DOI: 10.1002/chem.201002149

Photoactive RuII–Polypyridyl Complexes that Display Sequence Selectivityand High-Affinity Binding to Duplex DNA through Groove Binding

Amrita Ghosh,[a] Priyadip Das,[a] Martin R. Gill,[b] Prasenjit Kar,[a] Michael G. Walker,[b]

Jim A. Thomas,*[b] and Amitava Das*[a]

Introduction

Over the last two decades, studies on the interaction ofmetal complexes with DNA have burgeoned.[1] In this con-text, systems that incorporate redox-active and photoexcita-ble octahedral d6 metal centers have attracted particular at-tention because they can add functionality to the bindingsubstrate. For example, the photoactivation of metallointer-calators derived from the {RhIII ACHTUNGTRENNUNG(phi)} (phi =9,10-phenanthri-nequinone diimine) unit results in direct cleavage of duplexDNA.[2] By using this phenomenon to probe the structure ofthe binding site, sequence-selective binding has been accom-plished.[3] By employing ligands related to phi, structuralprobes capable of recognizing mismatches with fidelity havealso been synthesized.[4] Although RuII-based photocleavagesystems have previously been produced, the majority of re-ports on such metal complexes have investigated binding-in-

duced modulation of their 3MLCT-based (MLCT =metal–ligand charge transfer) luminescence.[5]

In particular, many studies on the “DNA light switch”effect observed for metallointercalators containing the {RuII-ACHTUNGTRENNUNG(dppz)} (dppz =2,5-bis(2-pyridyl)pyrazine) unit have beenreported.[6] This work has also been extended to other d6

metal ions, with the binding properties of dppz complexesbased on ReI, OsII, and IrIII centers all having been de-scribed.[7] Related homo- and heterometallic dinuclear inter-calators are also known.[8] In contrast, much less work onmetal complexes capable of groove binding has been report-ed. This is somewhat surprising, because substrates that ex-ploit this interaction often have highly attractive bindingcharacteristics; for example, classical minor-groove binderssuch as distamycin and netropsin exhibit higher binding af-finities and selectivity than most intercalators. Their high af-finity for such sequences is mainly due to the curvature oftheir hydrophobic surface being complementary to that ofduplex DNA.[9] Generally, these binding substrates have apreference for AT-rich regions because the minor groove ofAT sequences is particularly deep and narrow, which thusfacilitates and enhances favorable van der Waals contacts.However, hydrogen-bonding sites also enhance binding af-finities and largely drive more specific sequence preferen-ces.[10]

Only a handful of RuII complexes intentionally designedfor groove binding have been reported. Hannon et al. havereported a cytotoxic helicate that recognizes DNA with high

Abstract: The duplex-DNA bindingproperties of a nonintercalating poly-pyridyl ruthenium(II) complex that in-corporates a linear extended ligandwith a catechol moiety has beenprobed with a variety of photo- andbiophysical techniques. These studiesreveal that the complex groove bindsto DNA sequences biphasically, anddisplays binding constants equivalent

to those of high-affinity metallointerca-lators. The complex also displays pref-erential binding to AT-rich sequences.Changes in the structure of the coordi-nated catechol ligand and the incorpo-

ration of intercalating ancillary ligandsinto the complex were found to modu-late both the optical-binding responseand binding parameters of the system,which indicates that the catecholmoiety plays a crucial role in the ob-served enhancement to binding affini-ties.

Keywords: DNA · groove binding ·luminescence · molecular recogni-tion · ruthenium

[a] A. Ghosh, P. Das, P. Kar, Dr. A. DasCentral Salt & Marine Chemicals Research Institute (CSIR)Bhavnagar, 364002, Gujarat (India)Fax: (+91) 2782567562E-mail : amitava@csmcri.org

[b] M. R. Gill, M. G. Walker, Dr. J. A. ThomasDepartment of Chemistry, University of SheffieldSheffield: S3 7HF (UK)Fax: (+44) 1142229346E-mail : james.thomas@sheffield.ac.uk

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affinity through major-groove binding,[11] whereas Keeneand collaborators have explored the application of opticallyresolved mono- and dinuclear RuII complexes as minor-groove binding probes for canonical duplexes, mismatches,and hairpins.[12] In recent work, the Thomas and Otsukigroups have demonstrated that a dinuclear RuII complex,which is structurally related to the minor-groove binder be-renil, functions as a colorimetric sensor for DNA sequenceand structure.[13] However, the effect of hydrogen-bondingmoieties on the physical and biophysical properties of suchsystems is virtually unstudied.

The DNA-binding properties of a compound with a pend-ant catechol moiety capable of hydrogen bonding is ex-plored herein. This complex has a catechol unit held at theend of an extended ligand, (E)-4-[2-(4’-methyl-2,2’-bipyri-din-4-yl)vinyl]benzene-1,2-diol (L1), the curvature of whichcomplements that of the duplex. The interaction of thiscomplex with both genomic duplex DNA and synthetic se-quences is explored. These studies reveal that the complexdisplays unusual biphasic binding to DNA and high affinities(Kb>107

m�1 in 50 mm tris-HCl buffer (pH 7.2; tris(hydroxy-

methyl)aminomethane) that contains 50 mm of NaCl) forspecific sequences. By changing the ancillary ligands aroundthe metal center and removing the extended linker on thecatechol-based ligand, we have also explored the consequen-ces of changing steric demand and/or shape on the bindingproperties of this system.

Results and Discussion

Synthesis : RuII–polypyridyl complexes 1, 2, 3, and 4, whichcontain the catechol ligands, L1 and 1,10-phenanthroline-5,6-diol (L2), were all synthesized as hexafluorophosphate saltsin a similar manner; details of their syntheses are describedin the Experimental Section. The complexes were character-ized by using standard spectroscopic and analytical tech-niques. The water-soluble chloride salt of 1 was then pre-pared through counteranion metathesis. Electronic spectrafor all of these complexes are dominated by typical Rudp!

bpy/L1p*-based MLCT transitions in the 380–580 nm region,

and an intra-/inter ACHTUNGTRENNUNGli ACHTUNGTRENNUNGgand (bpy/L1p!bpy/L1

p*) (bpy= bipyri-dine) transition at around 300 nm.[14] On excitation into theMLCT transition band, all these complexes (1–4) showedemission bands with maxima at around 620 nm.

Due to their lower HOMO–LUMO gap, lower quantumyields (see Experimental Section) for complexes with ex-tended conjugation is not uncommon, and this complies wellwith the energy-gap law.[15] The quantum yield of 4 in aceto-nitrile matches well with the data reported for the [Ru-ACHTUNGTRENNUNG(phen)2ACHTUNGTRENNUNG(dppz)2+] (phen =1,10-phenanthroline; f= 0.0073 inair-equilibrated acetonitrile).[14c]

Absorption titration : The interaction of the complex withDNA was initially assessed with UV/Vis spectroscopy. Asshown in Figure 1, the electronic spectrum of the complex intris buffer was found to change on addition of increasingamounts of calf-thymus DNA (CT-DNA).

The spectrophotometric titration reveals an unusual bi-phasic response: Initially, several bands, including the1MLCT transition centered at around 450 nm, display clearhypochromism; large red- and blueshifting of band maximaare also observed. Continued addition of DNA leads to adecreasing rate of spectral change until a CT-DNA concen-tration of 4.5 �10�5

m is reached; further addition of DNAtriggers a second phase of spectral changes, which continueuntil [DNA]=5.0 � 10�4

m. Complex 2 also displays two setsof equilibria, for which spectral changes terminate at [CT-DNA]= 5.1 �10�5 and 2.6 �10�4

m. For 3, the terminal [CT-DNA] for the two equilibria were 5.5 � 10�5 and 5.2 � 10�4

m.For 4, only a single equilibrium was observed at a saturationpoint of [CT-DNA]=6.6 � 10�5

m. After this point, no further

Figure 1. UV/Vis titration for a) 1, b) 2, c) 3, and d) 4 (RuII complex con-centration=1.0� 10�5

m) with CT-DNA. Blue lines= changes in spectrumup to 1st phase; red lines= changes in spectrum up to 2nd phase. Eachspectrum was corrected for increasing [DNA] during titration.

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changes occur, even on further addition of DNA. In allcases, bands at 270 and 440 nm showed hypochromicity. Bi-phasic changes in absorption spectra have occasionally beenobserved during the interaction of metallointercalators withDNA,[16] with this phenomenon being assigned to stackingbetween bound metal complexes. However, in these cases,the hypo-/hyperchromic changes are much smaller,[16a,c] andno large-scale shifts in band maxima are observed, which in-dicates that the changes in the spectra of complexes 1–4 arenot simply due to similar stacking effects.

Luminescence titrations with CT-DNA : Complex 1 also dis-plays 3MLCT-based emission. Therefore, luminescence titra-tions were carried out by using a procedure similar to theUV/Vis titrations. The changes in emission intensity inducedby interaction with CT-DNA at 25 8C were measured byusing an excitation wavelength lext = 440 nm. Again, com-plex 1 showed two distinct sets of changes on binding to CT-DNA. Initially, the addition of DNA causes large reductionsin emission intensity, but, after reaching a minimum, furtheradditions lead to increasing emission intensity until it ishigher than that observed before the addition of any DNA.Significant modulations in the structure of the emissionband also accompany the changes in intensity (see Figure 2).This is the first time such a phenomenon has been observedand reported.

Clearly, the catechol moiety of L1 has a greater effect onthe DNA-induced optical response of 1 than on other relat-ed ruthenium(II) cations. The complex [RuACHTUNGTRENNUNG(bpy)3]

2+ displaysvirtually no emission changes on binding, whereas [Ru-ACHTUNGTRENNUNG(phen)3]

2+ only displays hypochromicity in absorption titra-tions and enhancements of luminescence intensities.[17] Usu-ally, DNA-induced decreases in luminescence indicatequenching of the excited state through photooxidation pro-cesses that involve nucleobases—commonly G sites.[18] How-ever, this process can be discounted because it would re-quire a further decrease in emission at higher [DNA]/ ACHTUNGTRENNUNG[com-plex] ratios, not an increase, as is observed.

The possibility that quenching of the complex excitedstate involves the catechol unit can also be discounted; pre-viously reported time-resolved absorption studies on 1 haveshown that the catechol functionality does not influence thedynamics of photophysical processes associated, on excita-tion, with the Rudp!L1

p*/bpy-based MLCT band.[19,20] Wealso recorded the time-resolved emission-decay kinetics for1 and the analogous compound ([Ru ACHTUNGTRENNUNG(bpy)2{(E)-4-(3,4-dime-thoxystyryl)-4’-methyl-2,2’-bipyridine}]ACHTUNGTRENNUNG[PF6]2) 5,[20] in whichthe redox-active catechol moiety is substituted with theredox-inactive dimethoxy moiety. Time constants for theemission-decay profile of 1 following excitation were foundto be (3.6�0.12) ns (7%) and (150�5) ns (93%), respec-tively, while for complex 5 the respective time constants are(34�1.4) ns (12%) and (170�7) ns (88 %), respectively.The absence of any substantial differences in the excited-state decay profiles of 1 and 5 confirms that the catecholfunctionality is not involved in quenching the triplet excitedstate of 1. However, these studies do show that, when thecatechol unit of complex 1 is involved in hydrogen bonding,a decrease in luminescence (due to an increase in nonradia-tive decay) is observed;[20] this effect has also been observedin analogous complexes that incorporate other urea-based,hydrogen-bonding ligands.[21]

Changes in the emission for two different phases (initialdecrease and then enhancement in emission) as a functionof the ratio [DNA]/[Ru] are shown in Figure 2. The initialobservation of a decrease in luminescence insinuates thatthe catechol unit of 1 hydrogen bonds to available nucleo-base-acceptor residues within the minor groove of the DNAduplex; conventional minor groove binders commonly inter-act with non-Watson–Crick motif nucleobase residues in thisway. After these sites are occupied, binding that does not in-volve catechol hydrogen bonding occurs. In the latter case,emission intensities increase as the luminophore movesaway from the polar bulk solvent into a more nonpolar, sol-vent-inaccessible binding site. This is the phenomenon re-sponsible for the emission enhancements usually observedwhen cations such as [Ru ACHTUNGTRENNUNG(phen)3]

2+ interact with DNA.To quantify the binding interactions of the complex with

DNA, changes in the absorption and emission titrationswere used to construct binding curves for each phase ofbinding; see Figures 3 and 4. Thus, binding parameters forthe interaction of the complex with CT-DNA were estimatedthrough fits of the data to the McGee–von Hippel model[22]

Figure 2. Emission titration for 1 (2.5 � 10�5m) with CT-DNA. blue lines=

changes up to 1st phase (decreasing intensity) ([CT-DNA] =0–2.6 �10�5

m); red lines= changes up to 2nd phase (increasing intensity) ([CT-DNA]=2.6 � 10�5–4.0 � 10�4

m).

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FULL PAPERPhotoactive RuII–Polypyridyl Complexes

[see Eqs. (2) and (3) in the Experimental Section]; thesedata are summarized in Table 1.

The fits for the two sets of binding titrations are in goodgeneral agreement, especially for the lower-affinity processand, although values for the higher-affinity binding processagree slightly less well, it is clear that Kb1 is at least fivetimes bigger in magnitude than Kb2. Clearly, the effect ofthe catechol ligand on binding affinity is also profound; therelated [Ru ACHTUNGTRENNUNG(bpy)3]

2+ cation binds DNA solely through elec-trostatic interactions, with Kb�103

m�1.[17] With an increase

of at least three orders of magnitude in binding affinity, thisdata confirms that the binding mode of 1 is very differentfrom that of [Ru ACHTUNGTRENNUNG(bpy)3]

2+ . The large affinities suggest thatboth modes involve groove binding, whereas the differencein optical response reveals that in only one of the twomodes are hydrogen-bonding contacts made; it seems likelythat this is a major reason for the enhanced affinity of thisbinding phase.

A second possibility is that the changes in optical proper-ties are not due to two binding sites, but are due to a singlebinding site with different spectral properties for isolatedand crowded complexes. Such a model has been used to ex-plain the biexponential decay of luminescent {RuIIACHTUNGTRENNUNG(dppz)}systems when bound to DNA[23] However, in this latter case,the optical changes observed are much more subtle than thelarge changes observed for 1 binding to DNA. Furthermore,it is difficult to envisage why changes in loading could pro-duce such diametrically different optical responses. Howev-er, even if this is the case, even at low [Ru]/ ACHTUNGTRENNUNG[DNA] bindingratios, complex 1 binds to duplex DNA with an affinity thatis appreciably higher than [Ru ACHTUNGTRENNUNG(bpy)3]

2+ , which clearly indi-cates that the catechol moiety has a profound effect onDNA binding.

It should be noted that 4 has a similar catechol unit to 1,but does not show any quenching in the presence of DNA.This also confirms that the catechol functionality is not re-sponsible for the initial quenching of emission from com-pounds 1, 2, and 3 in the presence of varying [DNA].

Luminescence titrations with other polynucleotides : Asmentioned above, classical groove binders display distinctivebinding preferences towards AT-rich sequences. To furtherprobe whether these L1-containing metal complexes func-tion as groove binders and also display similar preferences,the binding properties of 1 with polynucleotides possessingvarious AT- or GC-rich sequences was also investigated.The emission-spectral response of 1 in the presence of vary-ing concentrations of the polyACHTUNGTRENNUNG(dA–dT)2, poly ACHTUNGTRENNUNG(dG–dC)2,poly(dA)–poly(dT), and poly(dG)–poly(dC) are shown inFigure 5.

Table 1. Estimated binding parameters for the biphasic interaction ofcomplex 1 with CT-DNA obtained from absorption-titration data.

Titration Kb [m�1] N [bp]

absorptionKb1=2.9 � 106 3.3Kb2=2.0 � 105 4.1

luminescenceKb1=8.6 � 105 1.5Kb2=1.6 � 105 2.9

Figure 3. Binding curves for the interaction of 1 with CT-DNA construct-ed from DNA-induced changes in the absorption spectrum of 1. Both the1st phase (top) and 2nd phase (bottom) are shown.

Figure 4. Binding curves for the interaction of 1 with CT-DNA; The areaof each emission spectrum is plotted against the [DNA]/[Ru] in its emis-sion titration. Both the 1st phase (red line shows the best fit plot withR2 =0.98) (top) and 2nd phase (red line shows the best fit plot with R2 =

0.997) (bottom) are shown.

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J. A. Thomas, A. Das et al.

The emission-spectral response of 1 on addition of poly-nucleotides is dependent on the nature of the sequence,which offers further evidence that the optical changes arenot merely a function of loading. For both polyACHTUNGTRENNUNG(dA–dT)2

and poly ACHTUNGTRENNUNG(dG–dC)2, a large initial decrease in emission inten-sity was observed. This was followed by a much smaller in-crease in intensity than that observed for the genomic DNAtitrations, with changes being significantly smaller for poly-ACHTUNGTRENNUNG(dG–dC)2.

Addition of poly(dA)–poly(dT) and poly(dG)–poly(dC)induced a different response; in both cases only a decreasein emission intensity was prominently observed. Indeed,while continued addition of poly(dA)–poly(dT) did induce anominal enhancement in emission intensity, similar experi-ments with poly(dG)–poly(dC) produced no emission en-hancement at all, even when large excesses of the polynuc-leotide were added (see Figure 5). These experiments sug-gest that the interaction with poly(dG)–poly(dC) is mono-phasic in nature. Note that, for all four polynucleotides, al-though the emission changes for the first binding phasewere at least of the same magnitude as those observed forbinding to CT-DNA, the changes in the second phase weregreatly reduced or entirely absent. However, these differen-ces are not entirely reflected in a change in the magnitudeof binding affinities (see below).

The emission data collected in these experiments wasagain used to estimate respective binding constants; thisdata is presented in Table 2. From the data, it is clear that 1does display a preference for purine–pyrimidine steps, andparticularly binds to [AT.TA] steps with very high affinities(>107

m�1), with a fiftyfold preference for this alternating

step sequence over A tracks and G tracks. These data are

consistent with those obtained for conventional minor-groove binders,[9] and reflect the structural nature of thesesequences; it is known that alternating purine–pyrimidinesteps, especially those containing the AT pair, are particular-ly flexible, which is exemplified by the TATA box se-quence.[24] Although nonalternating sequences have con-trasting properties (due to bifurcated hydrogen bonding, Atracks display a compressed minor groove and widenedmajor groove[25] , whereas extended G tracks possess anarrow and deep major groove and a widened minorgroove[26]) they are both more rigid than the alternatingforms. Thus, complex 1 binds at flexible sites most able toaccommodate the steric demand of the metal complexwithin the minor groove. Although the site sizes obtained ina number of the titrations that involve the polynucleotidesare anomalously low, site sizes lower than one have been re-ported by using the McGhee–von Hippel fits in a number ofcases.[27]

Modifying the catechol and ancillary ligands : To investigatewhether a modulation in binding properties of the newsystem could be effected by a change in ancillary ligand, theCT-DNA binding properties of the new complex 2 werestudied. We postulated that, by substituting the bpy ancillaryligands of 1 with the more sterically demanding phen li-gands, a higher selectivity towards more open and accessiblesites may occur; a phenomenon that has been observed for{RhIII ACHTUNGTRENNUNG(phi)} systems.[28] To discover how much of the changein DNA-binding properties of 1, relative to [Ru ACHTUNGTRENNUNG(bpy)3]

2+ , isattributable to the curved shape of L1 or solely to the cate-chol group of L1, similar studies were also carried out usingcomplex 3, which contains ligand L2 with a catechol attachedmore directly to the metal center. Finally, the effects ofadding an intercalator moiety into the original groove-bind-ing system were probed using the newly synthesized com-plex 4, which contains the well-established intercalative unitdppz.

Following the syntheses of 2, 3, and 4, emission titrationsinvolving CT-DNA revealed that the optical response toDNA differed in each case. Luminescence titrations re-vealed that 2 and 3 displayed behavior closest to 1. For com-plex 2, a relatively small decrease in the emission maximum,at 606 nm, was observed for the first equilibrium process.On further addition of DNA, whereas no shift in lmax oc-curred, large enhancements in the emission intensity were

Figure 5. Changes in the emission spectrum of 1 on addition of selectedpolynucleotides. a) poly ACHTUNGTRENNUNG(dA–dT)2, ([DNA]=0–7.3 � 10�7

m ; 7.3� 10�7–7.8� 10�6

m), [1] =2.5� 10�5m ; b) polyACHTUNGTRENNUNG(dG–dC)2, ([DNA]=0–9.1 � 10�7

m ;9.1� 10�7–8.6 � 10�6

m), [1]=2.0 � 10�5m ; c) poly(dA)–poly(dT), ([DNA]=

0–1.1 � 10�6m ; 1.1� 10�6–3.6 � 10�6

m), [1]=1.0 � 10�5m. d) poly(dG)–

poly(dC), ([DNA] =0–8.8 � 10�6m ; [1]=1.0 � 10�5

m.

Table 2. Estimated binding parameters for the biphasic interaction of 1with polynucleotides obtained from emission-titration data.

DNA Kb [m�1] N [bp]

poly ACHTUNGTRENNUNG(A–T)2 with (1)Kb1 =2.4� 107 0.69Kb2 =1.03 � 106 1.5

poly ACHTUNGTRENNUNG(G–C)2 with (1)Kb1 =7.1� 106 1.05Kb2 =6.2� 105 0.49

poly(dA)–poly(dT) with (1)Kb1 =2.3� 106 0.89Kb2 =4.9� 105 0.73

poly(dG)–poly(dC) with (1) Kb =8.5� 105 1.5

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FULL PAPERPhotoactive RuII–Polypyridyl Complexes

observed, until no further changes were observed at [CT-DNA] of 3.0 � 10�4

m (see Figure 6).In contrast to both 1 and 2, although two phases of spec-

tral change were also observed for 3, the magnitude ofchange for both of these phases was relatively small (seeFigure 7).

Compared with 1, 2, and 3, complex 4 displays a very dif-ferent response to binding. Unlike the parent compound[Ru ACHTUNGTRENNUNG(bpy)2ACHTUNGTRENNUNG(dppz)]2+ , complex 4 does display some initial lu-minescence in aqueous solution, but, on addition of DNA, itshows a large increase in luminescence emission; the charac-teristic “light-switch” effect is a marker for the interactionof {RuII ACHTUNGTRENNUNG(dppz)} with DNA[6–8] (see Figure 8). This effect isusually taken as the sign of an intercalative binding interac-tion that involves the {RuII ACHTUNGTRENNUNG(dppz)} unit, even though interca-lation is not always required.[28]

The luminescence titration data obtained for all threenew complexes was fitted to the McGhee–von Hippel equa-tion to give the binding parameters outlined in Table 3.

As our results show, changing the ligand set has a largeeffect on the characteristics and magnitude of DNA binding.A comparison of the binding properties of 1 and 2 showsthat, although affinities are comparable, the reduction in

emission intensity for 2 in the first binding phase is appreci-ably smaller than for 1. More interestingly, for both bindingequilibria, the binding sites for 2 are appreciably larger thanfor 1. This observation insinuates that 2 is interacting withDNA more selectively than 1, since, although binding affini-ties are virtually equivalent, binding sites are now, on aver-age, up to 14 bp (bp =base pairs) apart, instead of every 2–3 bp.

A comparison of 1 and 3 is also informative. Althoughthe optical changes for 3 are significantly lower in magni-tude than those for 1, the actual binding parameters arealmost identical. The magnitude of the change in emissiondata reflects two factors: the strength of hydrogen bond-ing;[16,17] and how much the excited state is isolated fromcontact with bulk solvent.[29] The luminescence titrationssuggest that 1 binds to DNA with a stronger hydrogen-bond-ing interaction, and is more isolated from the bulk solventin the second binding site. This indicates closer contact tothe DNA groove, which is consistent with the elongatedform of L1 compared to L2. However, somewhat surprisingly,the binding-affinity data for the two complexes is almostidentical, which indicates that the enhanced binding of 1(and 2), relative to the [RuACHTUNGTRENNUNG(bpy)3]

2+ cation, is to a largeextent due to the inclusion of the catechol moiety.

Finally, although 4 now appears to interact with DNAthrough the dppz ligand in a single phase of intercalativebinding, its binding affinity is two orders of magnitude lowerthan both 1 and previously reported {RuII ACHTUNGTRENNUNG(dppz)} sys-tems.[5–7, 30] This is also caused by steric effects, which modu-late DNA binding. Clearly, if L1 of 4 slots into the narrowminor groove, the octahedral coordination geometry of thecomplex means that the bulky and elongated dppz ligandwill project into the groove wall, which would produce large

Figure 6. Emission titration for 2 (1.0 � 10�5m) with CT-DNA. Blue

lines=changes in spectrum up to 1st phase (decreasing intensity) ([CT-DNA]=0–4.6 � 10�5

m); red lines= changes in spectrum up to 2nd phase(increasing intensity) ([CT-DNA] =4.6� 10�5–3.4 � 10�4

m).

Figure 7. Emission titration of 3 (2.5 � 10�5m) with CT-DNA. blue lines=

changes up to 1st phase (decreasing intensity) ([CT-DNA] =0–2.4 �10�5

m); red lines= changes up to 2nd phase (increasing intensity) ([CT-DNA]=2.4 � 10�5–6.2 � 10�4

m).

Figure 8. Emission titration for 4 (1.0 � 10�5m) with CT-DNA. ([CT-

DNA]=0–8.8 � 10�5m).

Table 3. Estimated binding parameters from luminescence titrations ofcomplexes 2, 3, and 4 with CT-DNA.

Compound Kb [M�1] N [bp]

2Kb1=3.3 � 105 7.7Kb2=4.0 � 105 14.0

3 Kb1=8.2 � 105 1.2Kb2=2.0 � 105 4.1

4 Kb =6.5 � 104 2.2

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J. A. Thomas, A. Das et al.

unfavorable steric interactions. Consequently, intercalationbecomes the favored binding motif for the complex. Howev-er, even in intercalative binding mode, unfavorable steric in-teractions dominate, as now the sterically demanding L1 isan ancillary ligand. In such cases the overlap of aromaticbase residues with dppz are restricted, and there is a conse-quent lowering of binding affinities.[31]

Viscosity : To confirm the assigned binding modes of com-plexes 1, 2, 3, and 4, viscosity studies were carried out.Whereas intercalators increase the viscosity of short rodlikeDNA by increasing hydrodynamic lengths, classical groovebinders do not perturb the structure of the duplex on bind-ing; therefore, the addition of groove binders does notaffect the viscosity of DNA solutions.[32] The effects of 1, 2,and 3, and the established minor groove binder H33258 onthe relative viscosity of CT-DNA are shown in Figure 9.

This shows that, in these four cases, the addition of the bind-ing substrate produces no overall change in relative viscosityof the DNA, which thus confirms that the complexes areindeed groove binders. Contrastingly, complex 4 produces adistinct increase in viscosity; this confirms that it is interca-lating with the DNA through the {RuII ACHTUNGTRENNUNG(dppz)} unit.

Conclusion

The previously reported photoactive RuII–polypyridyl com-plex 1, which contains a coordinated ligand L1 with a pend-ant catechol functionality, was studied as a binding substratefor CT-DNA, and for the four different polynucleotidespoly ACHTUNGTRENNUNG(dA–dT)2, poly ACHTUNGTRENNUNG(dG–dC)2, poly(dA)–poly(dT), andpoly(dG)–poly(dC). It was found that the complex binds toCT-DNA in a unique biphasic manner. In the first bindingphase, the characteristic luminescence of the complex pro-gressively decreases. This is consistent with the catecholfunctionality of the coordinated ligand L1 participating inhydrogen bonding with non-Watson–Crick sites of theduplex minor groove. In the second phase of binding, the lu-minescence intensity increases, which is typically seen for

minor groove binders because they move into the solvent-isolated, hydrophobic environment of DNA. Studies withthe polynucleotides reveal that 1 preferentially binds tomore flexible, alternating purine–pyrmidine sequences, par-ticularly those that contain AT.TA steps, with very high af-finities (Kb>107

m�1).

To investigate the possible modulation of binding throughstructural changes to the basic molecular architecture of 1,the interactions with CT-DNA of complexes 2, 3, and 4were also investigated. In these complexes, either the ancil-lary ligands or L1 were exchanged for related structures.These studies revealed that the ancillary ligands can affectthe binding affinity and selectivity of the system, and suggestthat the catechol unit plays a large part in the distinctivebinding properties of these systems.

Further investigations into the details of the unusual bind-ing modes of complexes such as 1–3 and their unique opticalproperties are currently underway. These initial studies havebeen carried out on a racemic mixture of the L and D formsof complexes 1 to 4 ; routes to the isolation of resolved sam-ples and methods to modulate the binding properties ofthese systems will be outlined in future publications.

Experimental Section

Analytical measurements : 1H NMR spectra were recorded on Bruker200 MHz FT NMR (model: Advance-DPX 200) spectrometer at RT(25 8C). TMS was used as an internal standard for all 1H NMR spectro-scopic studies. ESIMS measurements were carried out on a Waters QTof-Micro instrument. Microanalysis (C, H, N) was performed using aPerkin–Elmer 4100 elemental analyzer. IR spectra were recorded byusing KBr pellets on a Perkin–Elmer Spectra GX 2000 spectrometer.UV/Vis spectra were obtained by using either a Shimadzu UV-3101 PCor Cary 500 Scan UV/Vis–NIR spectrometer. RT emission spectra wereobtained using a FluoroLog Horiba Jobin Yvon luminescence spectro-fluorimeter.

Materials and methods : [Ru ACHTUNGTRENNUNG(bpy)2]Cl2·2H2O and [Ru ACHTUNGTRENNUNG(phen)2Cl2] wereprepared according to standard literature procedures.[33] RuCl3·xH2O,bpy, phen, and CT-DNA were purchased from Aldrich (USA), and wereused as received. [tBu4N]PF6 was recrystallized from ethanol before use.K4[Fe(CN)6], oleum, nitric acid, and acetic acid were obtained from S.D.Fine Chemicals (India), and were used without further purification. Allsolvents were dried and distilled prior to use following standard proce-dures. pBR322 plasmid DNA was obtained from Bangalore Genei(India). The CT-DNA concentration per nucleotide was determined byabsorption spectroscopy by using the molar absorption coefficient(6600 mol�1 dm3 cm�1) at 260 nm.[34a] The absorption and emission datawere used to construct nonlinear Scatchard plots (r/c versus r) and fittedto the McGhee–von Hippel model, in which neither the site size or bind-ing constant was set. Each titration was carried out several times, andonly data that led to R2> 95% was accepted. Uncertainties were esti-mated at 20%.

The ligands L1, L2, dppz,[6] and [Ru ACHTUNGTRENNUNG(dmso)2Cl2] were synthesized accord-ing to reported procedures.[20b, 35, 36]

Synthesis of 1: [Ru ACHTUNGTRENNUNG(bpy)2Cl2]·2H2O (0.07 g, 0.072 mmol) and L1 (0.02 g,0.07 mmol) dissolved in ethanol–water (50 mL, 1:1, v/v) were heated atreflux for 8 h with constant stirring under an inert atmosphere. The etha-nol was then removed under vacuum, and the desired crude complex wasprecipitated as a red–orange solid by adding an excess aqueous solutionof KPF6. The solid was filtered off, washed with cold water, and air dried.This crude product was further purified by gravity chromatography withsilica as the stationary phase and CH3CN/saturated aqueous solution of

Figure 9. Relative viscosity of CT-DNA on addition of 1, 2, 3, 4, and theestablished minor groove binder H33258.

Chem. Eur. J. 2011, 17, 2089 – 2098 � 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim www.chemeurj.org 2095

FULL PAPERPhotoactive RuII–Polypyridyl Complexes

KPF6 (98:2, v/v) as eluent. The acetonitrile was removed under vacuum,and the desired pure complex was extracted into dichloromethane. Thedichloromethane was removed under reduced pressure to afford the purecompound (0.070 g, 52%). 1H NMR (CD3CN): d=8.5 (d, J =6.6 Hz, 6H;2H6 (bpy), 2H6’ (bpy) H6 and H6’ L1), 8.07 (t, J =7.6 Hz, 4 H; 2H4 (bpy),2H4’ (bpy)), 8.01 (s, 1H; H3 (L1)), 7.82 (d, J=5.8 Hz, 1 H; H3’ (L1)), 7.76–7.56 (m, 5 H; 2H5 (bpy), 2H5’ (bpy), HCH=CH), 7.43–7.35 (m, 6H; 2H3

(bpy), 2H3’ (bpy), 2H5,5’ (L1)), 7.23 (d, J=8.2 Hz, 1 H; H5 (phenyl)), 7.03(d, J= 8.2 Hz, 1H; H6 (phenyl)), 7.13 (d, J=16.6 Hz, 1 H; HCH=CH), 6.86(d, J=8.2 Hz, 1H; H3 (phenyl)), 2.56 ppm (s, 3H; CH3); FTIR (KBr):n= 3449 (�OH), 1603 (C=C, C=N), 844 cm�1 (PF6); elemental analysiscalcd (%) for RuC39H32N6O2P2F12: C 46.5, H 3.2, N 8.3; found: C 46.7, H3.2, N 8.1; ESIMS: m/z (%): 863 [M�PF6]

+ (15), 717 [M�2PF6]+ (5).

Synthesis of 2 : [Ru ACHTUNGTRENNUNG(Phen)2Cl2]·2H2O (0.318 g, 0.55 mmol) and L1 (0.170 g,0.55 mmol) dissolved in ethanol–water mixture (50 mL, 1:1, v/v) wereheated at reflux for 24 h with constant stirring under an inert atmosphere.The ethanol was then removed under vacuum, and the desired crudecomplex was precipitated as a red–orange solid by adding an excessaqueous solution of KPF6. The solid was filtered off, washed with coldwater, and air dried. This crude product was further purified by gravitychromatography with neutral Al2O3 (grade III) as the stationary phaseand CH3CN as eluent. The acetonitrile was removed under vacuum andthe desired pure complex was extracted into dichloromethane. The di-chloromethane was removed under reduced pressure to afford the purecompound (0.29 g, 44 %). 1H NMR (CD3CN,): d=10.24 (s, 1H; H3’(L1)),9.84 (s, 1 H; H3(L1)), 9.73 (s, 1H; Hphenyl(L1)), 8.86 (d, J =10 Hz, 1 H;Hphenyl(L1)), 8.75 (d, J =10 Hz, 2 H; HPhen), 8.69 (d, J=9 Hz, 2H; HPhen),8.43 (d, J =8 Hz, 2H; HPhen), 8.29 (d, J=8 Hz, 4 H; HPhen), 8.24–8.09 (m,6H; HPhen), 7.91 (s, 1 H; H�CH=CH�(L1)), 7.75–7.73 (m, 2 H; H5’(L1),Hphenyl(L1)), 7.43–7.41 (m, 1 H; H5(L1)), 7.35–7.32 (m, 2H; H�CH=CH�(L1),Hphenyl (L1)), 2.214 ppm (s, 3H; CH3); FTIR (KBr): n=3449 (�OH), 1604(C=C, C=N), 846 cm�1 (PF6); elemental analysis calcd (%) forRuC43H32N6O2P2F12: C 48.9, H 3.2, N 8.0; found: C, 49.1; H, 3.1; N, 8.0;ESIMS: m/z (%): 1055 [M+1]+ , 805 [M�2PF6+K+]+ .

Synthesis of 3 : [Ru ACHTUNGTRENNUNG(bpy)2Cl2]·2H2O (0.113 g, 0.21 mmol) and L2 (0.046 g,0.21 mmol) dissolved in DMF (20 mL) were heated at reflux for 8 h withconstant stirring under an inert atmosphere. The DMF was removedunder vacuum, and the desired crude complex was precipitated as a red–orange solid by adding an excess aqueous solution of KPF6. The solidwas filtered off, washed with cold water, and air dried. This crude prod-uct was further purified by gravity chromatography with silica as the sta-tionary phase and CH3CN/saturated aqueous solution of KPF6 (98:2, v/v)as eluent. The acetonitrile was removed under vacuum and the desiredpure complex was extracted into dichloromethane. The dichloromethanewas removed under reduced pressure to afford the pure compound(0.080 g, 37%). 1H NMR (CD3CN): d=8.5 (d, J =6.6 Hz, 6H; 2H6 (bpy),2H6’ (bpy) 2H6,6’(L2)), 8.09 (t, J=8 Hz, 5 H; 2H4, (bpy), 2H4’ (bpy), H5

(bpy)), 7.97 (d, J =8.5 Hz, 2 H; H3,3’ (bpy), 7.83 (d, J=5.8 Hz, 2H; H5,5’(bpy)), 7.74 (d, J =8.2 Hz, 2H; 2H3,3’ (bpy), 7.62 (d, J=8 Hz, 1H; H4

(L2), 7.58 (d, J =8.3 Hz, 1H; H4’ (L2)) 7.46–7.39 ppm (m, 3 H; H5’, ACHTUNGTRENNUNG(bpy),

H5,5’ (L2)); FTIR (KBr): n=3449 (�OH), 1601 (C=C, C=N), 847 cm�1

(PF6); elemental analysis calcd (%) for RuC32H25N6O2P2F12: C 41.98, H2.64, N 9.18; found: C 41.5, H 2.4, N 9.3; ESIMS: m/z (%): 625[M�2PF6]

+ (30).

Synthesis of 4 : [Ru ACHTUNGTRENNUNG(bpy) ACHTUNGTRENNUNG(dmso)2Cl2] (0.162 g, 0.33 mmol) was dissolved inDMF (15 mL) in a 100 mL round-bottomed flask fitted with a condenser.A solution of dppz (0.094 g, 0.33 mmol) in DMF (10 mL) was added tothe flask. The mixture was heated for 5 h at 95 8C with constant stirringunder an inert atmosphere. A solution of L1 (0.102 g, 0.33 mmol) in theminimum volume of DMF was added, and the reaction stirred at 115 8Covernight. The DMF was removed, and the product was dried under re-duced pressure. The crude solid was redissolved in methanol/water(25 mL, 1:4, v/v). The desired complex was precipitated by adding anexcess aqueous solution of KPF6. The product was filtered off, washedwith cold water, and air dried. This crude product was further purified bycolumn chromatography with silica as the stationary phase and CH3CN/saturated aqueous solution of KPF6 (98:2, v/v) as eluent (0.100 g, 22%).1H NMR (CD3CN): d =9.6 (dd, J=4.0 Hz, 2H; dppz), 8.48 (m, 6 H; bpy),

8.34 (s, 1 H; H3 (L1)), 8.16 (m, 4 H; dppz), 7.88 (m, 1H; HCH=CH), 7.76 (m,2H; dppz), 7.74 (m, 2 H; L1), 7.54 (m, 3 H; 2H5 (bpy), 1H (L1)), 7.24 (m,2H; L1),7.12 (s, 2 H; dppz), 7.10–7.035 (m, 3 H; phenyl, L1), 6.84 (d, J=

16.6 Hz, 1H; HCH=CH), 2.52 ppm (s, 3 H; CH3); FTIR (KBr): n =3436(OH), 1600 (C=C, C=N), 843 cm�1 (PF6); elemental analysis calcd (%)for RuC47H34N8O2P2F12: C 49.79, H 3.02, N 9.88; found: C 49.5, H 3.1, N9.7; ESIMS: m/z (%): 989 [M�PF6]

+ (15), 1012 [M�PF6+Na+]+ (10),1134 [M]+ (5).

The fluorescence of compounds 1, 2, 3, and 4 was measured in air-equili-brated acetonitrile solution. Relative quantum yields were calculated byusing Equation (1):

�f ¼ �f0ðIsample=IstdÞðAstd=AsampleÞðh2

sample=h2stdÞ ð1Þ

in which ff’ is the absolute quantum yield for [RuACHTUNGTRENNUNG(bpy)3ACHTUNGTRENNUNG(PF6)2], in aceto-nitrile (0.06), which was used as a reference; Isample and Istd are the inte-grated emission intensities; Asample and Astd are the absorbances at the ex-citation wavelength. Values of h2

sample and h2std, the respective refractive

indices, are considered to be same in this study. Quantum yields for 1, 2,3, and 4 were found to be 0.003, 0.0028, 0.0024, and 0.0054, respectively.

The observed fluorescence is assumed to be a sum of the weighted contri-butions of free (cf) and bound RuII complex (cb), as shown in Equa-tion (2):

cb ¼ c½ðI�I0Þ=ðImax�I0Þ� ð2Þ

in which c is the total RuII complex concentration, I and I0 are the emis-sion intensities in the presence and absence of DNA, and Imax is the fluo-rescence of the totally bound complex. The concentration of the freecomplex, cf, is equal to c�cb. A plot of r/cf versus r, in which r iscb/ ACHTUNGTRENNUNG[DNA], was constructed according to the McGhee–von Hippel equa-tion [Eq. (3)]:

2r=cf ¼ Kbð1�2nrÞ½ð1�2nrÞ=f1�2ðn�1Þrg�n�1 ð3Þ

in which Kb represents the intrinsic binding constant of the complex withDNA and n is the size of a binding site in base pairs.

DNA binding studies : Solutions of CT-DNA were obtained from Sigma–Aldrich (USA). The DNA concentration per nucleotide was determinedby absorption spectroscopy, for which a molar absorption coefficientvalue of 6600 mol�1 dm3 cm�1 at 260 nm was used.[34a] The ratio of the ab-sorbances at 260 (A260) and 280 nm (A280) for CT-DNA was used to checkthe purity of the DNA solution with respect to protein impurities. Simi-larly, for the other polynucleotides, the molar absorption coefficientvalues were 6550, 8400, 6000, and 7000 mol�1 dm3 cm�1 for poly ACHTUNGTRENNUNG(dA–dT)2,poly ACHTUNGTRENNUNG(dG–dC)2, poly(dA)–poly(dT), and poly(dG)–poly(dC), respectively,at 260 nm.[34b,c] tris-buffer (5 mm, pH 7.1, 25 mm NaCl) was used for theabsorption and fluorescence titration experiments.

Acknowledgements

A.D. acknowledges Department of Biotechnology, New Delhi, India.M.W. and M.G. are grateful to the EPSRC for QUOTA and DTC stu-dentships, respectively. We acknowledge the support of The BritishCouncil/DST (New Delhi) support through the DST-UKIERI scheme,which has enabled visits by A.G., P.D., and A.D. to J.A.T�s lab, and byM.W. and J.A.T. to A.D.�s lab.

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Received: April 15, 2010Revised: September 7, 2010

Published online: January 19, 2011

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