectral change is observed upon addition of DNA, demonstrating selectivity for RNA over DNA. The binding preference for D2 and D4 is not as apparent as that for D6 because the emission intensity is decreasing with increasing RNA concentration but increasing with higher DNA concentration. These complexes thus bind to both nucleic acids to some extent. The relative affinity for RNA compared with that for DNA, however, appears to be higher for D4 because the emission intensity, for the same concentration of nucleic acid, is more similar to the intensity in pure RNA compared with the corresponding spectra with D2. Information regarding the DNA binding mode of the complexes can be obtained by flow linear dichroism spectroscopy. The ct-DNA (but not the rRNA) molecules are long enough to orient in the shear flow of the Couette cell with the helix axis

Figure 4. Emission spectra of D2, D4, and D6 (2 μM) in pure ct-DNA (40 μM bases, thick black line) and in pure rRNA (40 μM bases, thick red line). When the ct-DNA sample is gradually mixed with the rRNA sample (thin black lines), the intensity of the ct-DNA samples is decreasing, and for D2 and D4, an increase is shown for the opposite procedure (thin red lines), whereas the emission of RNA-bound D6 does not change upon addition of ct-DNA. 1709

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Figure 5. LD spectra of the ruthenium complexes (10 μM) bound to ctDNA (100 μM) and an enlarged inset of the region 320-550 nm. D2 (green), D4 (black), and D6 (red) are compared with the parent complex Ru(phen)2dppz2þ (gray) and pure ct-DNA (blue).

parallel to the flow direction, as shown by the negative LD peak at 260 nm originating from the perpendicularly oriented nucleobases. The ruthenium complexes themselves are too small to align in the shear flow, but when intercalated into DNA, the complexes become oriented and show a characteristic pattern of negative and positive bands between 320 and 550 nm. (See the inset in Figure 5.)27-29 All complexes exhibit the same LD pattern with some variations of the amplitude, which shows that they indeed bind DNA by intercalation, although the amplitude for D6 is quite low.

’ DISCUSSION The CLSM imaging of fixed cells costained with ruthenium complex and commercial RNA probe shows that the localization of the two more lipophilic complexes is very similar to that of the RNA probe. The staining patterns of the complexes differ slightly, however, with D4 having somewhat stronger emission in the nucleus and D6 less in the nucleoli compared with the RNA probe. In contrast with D4 and D6, D2 only stains the nucleus and shows an inverted staining compared with the RNA probe, which is clearly seen by the absence of nucleoli staining in the CLSM images and in the intensity profiles in Figure 1. When the RNA probe is compared with the membrane probe FM 4-64, the staining of the probes is similar, but the nucleoli dyeing is less intense for FM 4-64, indicating that the staining of D4 cannot solely be explained by membrane binding. In cells, DNA is located in the nucleus, whereas membranes are mainly found in the cytoplasm and in the cell membrane. There are many different types of RNA, most of which are located in the cytoplasm, but in the nucleoli where the ribosomal subunits are formed, a particular high concentration of rRNA is found.45 Size and number of nucleoli vary during the cell cycle; therefore, the subunits are not always seen in the cells. Emission spectroscopy shows that the RNA probe has approximately four times more intensive emission when bound to rRNA compared with ctDNA and, importantly, no emission is exhibited in the LUV solution. Hence, we conclude that “SytoRNA select” exclusively stains RNA inside the cell. The corresponding emission spectra for D4 show that the complex has the highest intensity when bound to DNA and much lower when bound to rRNA, but interestingly, the CLSM images show that the cytoplasm and the rRNA-rich nucleoli are clearly stained and that the lowest

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intensity is found in the DNA-rich nucleus. These findings support the concluded significance for RNA binding for this complex in the cellular milieu. In the search for an explanation of why the three complexes tend to accumulate in different parts of the cell, the binding preference was investigated, comparing the affinity with rRNA, ct-DNA, and lipid membranes. The complexes have previously been shown to have different preferences to DNA and membranes with 20% negatively charged lipids, where the least lipophilic complex D2 favors DNA binding, whereas D4 and D6 both bind more strongly to membranes.42 Here we show that with zwitterionic LUVs the least lipophilic complex, D2, still prefers binding to DNA and D6 favors membrane binding but that D4 switches preference and favors binding to DNA instead of membrane. Regarding the RNA binding, in the presence of negatively charged LUVs, all complexes prefer membrane binding over rRNA, but with zwitterionic LUVs, the preference is changed for D2, which favors rRNA, and for D4, which seems to bind to both rRNA and LUVs. The lipophilic character of D6 results in preferred membrane binding, even to zwitterionic LUVs, showing the importance of small changes in hydrophobicity for the binding preferences of the complexes. When the preference regarding nucleic acid binding is compared, the results clearly show that D6 favors RNA binding because the emission spectral properties of the RNA-bound complex do not change upon the addition of DNA. (See Figure 3.) The other two complexes show a distribution of both RNA and DNA binding, where D4 seems to have higher relative affinity for RNA than D2. Hence, structural differences of the nucleic acids appear to be essential for the binding properties of the complexes.34 RNA has many different secondary structures such as A-form helices, internal loops, junctions, and hairpin loops, which are not commonly found for DNA.46 These structures are important for the function of the RNA and might provide more lipophilic binding sites than DNA and hence be a reason why the more lipophilic complexes have higher affinity for rRNA compared with DNA. Low luminescence for the RNAbound complex suggests that the RNA binding mode is less shielded from hydrogen bonding of the aza nitrogens on the dppz ligand with water, resulting in quenching of the emission, compared with when the complex is bound to DNA or embedded in the lipid membrane. LD results confirm that all complexes bind to DNA by intercalation, although the most lipophilic complex D6 shows weak LD signal from the dppz transition. The decreased LD magnitude suggests that a smaller fraction of this complex is intercalating into DNA and that a significant number of complexes bind in a disordered fashion. The DNA signal at 260 nm is also decreased, which suggests that D6 causes some distortion (compaction and/or aggregation) and thereby impairs the orientation of the DNA helix. D6 has a tendency to aggregate at high salt concentration, as evidenced from the fact that its luminescence in high salt buffer is not completely quenched. The unspecific DNA binding mode of D6 can explain why the emission is lower compared with DNAbound D2 and D4 in Figure 4.

’ CONCLUSIONS In conclusion, we demonstrate that three lipophilic ruthenium dppz complexes with tunable lipophilicity show major variations in binding preference comparing RNA with DNA and phospholipid membrane. The binding preferences studied by emission 1710

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The Journal of Physical Chemistry B spectroscopy correlate with confocal microscopy analysis of their cellular staining patterns, and new insight into the macromolecular targeting is provided by costaining with RNA- and membrane-specific dyes. The least lipophilic complex, D2, binds to DNA in the cell nucleus, and no nucleoli staining is observed, which is consistent with the low emission and low affinity for pure rRNA. D6 binds selectively to membrane structures and stains the cytoplasm, but although this lipophilic complex also shows higher emission and affinity for pure RNA, the nucleoli are only weakly stained. D4, in contrast, is the complex that most strongly stains the nucleoli with a cellular staining pattern very similar to the “SytoRNA select” probe, and despite its high emission when intercalated into DNA, the emission from the nucleus is weak. This staining pattern can be explained only by higher affinity for RNA and lipid membranes than for the nuclear DNA. The finding that the complexes show such different preferences as a response to such small structural variations highlights the potential of transition-metal complexes as selective and sensitive cellular and biomacromolecular probes.

’ AUTHOR INFORMATION Corresponding Author

*Tel: þ46317723069. Fax: þ46317723858. E-mail: frida.svensson@ chalmers.se. † E-mail: [email protected].

’ ACKNOWLEDGMENT This work was funded by grants from the Swedish Research Council (VR) and the European Commission. ’ REFERENCES (1) Friedman, A. E.; Chambron, J. C.; Sauvage, J. P.; Turro, N. J.; Barton, J. K. J. Am. Chem. Soc. 1990, 112, 4960–4962. (2) Fernandez-Moreira, V.; Thorp-Greenwood, F. L.; Coogan, M. P. Chem. Commun. (Cambridge, U. K.) 2010, 46, 186–202. (3) Gill, M. R.; Garcia-Lara, J.; Foster, S. J.; Smythe, C.; Battaglia, G.; Thomas, J. A. Nat. Chem. 2009, 1, 662–667. (4) Rajendiran, V.; Palaniandavar, M.; Periasamy, V. S.; Akbarsha, M. A. J. Inorg. Biochem. 2010, 104, 217–220. (5) Amoroso, A. J.; Coogan, M. P.; Dunne, J. E.; Fernandez-Moreira, V.; Hess, J. B.; Hayes, A. J.; Lloyd, D.; Millet, C.; Pope, S. J.; Williams, C. Chem. Commun. (Cambridge, U. K.) 2007, 3066–3068. (6) Lamoureux, M.; Seksek, O. J. Fluoresc. 2010, 20, 631–643. (7) Lau, J. S.; Lee, P. K.; Tsang, K. H.; Ng, C. H.; Lam, Y. W.; Cheng, S. H.; Lo, K. K. Inorg. Chem. 2009, 48, 708–718. (8) Lo, K. K. W.; Lee, P. K.; Lau, J. S. Y. Organometallics 2008, 27, 2998–3006. (9) Pisani, M. J.; Weber, D. K.; Heimann, K.; Collins, J. G.; Keene, F. R. Metallomics 2010, 2, 393–396. (10) Scharwitz, M. A.; Ott, I.; Geldmacher, Y.; Gust, R.; Sheldrick, W. S. J. Organomet. Chem. 2008, 693, 2299–2309. (11) Scharwitz, M. A.; Ott, I.; Gust, R.; Kromm, A.; Sheldrick, W. S. J. Inorg. Biochem. 2008, 102, 1623–1630. (12) Yu, M.; Zhao, Q.; Shi, L.; Li, F.; Zhou, Z.; Yang, H.; Yi, T.; Huang, C. Chem. Commun. (Camb.) 2008, 2115–2117. (13) Lo, K. K.; Lee, T. K.; Lau, J. S.; Poon, W. L.; Cheng, S. H. Inorg. Chem. 2008, 47, 200–208. (14) Lo, K. K. W.; Louie, M. W.; Zhang, K. Y. Coord. Chem. Rev. 2010, 254, 2603–2622. (15) Neugebauer, U.; Pellegrin, Y.; Devocelle, M.; Forster, R. J.; Signac, W.; Moran, N.; Keyes, T. E. Chem. Commun. (Cambridge, U. K.) 2008, 5307–5309.

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(16) Puckett, C. A.; Barton, J. K. J. Am. Chem. Soc. 2007, 129, 46–47. (17) Puckett, C. A.; Barton, J. K. Biochemistry 2008, 47, 11711– 11716. (18) Puckett, C. A.; Barton, J. K. J. Am. Chem. Soc. 2009, 131, 8738– 8739. (19) Puckett, C. A.; Barton, J. K. Bioorg. Med. Chem. 2010, 18, 3564– 3569. (20) Puckett, C. A.; Ernst, R. J.; Barton, J. K. Dalton Trans. 2010, 39, 1159–1170. (21) Schatzschneider, U.; Niesel, J.; Ott, I.; Gust, R.; Alborzinia, H.; Wolfl, S. ChemMedChem 2008, 3, 1104–1109. (22) Zava, O.; Zakeeruddin, S. M.; Danelon, C.; Vogel, H.; Gratzel, M.; Dyson, P. J. ChemBioChem 2009, 10, 1796–1800. (23) Zhang, K. Y.; Li, S. P.; Zhu, N.; Or, I. W.; Cheung, M. S.; Lam, Y. W.; Lo, K. K. Inorg. Chem. 2010, 49, 2530–2540. (24) Barton, J. K.; Danishefsky, A. T.; Goldberg, J. M. J. Am. Chem. Soc. 1984, 106, 2172–2176. (25) Biver, T.; Secco, F.; Venturini, M. Coord. Chem. Rev. 2008, 252, 1163–1177. (26) Erkkila, K. E.; Odom, D. T.; Barton, J. K. Chem. Rev. 1999, 99, 2777–2795. (27) Hiort, C.; Lincoln, P.; Norden, B. J. Am. Chem. Soc. 1993, 115, 3448–3454. (28) Holmlin, R. E.; Stemp, E. D. A.; Barton, J. K. Inorg. Chem. 1998, 37, 29–34. (29) Lincoln, P.; Broo, A.; Norden, B. J. Am. Chem. Soc. 1996, 118, 2644–2653. (30) Moucheron, C. New J. Chem. 2009, 33, 235–245. € (31) Onfelt, B.; Lincoln, P.; Norden, B. J. Am. Chem. Soc. 1999, 121, 10846–10847. (32) Vos, J. G.; Kelly, J. M. Dalton Trans. 2006, 4869–4883. (33) O’Connor, N. A.; Stevens, N.; Samaroo, D.; Solomon, M. R.; Marti, A. A.; Dyer, J.; Vishwasrao, H.; Akins, D. L.; Kandel, E. R.; Turro, N. J. Chem. Commun. 2009, 2640–2642. (34) Xu, H.; Liang, Y.; Zhang, P.; Du, F.; Zhou, B. R.; Wu, J.; Liu, J. H.; Liu, Z. G.; Ji, L. N. J. Biol. Inorg. Chem. 2005, 10, 529–538. (35) Spillane, C. B.; Smith, J. A.; Buck, D. P.; Collins, J. G.; Keene, F. R. Dalton Trans. 2007, 5290–5296. (36) Mei, H. Y.; Barton, J. K. J. Am. Chem. Soc. 1986, 108, 7414– 7416. (37) Luedtke, N. W.; Hwang, J. S.; Glazer, E. C.; Gut, D.; Kol, M.; Tor, Y. ChemBioChem 2002, 3, 766–771. (38) Xu, H.; Deng, H.; Zhang, Q. L.; Huang, Y.; Liu, J. Z.; Ji, L. N. Inorg. Chem. Commun. 2003, 6, 766–768. (39) Luedtke, N. W.; Hwang, J. S.; Nava, E.; Gut, D.; Kol, M.; Tor, Y. Nucleic Acids Res. 2003, 31, 5732–5740. (40) Ardhammar, M.; Lincoln, P.; Norden, B. J. Phys. Chem. B 2001, 105, 11363–11368. (41) Svensson, F. R.; Li, M.; Norden, B.; Lincoln, P. J. Phys. Chem. B 2008, 112, 10969–10975. (42) Svensson, F. R.; Matson, M.; Li, M.; Lincoln, P. Biophys. Chem. 2010, 149, 102–106. (43) Norden, B.; Rodger, A.; Dafforn, T. Linear Dichroism and Circular Dichroism: A Textbook on Polarized-Light Spectroscopy; RSC Publishing: Cambridge, U.K., 2010. (44) Haugland, R. P. The Handbook: A Guide to Fluorescent Probes and Labeling Technology, 10th ed.; Spence, M.T.Z., Ed.; Invitrogen, Molecular Probes: Eugene, OR, 2005. (45) Boisvert, F. M.; van Koningsbruggen, S.; Navascues, J.; Lamond, A. I. Nat. Rev. Mol. Cell Biol. 2007, 8, 574–585. (46) Keene, F. R.; Smith, J. A.; Collins, J. G. Coord. Chem. Rev. 2009, 253, 2021–2035.

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