Synthesis, characterization, and crystal structure of RNA targeted L- and D-phenylalanine-(1,10-phen)–copper(II) conjugate complexes: comparative in vitro RNA binding profile of enantiomers and their biological evaluation by morphological studies and antibacterial activity

Abstract

New ternary chiral Cu(II) complexes 1a and b derived from L- and D-phenylalanine and 1,10-phenanthroline were synthesized and characterized thoroughly by single crystal X-ray diffraction and other spectroscopic techniques viz. UV-vis, IR, EPR, ESI-MS and elemental analysis. The complexes crystallized in the monoclinic P21 space group, possessing the lattice parameters a = 5.74690(10) Å, b = 20.6365(2) Å, c = 9.28010(10) Å, α = γ = 90°, and β = 98.2040(10) in complex 1a and a = 5.728(5) Å, b = 20.587(5) Å, c = 9.252(5) Å, α = γ = 90°, and β = 98.308(5) in complex 1b per unit cell, respectively. Comparative in vitro RNA binding studies of the L- and D-enantiomeric complexes, 1a and b, were carried out by a variety of optical spectroscopy techniques viz. UV-vis, fluorescence, and circular dichroism. Because copper is a redox metal ion, cyclic voltammetry was employed to evaluate the enantioselective RNA binding of the complexes. The results demonstrated that the L-enantiomer of Cu(II) complex, 1a, binds more strongly to the t-RNA motif than the D-enantiomer, thereby underlining the differential disposition of the enantiomers and the site preference of RNA for the L-enantiomer over the D-enantiomer. Furthermore, the comparative K_b, K and K_sv values of the L- and D-complexes demonstrated significant increases for the L-enantiomer of the copper complex, 1a, in comparison to its D-enantiomeric form, 1b. SEM analysis divulged surface morphological alteration of complexes 1a and b, evidenced by the formation of hollow tubes and a concrete-like structure with the RNA condensate, which was less pronounced in SEM micrographs of the complex 1b condensate. Complexes 1a and b were evaluated by the agar well diffusion method and demonstrated significant antibacterial activity.

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Introduction

The interaction of transition metal complexes with nucleic acids is one of the most interesting areas of bioinorganic chemistry and medicinal chemistry owing to the possible applications of these complexes as therapeutic agents for drug design,^1,2 nucleic acid structural probes³ and artificial nucleases.^4–6 Nucleic acids, particularly DNA and RNA, are primary targets for most drugs for treating infectious diseases,^7,8 viz., HIV, AIDS, hepatitis C and cancer.⁹ RNA, particularly microRNA (miRNA), plays an indispensable role in various biological processes, including development, cell proliferation, differentiation and apoptosis. Thus, altered miRNA expression is likely to contribute to many human diseases, including cancer.^10–12 RNA differs from CT-DNA not only in the composition of the bases but also in structure. CT-DNA has a B-form double-helical configuration with a wide major groove and a relatively narrow minor groove, while yeast t-RNA has an A-form conformation with an L-shaped tertiary structure (mainly unispiral), a deep major groove, and a wide, shallow minor groove. These differences are expected to lead to different binding modes and affinities.^13,14

The in vivo structures of DNA and RNA are very different; whereas DNA typically remains in the helical double stranded form, RNA folds into diverse structures, adopting folds that are similar to proteins; this is responsible for its diverse functions in cells.¹⁵ The specific recognition of pockets available in RNA by suitable ligands, involving non-covalent binding usually based on van der Waals forces, electrostatic interactions and surface complementarity, is considered to be an important asset for RNA targeted therapeutics.¹⁶ Therefore, studies on RNA–metal complex interactions could be highly instrumental for envisaging drug design strategies.¹⁷

Antibiotic resistance, particularly multi-drug resistance (MDR) of pathogenic bacteria, is one of the most crucial problems facing modern health care. Hence, the focus of pharmaceutical R&D is to discover and develop new improved antibiotics/anti-infectives with specific biological targets, viz. Riboswitch RNAs¹⁸ and RNA secondary structures, such as bulges and helices.^14,19 Previous literature reports reveal that most antibacterials exert their activity by modulating ribosome functions and RNA fragments/motifs by the binding of a suitable ligand framework, which acts as a pharmacophore. A combinatorial synthetic strategy involving several ligand fragments, in our case L- and D-phenylalanine (as a chiral centre and metal binding domain), 1,10-phenanthroline (as a recognition element that induces intercalation) and a redox active copper centre, is highly directed for specific interactions at RNA binding sites. Both the L- and D-enantiomers were proven to be good anti-bacterial agents against Gram-positive Bacillus subtilis [MTCC 121] and Staphylococcus aureus [NRRLB 767] as well as Gram-negative Escherichia coli [K12] and Pseudomonas aeruginosa. Because metal complexes (electropositive tendencies) attract bacterial membranes and other nucleic acid and protein targets, they are logical options for metal-based therapeutics which can perform dual antibacterial and antitumor functions.²⁰ The metal centre acts as a scaffold, holding the 3D structure of the biological ligands firmly in place and tuning/synchronizing the drug entity to reach a specific site of action. Chiral molecules play a critical role in the exploitation of three-dimensional space at the target site and regulate stereoselectivity in a highly organized fashion.²¹ Furthermore, the use of ancillary chiral amino acid ligands provides insight into the mechanism of action, allowing discrimination between unspecific interactions which are common to both enantiomers and specific contacts that give rise to enantioselectivity. An amino acid with a side chain aromatic ring, e.g. phenylalanine, contributes mainly to the stabilization of proteins through hydrophobic interactions and the formation of hydrophilic environments.²² These ligands play a crucial role in modulating the hard/soft properties of the metal ions and the lipophilic/hydrophilic balance of the metal complexes, thereby controlling their solubility as well as their permeability to cross cell barriers.

Copper is a biological redox element and has efficacious biocidal properties. Because of its biological congruity, a large number of copper(II) complexes have been synthesized and their biological activities have been scrutinized.^23–25 Among all these complexes, copper(II) complexes containing amino acids and 1,10-phenanthroline ligand have been receiving considerable interest due to their biological relevance, good nucleic acid binding, and antimicrobial and anticancer activities.^26–28 Recently, J. R. Aldrich-Wright et al. reported a number of copper(II) analogs with different derivatives of 1,10-phenanthroline ligand. The antimicrobial properties of these complexes against bacterial strains of S. aureus, E. coli, P. aeruginosa, and yeast S. cerevisiae were characterized, as well as the activity of their phen-based intercalating ligands.²⁹

In the continuation of our recent interest in RNA targeted chemotherapeutics,^30–32 we have carried out the synthesis, single X-ray diffraction studies and in vitro t-RNA binding studies of enantiomeric Cu(II) complexes derived from L- and D-phenylalanine and the intercalating ligand 1,10-phenanthroline, which has proven antibacterial activity. Both complexes are highly soluble in most organic solvents and are readily soluble in water, which is a prerequisite condition for the design of novel therapeutics, as water soluble complexes show superior ability to cross biological membranes. The binding propensities of the L- and D-conformations with yeast t-RNA were studied by UV-vis absorption and emission, circular dichroism spectroscopy and cyclic voltammetry.

Results and discussion

Synthesis and characterization

Complexes 1a and b were synthesized by reacting methanolic solutions of L- and D-phenylalanine (deprotonated by NaOH), 1,10-phenanthroline and Cu(NO₃)₂·3H₂O in a 1 : 1 : 1 stoichiometric ratio under reflux conditions, as depicted in Scheme 1. A dark blue transparent solution was formed and maintained at room temperature. Crystals of complexes 1a and b suitable for single X-ray crystallography were obtained by slow evaporation of the reaction mixture. The synthesized complexes were found to be stable towards air and soluble in organic solvents, such as MeOH, DCM, DMF and DMSO. The molecular structures of both complexes were established by single X-ray diffraction studies, which corroborated well with the elemental analysis and other spectroscopic measurements. The syntheses were repeated several times and were found to be reproducible.

Scheme 1 Synthetic route of complexes 1a and b.

A single crystal X-ray structural study revealed that complexes 1a and b both crystallized in the monoclinic P21 space group, possessing the lattice parameters a = 5.74690(10) Å, b = 20.6365(2) Å, c = 9.28010(10) Å, α = γ = 90°, and β = 98.2040(10) and a = 5.728(5) Å, b = 20.587(5) Å, c = 9.252(5) Å, α = γ = 90°, and β = 98.308(5) per unit cell, respectively. An ellipsoid view of complexes 1a and b along with the atom numbering scheme is shown in Fig. 1. The packing diagram is depicted in Fig. S1,† and selected bond lengths and angles of complexes 1a and b are listed in Tables S1–S4,† respectively. The complexes are mononuclear, and the asymmetric units of complexes 1a and b contain one NO₃⁻ ion, one water molecule and a Cu(II) ion; they possess a distorted square pyramidal geometry in which L- and D-phenylalanine, respectively, coordinate through the amino nitrogen and the deprotonated carboxylato oxygen atom, i.e. the amino acid is N, O chelated, forming a five-membered ring with an “envelope” conformation. The ligands 1,10-phen [Cu(1)–N(1) = 2.029(3) Å, Cu(1)–N(2) = 2.004(3) Å] and L-phe [Cu(1)–O(2) = 1.941(2) Å, Cu(1)–N(3) = 1.993(3) Å] in complex 1a and 1,10-phen [Cu(1)–N(2) = 2.025(3) Å, Cu(1)–N(3) = 2.001(3) Å] and D-phe [Cu(1)–O(1) = 1.940(2) Å, Cu(1)–N(1) = 1.985(3) Å] in complex 1b form the square base, while the water molecule has a long axial coordination [Cu(1)–O(1) = 2.199(3) Å] and [Cu(1)–O(1) = 2.191(3) Å], respectively. The values of the C–O and C–N bonds are in agreement with those in the literature for similar geometries.³³ The square base formed by the coordinated atom in complexes 1a and b is almost planar, with deviations of 0.063 Å and 0.066 Å, respectively; the Cu(1) atom is out of the plane of N1N2N3O2 by ca. 0.24 Å and 0.23 Å, respectively, towards the axial water molecule. Further, extensive hydrogen bonding (NH⋯O, O–H⋯O, C–H⋯O, O–H⋯N) interactions involving the amine nitrogen, carboxylato oxygen, distorted nitrato oxygen, and lattice water molecule stabilize the molecules in the crystal lattice.

Fig. 1 ORTEP view of (a) complex 1a and (b) complex 1b with partial numbering. Solid thermal ellipsoids are reported at the 50% probability level. Hydrogen atoms have been omitted for clarity.

The IR spectra of complexes depicted diagnostic bands in the range of 3239 to 3131 cm⁻¹, which were attributed to the coordinated NH₂ group vibrations of L- and D-phenylalanine, and two broad bands at 3469 and 3467 cm⁻¹, ascribed to the stretching vibration of the water molecule in complexes 1a and b, respectively. The IR spectra also facilitated prediction of the coordination mode of the carboxylate group. The characteristic bands of the carboxylate group were observed in the range of 1580 to 1630 cm⁻¹ for the asymmetric stretch and 1358 to 1410 cm⁻¹ for the symmetric stretch; the differences in the stretching frequencies of 1620 to 1369 cm⁻¹ in both the complexes (Δν = ν_asym − ν_sym) suggested a value >200, which indicated the terminal (monodentate) coordination mode of the carboxylate group of L- or D-phenylalanine with the metal in both complexes.³⁴ Strong bands at 912, 858, 765 and 705 cm⁻¹ were identified as the aromatic hydrocarbons with the motion of the ring hydrogens in the complexes. In the far-IR region, the presence of medium intensity bands at 535 and 428 cm⁻¹ correspond to Cu–N and Cu–O vibrations, respectively, in both complexes.³²

The ESI-MS spectra of complexes 1a and b (Fig. S2 and S3†) displayed a prominent peak at m/z 487, corresponding to the molecular ion peak [C₂₁H₂₀CuN₄O₆]⁺. Other fragmentation peaks for complexes 1a and b were observed at m/z 425 and 407, which could be assigned to [C₂₁H₂₀CuN₄O₆–NO₃]⁺ and [C₂₁H₂₀CuN₄O₆–NO₃–H₂O]⁺, respectively.

The solid state X-band EPR spectra of L- and D-Cu(II) complexes 1a and b were recorded in the solid state at room temperature (RT) on an X-band frequency of 9.1 GHz under a magnetic field strength of 3000 G (Fig. S4†). In the solid state, both complexes showed an isotropic band centered at g = 2.05.³⁵ These values were in good agreement with the unpaired electrons mainly located in the d_z² orbital, i.e. (eg)⁴(a₁g)²(b₁g)²(b₂g)¹. This pattern is typical for copper complexes with square pyramidal environments.

The electronic spectra of complexes 1a and b were recorded for 1 × 10⁻⁴ molar solutions of the metal complexes in DMSO; they displayed peaks at 265 nm and 296 nm, attributed to π–π* and low energy n–π* ligand centered transitions, while in the visible region, a broad d–d transition band at 618 nm validated the pentacoordinated geometry around the central Cu(II) metal in both the complexes.³⁶

Solution stability studies

The stabilities of the complexes have been confirmed by UV-vis spectroscopy. UV-visible absorption spectra of complexes 1a and b were recorded in DMSO as well as in water at different time intervals (0 h, 3 h, 8 h and 24 h). The results revealed no appreciable change either in the intensity or the position of the absorption bands. Moreover, no significant changes in the absorption spectra were observed, even after a period of 24 h, indicating the stability of their geometries and coordinating spheres (Fig. 2(a) and (b)). These results are quite consistent with the stability of analogous complexes reported in the literature.^37,38

Fig. 2 (a) UV-vis absorption spectra of (a) complex 1a and (b) complex 1b in DMSO at different time intervals (0 h, 3 h, 8 h and 24 h). (b) UV-vis absorption spectra of (a) complex 1a and (b) complex 1b in water at different time intervals (0 h, 3 h, 8 h and 24 h).

In vitro binding studies of complexes with t-RNA

The binding propensity of complexes 1a and b with t-RNA was studied using electronic absorption spectroscopy. Upon addition of increasing amounts of t-RNA (0.00 to 5.0 × 10⁻⁵ M) to fixed concentrations of complexes 1a and b (2.0 × 10⁻⁵ M), a ‘hypochromic effect’ was observed, with a slightly bathochromic shift of 4 to 7 nm in the intraligand band at 265 nm (Fig. 3). Hypochromism is a typical characteristic of intercalation because of the π–π stacking between the base pairs of t-RNA and complexes 1a and b. Furthermore, in the intercalative mode of binding, the π* orbital of the intercalated ligand can couple with the π orbital of the nucleic acid base pair; this decreases the π–π* transition energy, resulting in bathochromism, while the partial electron filling of the coupling π orbital decreases the transition probabilities, concomitantly resulting in hypochromism.

Fig. 3 Absorption spectra of (a) complex 1a and (b) complex 1b in the absence and presence of increasing amounts of yeast t-RNA in Tris–HCl buffer at pH 7.2. Insets: plots of [RNA]/ε_a − ε_f (M² cm) vs. [RNA] for the titration with complex 1a and b, , experimental data points, full lines, and linear fitting of the data. [t-RNA] = 0.0 to 5.0 × 10⁻⁵ M, [complex 1] = 1.67 × 10⁻⁴ M. The arrows indicate the change in intensity with increasing [t-RNA].

Although there are subtle differences in the binding potentials of the different enantiomers, we can nevertheless discriminate between the two enantiomeric forms by observing their spectral titration profiles. The differences in binding of the enantiomeric L- and D-forms are quite discernible due to the alternate conformation, as there is greater hypochromism in the case of L-enantiomer 1a of 36% with a red shift of 6 nm, while D-enantiomer 1b exhibited hypochromism of 31% with a red shift of 4 nm.

In order to quantitatively compare the binding strengths of complexes 1a and b with the t-RNA intrinsic binding constant, K_b values were calculated using the Wolfe–Shimmer eqn (1).³⁹

[RNA]/ε_a − ε_f = [RNA]/ε_b − ε_f + 1/K_b|ε_b − ε_f| (1)

where [RNA] is the t-RNA concentration and ε_a, ε_f and ε_b are the apparent (A_abs/[Cu(II) complex]), free and bound complex extinction coefficients, respectively. In a plot of [RNA]/(ε_a − ε_f) vs. [RNA] with a slope of 1/(ε_b − ε_f) and an intercept of 1/[K_b(ε_b − ε_f)], K_b values were obtained by the ratio of the slope to the intercept. The intrinsic binding constant K_b values were found to be 2.6 (±0.08) × 10⁵ M⁻¹ and 1.5 (±0.08) × 10⁵ M⁻¹ (Table 1) for complexes 1a and 1b, respectively.

Table 1 The binding constant (K_b) values of complexes 1a and b with yeast t-RNA (mean standard deviation)

Yeast t-RNA	λmax (nm)	Kb (M^−1)	% hyperchromism
Complex 1a	265	2.6 (±0.08) × 10^5	36
Complex 1b	265	1.5 (±0.08) × 10^5	31

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Because DNA is a primary pharmacological target for many metal-based drugs, the comparative binding profiles for complexes 1a and b with CT-DNA were also studied; the results revealed a higher order of binding constants with t-RNA compared to CT-DNA (intrinsic binding constant, K_b = 2.73 (±0.05) × 10⁴ M⁻¹ and 1.97 (±0.05) × 10⁴ M⁻¹). H. Xu et al.⁴⁰ carried out similar binding studies on a ruthenium(II) polypyridyl complex; these observations revealed that the complex exhibited better potential for enantioselective binding to yeast t-RNA than to CT-DNA. This is quantitatively validated by the values of K_b, which were found to be 2.21 × 10⁶ M⁻¹ and 8.53 × 10⁵ M⁻¹ for yeast t-RNA and CT-DNA, respectively. This experimental finding demonstrates strong binding propensity and specificity for yeast t-RNA. The rationale for the specific RNA binding of the complexes could be (i) the highly exposed base pairs of RNA, which are easy to access and (ii) the bulge region of yeast t-RNA (which is absent in CT-DNA), which provides greater overlap in RNA, thereby enhancing the binding affinity of the complexes.⁴¹

The binding of complexes 1a and b with t-RNA was further validated by another, more sensitive fluorescence spectroscopy method. Upon excitation at a wavelength of 260 nm (λ_ex), the fluorescence spectra of complexes 1a and b exhibited appreciable fluorescence at ca. 380 nm (λ_em) and demonstrated a further increase upon concomitant addition of t-RNA (0–4.00 × 10⁻⁵ M) without any apparent shift in the emission maxima (Fig. S5†). This enhancement in the fluorescence intensity could be largely attributed to the penetration of complexes into the hydrophobic environment of the interior t-RNA helix, thereby reducing its accessibility to solvent molecules as well as restricting the mobility of the complexes at the binding site, which causes reduction of the vibrational modes and leads to higher emission intensity.⁴²

The binding strengths of complexes 1a and b with t-RNA were ascertained by the binding constant (K) value derived from a Scatchard equation⁴³ and were found to be 4.6 × 10⁵ M⁻¹ and 3.2 × 10⁵ M⁻¹, respectively. On the basis of the K values, we can evaluate the extent and mode of binding of the different enantiomers to underline the enantiospecific discrimination.

Competitive binding experiments were carried out on the EB–t-RNA system by varying the concentration of the complex to obtain further insight into the binding affinities of the complex–yeast t-RNA interactions. EB is a planar cationic dye that emits intense fluorescence in the presence of t-RNA at ca. 600 nm (λ_em) due to its strong intercalation between the adjacent base pairs. The enhanced fluorescence was quenched upon addition of the second molecule by replacing the bound EB or accepting the excited electron from EB.⁴⁴

It was observed that upon concomitant addition of complexes 1a and b to the EB–t-RNA system [complex]/[t-RNA] = 1 : 1 to 8 : 1, the emission intensity decreased progressively, indicative of competition between EB and the complexes towards yeast t-RNA binding (Fig. S6†). The extent of reduction of the emission intensity gives a measure of the binding propensity of the complex to t-RNA. Quantitative assessment of the magnitude of interaction was further ascertained by the classical Stern–Volmer equation as

I₀/I = 1 + K_svr (2)

where I₀ and I represent the fluorescence intensities in the absence and presence of complex, respectively, and r is the concentration ratio of complex 1 to t-RNA. K_sv is used to evaluate the quenching efficiency and is obtained as the slope of I₀/I vs. r. The calculated Stern–Volmer constants, K_sv, were found to be 1.62 and 1.0 for complexes 1a and b, respectively. The calculated K_b, K and K_sv values verified that the L-enantiomeric form of the complex binds more strongly to t-RNA than the D-enantiomeric form.

Circular dichroic titrations were performed as a function of complex concentration to monitor the conformational changes in t-RNA and the binding modes of the complexes upon interaction with t-RNA. The CD spectrum of free t-RNA displayed four major peaks at 208 and 240 nm (negative) and 221 and 270 nm (positive), consistent with the conformation of t-RNA. Upon addition of complex 1a (r = [complex]/[t-RNA] = 1), a lower decrease in the molar ellipticity band at 208 and 270 nm was observed, while the positive band at 227 and the negative band at 240 nm demonstrated relatively greater amplification. Complex 1b presented an inverse CD spectrum in which the intensity of the positive band at 227 significantly increased, while the intensity of the other bands decreased (Fig. S7†). The enhancement in intensity of the band at 270 nm in the spectra of the complex–t-RNA systems along with the major intensity changes of the bands at 210 and 227 nm could be attributed to the formation of t-RNA aggregates upon complexation.^45,46 From these CD assays, we could virtually discriminate the enantiopreferential binding of t-RNA with the L- and D-forms of the complexes; to the best of our knowledge, this is the first report of an enantiomeric binding profile of RNA in the literature.

The electrochemical profiles of complexes 1a and b were studied in H₂O/DMSO (95 : 5) at room temperature at a scan rate of 0.2 V s⁻¹ by cyclic voltammetry in the potential range from 0.8 to −1.0 V. Both complexes displayed well-defined cathodic waves; however, the corresponding anodic waves were drawn out, revealing that the very unstable Cu^I species are not discernible with the time scale of the CV experiments.⁴⁷ The cyclic voltammogram of 1a exhibited a quasi-reversible one-e⁻ redox process involving the Cu^II/Cu^I couple with a cathodic peak at E_pc = −0.48 V and an oxidation peak at E_pa = −0.12 V (Fig. S8(a)†). For this couple, the difference between the cathodic and anodic peak potential ΔE_p and the ratio of the cathodic and anodic peak currents I_pa/I_pc are −0.36 and 1.0, respectively. The formal electrode potential E_1/2, taken as an average of E_pc and E_pa, was −0.30 V in the absence of t-RNA. The CV of 1b (Fig. S8(b)†) featured reduction of the Cu^II/Cu^I form at a cathodic peak potential of E_pc = −0.45 V. Reoxidation of the complex occurred at E_pa = −0.11 V. The difference between the anodic and cathodic peak potential ΔE_p = −0.34 V and the I_pa/I_pc ratio = 0.77, which indicated a quasi-reversible redox process.

Addition of yeast t-RNA to the complexes resulted in significant reduction in the cathodic and anodic peak currents due to the slow diffusion of an equilibrium mixture of the free and RNA-bound complexes to the electrode surface. The observed shifts in the E_1/2 values suggested that the Cu^II and Cu^I forms of complexes 1a and b were bound to the t-RNA. However, the shifts observed in the ΔE_p, E_1/2 and I_pa/I_pc values were greater for complex 1a than for complex 1b, which further validated the greater binding affinity of the L-isomer of the complex towards t-RNA.

Morphological studies

Structural studies of RNA condensate with metal complexes can help to map/identify the specific interactions of enantiomeric drug entities to RNA; RNA possesses highly structured active sites that can dictate the formation of different shape particles due to its thermodynamic stability, base stacking capabilities and tertiary interactions.⁴⁸ Literature reports reveal that the morphology of the condensate mainly depends on the ion type, solution properties (i.e. ionic strength and solvent polarity), nature of the condensing agent (i.e. charge density) and surface of the substrate.⁴⁹ In this study, RNA condensate was achieved by evaporating an equimolar mixture of complex 1a and t-RNA or complex 1b and t-RNA, respectively, under neutral conditions in aqueous Tris–HCl buffer (50 mM NaCl, pH 7.2). Scanning electron microscopy was employed to analyze the morphological changes of the condensate in comparison to complexes 1a and b. Micrographs of complexes 1a and b are shown in Fig. 4(a) and (b) and indicate that the chemotherapeutic candidates 1a and b mostly display crystalline and irregular morphologies with rectangular, almost cubic shapes of various sizes. However, the SEM micrographs of complex 1a with t-RNA present different structural features, as evidenced by the formation of hollow tubes and some concrete-like structures, indicating the condensation of RNA molecules into a compact, massive structure. These morphological changes were observed in complexes 1a and b; however, 1a showed more prominent morphological changes with t-RNA compared to 1b.

Fig. 4 (a) SEM images (a and b) of complex 1a nanoparticles, showing the surface morphology; (c and d) condensation of t-RNA incubated in the presence of complex 1a after 24 h; and (e) a portion of the condensate image showing the formation of hollow tubes. (b) SEM images (a and b) of complex 1b nanoparticles, showing the surface morphology, and (c and d) condensation of t-RNA incubated in the presence of complex 1b after 24 h.

These results suggested the condensation of t-RNA by cationic molecules and that a major factor governing the condensation of t-RNA is specific interaction of the complexes with the base pairs of t-RNA, which correlates well with the in vitro t-RNA binding studies.

Molecular docking studies

Docking experiments were carried out with t-RNA (PDB ID: 6TNA) in order to predict the preferred orientation of the complexes inside the RNA helix and to substantiate the spectroscopic results. Yeast t-RNA has well-defined 3D structures, including regions such as the D arm, acceptor stem, T arm, ψ loop and anticodon arm. The most energetically favorable conformations of the docked pose of complexes 1a and b revealed that both the L- and D-complexes fitted snugly into the RNA intercalation site, i.e. inserted into the active pocket of the anticodon arm in close proximity of C-40, A-35, A-36, A-38, G-43, U-41 and C-28.⁵⁰ The optimal binding conformations of the complexes allow them to form intermolecular H-bonds, considered on the basis of the 3 Å distance between the O atoms of complex (H-bonding acceptor group) and the H (H-bonding donor group) atom of G37, such that G37:O2′⋯O1 of complex 1a = 2.39 Å and 1b = 2.37 Å, respectively. In addition to these forces, contributions from van der Waals forces and hydrophobic interactions play a major role in the stronger binding of complexes to yeast t-RNA (Fig. 5 and 6).

Fig. 5 Molecular docked model of complex 1a docked into the active pocket of the anticodon arm of yeast t-RNA (PDB ID: 6TNA) along with the possible hydrogen bonding interactions. Hydrogen atoms have been omitted for clarity.

Fig. 6 Molecular docked model of complex 1b docked into the active pocket of the anticodon arm of yeast t-RNA (PDB ID: 6TNA) along with the possible hydrogen bonding interactions. Hydrogen atoms have been omitted for clarity.

The resulting binding energies calculated for the best docked pose of complexes 1a and b with the yeast t-RNA target were found to be −275 and −256 kJ mol⁻¹, respectively. The more negative binding energy value of complex 1a signifies the greater binding propensity of the L-form of the metal complex, which correlates well with the results of the in vitro t-RNA binding studies.

Antibacterial activity

Complexes 1a and b were screened for their in vitro antibacterial activity against Gram-positive Bacillus subtilis [MTCC 121] and Staphylococcus aureus [NRRLB 767] as well as Gram-negative Escherichia coli [K12] and Pseudomonas aeruginosa; the relevant data are presented in Table 2. The antibacterial activities of complexes 1a and b were evaluated by measuring the zones of inhibition; both the complexes exhibited uncertain degrees of inhibitory effects on the growth of the test organisms, as the molecules contain the 1,10-phen moiety, which is capable of killing many bacterial strains.²⁹ Complex 1a shows very high activity against B. subtilis, with zone diameters of 27 and 33 mm, and against E. coli, with zone diameters of 20 and 27 mm at 100 μg and 200 μg concentrations, respectively. The screening results indicated that complex 1a exhibited higher activity than complex 1b against all tested bacteria (Fig. 7 and 8).

Table 2 Antibacterial data for complexes 1a and 1b

Compound	Zone of inhibition (mm)
	B. subtilis		S. aureus		E. coli		P. aeruginosa
	100 μg	200 μg	100 μg	200 μg	100 μg	200 μg	100 μg	200 μg
1a	27	33	11	17	20	27	—	10
1b	22	27	10	14	16	23	—	—

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Fig. 7 Antibacterial activity of complex 1a (A) and b (B) against E. coli and B. subtilis at 100 μg (A1 and B1) and 200 μg (A2 and B2) concentrations.

Fig. 8 The antimicrobial activities of complexes 1a and b at 100 μg concentration.

Conclusions

Much attention has been given to the rationale of the development of new, efficacious antibiotics owing to bacterial resistance to existing and currently used antibiotics; new chiral conjugate phenylalanine-(1,10-phen)–Cu(II) complexes, 1a and 1b, were synthesized for antibacterial treatments and were thoroughly characterized by spectroscopic techniques, viz., FT-IR, EPR, and ESI-MS, and by elemental analysis. Structure elucidation of both 1a and b was performed by single X-ray diffraction. The complexes are mononuclear and the asymmetric units of complexes 1a and b contain one NO₃⁻ ion, one water molecule and a Cu(II) ion; they possess a distorted square pyramidal geometry in which L- and D-phenylalanine, respectively, coordinate through the amino nitrogen and deprotonated carboxylato oxygen atom to form a five-membered ring with an “envelope” conformation. The complexes satisfy one prerequisite condition for the design of novel therapeutics in that they are readily soluble in water; therefore, they show superior ability to cross biological membranes. RNA, particularly miRNA, plays an indispensable role in various biological processes, such as cell proliferation, differentiation and apoptosis; therefore, RNA has been identified as a target for anti-infectives and cancer therapeutics. In consideration of this, in vitro interaction studies of complexes 1a and b with t-RNA employing UV-vis, fluorescence, circular dichroism and cyclic voltammetry techniques were carried out. We were interested to study the differential disposition of enantiomers 1a and b towards their biological target t-RNA on the basis of intrinsic binding constant values and the site preference of RNA for the L-enantiomer over the D-enantiomer. Furthermore, SEM analysis was performed to assess the surface morphological changes of complexes 1a and b; the images revealed the formation of hollow tubes and concrete-like structures with RNA condensate, which was more pronounced in the SEM micrographs of the L-enantiomer of the complex 1a condensate. The antibacterial activity of the complexes was also evaluated and displayed high antibacterial activity of complex 1a compared to 1b. Additionally, molecular docking studies of the complexes showed an intercalative mode of binding and a relatively high binding energy of the L-enantiomer, 1a, which correlated well with other spectroscopic techniques.

Experimental

Materials and instrumentation

L- and D-phenylalanine, 1–10 phenanthroline and copper(II) nitrate were purchased from Sigma Aldrich. All reagents were of the best commercial grade and were used without further purification. Elemental analysis was carried out on a Carlo Erba Analyser Model 1106. Molar conductance was measured at room temperature on a Eutech CON 510 electronic conductivity bridge. Fourier-transform infrared (FT-IR) spectra were recorded on Interspec 2020 and Spectrum Two (Perkin Elmer) FT-IR spectrometers. The EPR spectra of the copper complexes were acquired on a Varian E 112 ESR spectrometer using X-band frequency (9.5 GHz) at liquid nitrogen temperature. ESI-MS spectra were recorded on a Micromass Quattro II triple quadrupole mass spectrometer. Electronic spectra were recorded on a Perkin-Elmer Lambda 25 using cuvettes of 1 cm path length, and the data were reported in λ_max/nm. Emission spectra were acquired on a Shimadzu RF-5301PC spectrofluorophotometer. All the experiments involving the interaction of the complex with CT-DNA and yeast t-RNA were carried out in aerated buffer (5.0 mM Tris–HCl, 50.0 mM NaCl, pH = 7.3). The concentrations per base pairs for both DNA and RNA were determined spectrophotometrically by assuming the ε_{260 nm} values to be 6600 and 7700 M⁻¹ cm⁻¹, respectively. CD spectra were measured on a Jasco J-815-CD spectropolarimeter at room temperature. Cyclic voltammetry was carried out using a CH instrument electrochemical analyzer. High purity H₂O and DMSO (95 : 5) were employed for the cyclic voltammetry studies using 0.4 M KNO₃ as the supporting electrolyte. A three electrode configuration was used, comprising a Pt disk working electrode, a Pt wire counter electrode and Ag/AgCl as the reference electrode.

Synthesis of complexes 1a and b

To a methanolic solution of L-/D-phenylalanine (0.165 g, 1 mmol) deprotonated by NaOH (1 mmol) was added a methanolic solution of copper(II) nitrate trihydrate (0.241 g, 1 mmol). After 0.5 h of stirring, a methanolic solution of 1,10-phenanthroline (0.198 g, 1 mmol) in a stoichiometric ratio of 1 : 1 was added to this reaction mixture, affording a dark blue clear solution. Dark blue crystals of complexes 1a and b suitable for single X-ray crystallography were obtained by slow evaporation of the reaction mixture.

1a. Yield: (66%), mp 132 °C; anal. calc. for C₂₁H₂₂CuN₄O₇ (%): calc. C, 49.85; H, 4.38, N, 11.07. Found: C, 49.32; H, 4.41, and N, 11.09. UV-vis (1 × 10⁻³ M, DMSO, λ_max nm): 265, 296, 618 nm. FTIR (KBr pellet, ν_max/cm⁻¹) 3469, 3239–3133, 1620, 1336, 912, 858, 765, 535, 428. ESI-MS (DMSO) (+) (m/z): 487, 425, 407, 261 and 243. CCDC: 1405339.†

1b. Yield: (62%), mp 134 °C; anal. calc. for C₂₁H₂₂CuN₄O₇ (%): calc. C, 49.85; H, 4.38, N, 11.07. Found: C, 49.17; H, 4.13, and N, 11.13. UV-vis (1 × 10⁻³ M, DMSO, λ_max nm): 265, 296, 618 nm. FT-IR (KBr pellet, ν_max/cm⁻¹) 3467, 3238–3131, 1620, 1335, 912, 858, 765, 535, 428. ESI-MS (DMSO) (+) (m/z): 487, 425, 407, 261 and 243. CCDC: 1472957.†

Description of X-ray crystal structure

Suitable X-ray quality crystals of complexes 1a and 1b were obtained after slow evaporation of the in a DCM : MeOH solution (7 : 3) at room temperature. Single crystal X-ray structural studies of the complexes were performed on a CCD Oxford Diffraction X caliber Saphir 3 diffractometer employing graphite-monochromated Mo-Kα radiation generated from a fine-focus sealed tube (λ = 0.71073 Å) at 140(2) K. The data collection strategy was evaluated using Crys Alis Pro CCD software. The collections of data were observed by the standard ω scan techniques and were scaled and reduced using Crys Alis Pro RED software. The structure was solved by direct methods using SIR-97 (ref. 51) and refined by least-squares methods on F² using SHELXL-97.⁵² The positions of all atoms were obtained by direct methods. Anisotropic thermal parameters were assigned to all non-hydrogen atoms, and the remaining hydrogen atoms were placed in geometrically constrained positions and refined as riding atoms with a common fixed isotropic thermal parameter. The drawing of the complex was realized with PLATON.⁵³ A summary of selected crystallographic data for complexes 1a and 1b is given in Table 3.

Table 3 Crystallographic data for the complexes

a GoF is defined as {∑[w(F0^2 − Fc^2)]/(n − p)}^1/2, where n is the number of data and p is the number of parameters.b R = {∑||F0| − |Fc||/∑|F0|, wR^2 = {∑w(F0^2 − Fc^2)^2/∑w(F0^2)^2}^1/2.
Complex	1a	1b
Formula	C21H22CuN4O7	C21H22CuN4O7
Fw (g mol^−1)	505.96	505.97
Crystal system	Monoclinic	Monoclinic
Space group	P21	P21
a (Å)	5.74690(10)	5.728(5)
b (Å)	20.6365(2)	20.587(5)
c (Å)	9.28010(10)	9.252(5)
α (deg)	90	90
β (deg)	98.2040(10)	98.308(5)
γ (deg)	90	90
U (Å^3)	1089.32(2)	1079.6(11)
Z	2	2
ρcalc (mg cm^−3)	1.543	1.557
μ (mm^−1)	1.871	1.064
F(000)	522	522
Flack	0.027(11)	−0.016(13)
Temp (K)	150(2) K	150(2) K
Measured reflns	9949	12 893
Indep reflns	3741	3579
GOF^a	1.080	1.056
R^b[I > 2σ(I)]	0.0253	0.0301
wR2^b (all data)	0.0775	0.0684

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In vitro binding experiments

DNA/RNA binding experiments were performed in Tris–HCl/NaCl (5 : 50 mM) buffer at pH 7.2 and included absorption spectral traces, emission spectroscopy, viscosity measurements and CD, conforming to the standard methods and practices previously adopted by our laboratory.^54–56 While measuring the absorption spectra, equal amounts of CT-DNA and t-RNA were added to both the compound solution and the reference solution to eliminate the absorbance of the t-RNA itself, and the absorbance of the Tris buffer was subtracted through base line correction.

Molecular docking studies

Molecular docking studies were performed using HEX 8.0 software,⁵⁷ which is an interactive molecular graphics program for calculating and displaying feasible docking modes of enzymes and DNA molecules. The structures of complexes 1a and 1b were converted into PDB format from mol format by OPENBABEL (http://www.vcclab.org/lab/babel/). The crystal structure of RNA (PDB ID: 6TNA) was downloaded from the protein data bank (http://www.rcsb.org./pdb). Visualization of the docked pose was performed using CHIMERA (http://www.cgl.ucsf.edu/chimera), PyMol (http://pymol.sourceforge.net/) and the Discovery Studio molecular graphics program.

Antibacterial assay

Antibacterial screening of the complexes was performed by the Agar well diffusion method of Perez et al., also described earlier by Ahmad et al. 0.1 ml of diluted inoculum (10⁵ cfu ml⁻¹) of the test organism was spread on nutrient agar (NA)/SD plates. Wells of 8 mm diameter were punched into the agar medium and were filled with test solutions of different concentrations (100 mg ml⁻¹, 200 mg ml⁻¹ and 400 mg ml⁻¹), solvent blank (DMSO) and an antibacterial drug (nitrofurantoin, 100 mg ml⁻¹). The plates were incubated for 18 h at 37 °C for the test bacteria. The antibacterial activity was evaluated by measuring the zone of inhibition against the test organism.

Acknowledgements

The authors are thankful to SAIF, CIL, Panjab University, Chandigarh, for ESI-Mass and elemental analysis facilities. The financial support from the DST-PURSE programme and DRS-1 (SAP) from UGC, New Delhi is gratefully acknowledged. The authors are grateful to USIF, Aligarh Muslim University for providing SEM facilities. We are grateful to Dr Iqbal Ahmad and Mohammad Shavez Khan, Department of Agricultural Microbiology, Faculty of Agricultural Sciences, Aligarh Muslim University, Aligarh for carrying out the antibacterial studies. The author (S. Sharma) expresses his gratitude to the University Grants Commission (UGC), New Delhi, for a BSR Fellowship.

References

1. P. Nagababu, J. N. L. Latha and S. Satyanarayanana, Chem. Biodiversity, 2006, 3, 1219–1229.
2. D. M. Kolpashchikov, Chem. Rev., 2010, 110, 4709–4723.
3. D. Li, S. Song and C. Fan, Acc. Chem. Res., 2010, 43, 631–641.
4. M. Gaglione, G. Milano, A. Chambery, L. Moggio, A. Romanelli and A. Messere, Mol. BioSyst., 2011, 7, 2490–2499.
5. J.-H. Li, J.-T. Wang, Z.-W. Mao and L.-N. Ji, Inorg. Chem. Commun., 2008, 11, 865–868.
6. D. Desbouis, I. P. Troitsky, M. J. Belousoff, L. Spiccia and B. Graham, Coord. Chem. Rev., 2012, 256, 897–937.
7. J. Gallego and G. Varani, Acc. Chem. Res., 2001, 34, 836–843.
8. N. Foloppe, N. Matassova and F. Aboul-ela, Drug Discovery Today, 2006, 11, 1019–1027.
9. R. W.-Y. Sun, D.-L. Ma, E. L.-M. Wong and C.-M. Che, Dalton Trans., 2007, 4884–4892.
10. C.-Y. Hong, X. Chen, J. Li, J.-H. Chen, G. Chen and H.-H. Yang, Chem. Commun., 2014, 50, 3292–3295.
11. D. M. Pereira, P. M. Rodrigues, P. M. Borralho and C. M. P. Rodrigues, Drug Discovery Today, 2013, 18, 282–288.
12. S. Sassen, E. A. Miska and C. Caldas, Virchows Arch., 2008, 452, 1–10.
13. C. S. Chow and F. M. Bogdan, Chem. Rev., 1997, 97, 1489–1513.
14. L. Guan and M. D. Disney, ACS Chem. Biol., 2012, 7, 73–86.
15. G. Bischoff and S. Hoffmann, Curr. Med. Chem., 2002, 9, 312–348.
16. K. B. Turner, A. S. Kohlway, N. A. Hagan and D. Fabris, Biopolymers, 2008, 91, 283–296.
17. N. Delihas, S. E. Rokita and P. Zheng, Nat. Biotechnol., 1997, 15, 751–753.
18. K. F. Blount and R. R. Breaker, Nat. Biotechnol., 2006, 24, 1558–1564.
19. A. J. Alanis, Arch. Med. Res., 2005, 36, 697–705.
20. S. Ambika, S. Arunachalam, R. Arun and K. Premkumar, RSC Adv., 2013, 3, 16456–16468.
21. M. Mudasir, N. Yoshioka and H. Inoue, J. Inorg. Biochem., 2008, 102, 1638–1643.
22. N. Thanki, J. M. Thornton and J. M. Goodfellow, J. Mol. Biol., 1988, 202, 637–657.
23. V. Uma, M. Kanthimathi, T. Weyhermuller and B. U. Nair, J. Inorg. Biochem., 2005, 99, 2299–2307.
24. Q. Zhou and P. Yang, Inorg. Chim. Acta, 2006, 359, 1200–1206.
25. Y. Li, Y. Wu, J. Zhao and P. Yang, J. Inorg. Biochem., 2007, 101, 283–290.
26. F. P. Dwyer, I. K. Reid, A. Shulman, G. M. Laycock and S. Dixson, Aust. J. Exp. Biol. Med. Sci., 1969, 47, 203–218.
27. P. R. Reddy, S. Rajeshwar and B. Satyanarayana, J. Photochem. Photobiol., B, 2016, 160, 217–224.
28. T. Hara, H. Matsuzaki, T. Nakamura, E. Yoshida, T. Ohkubo, H. Maruyama, C. Yamamoto, S. Saito and T. Kaji, Fundam. Toxicol. Sci., 2016, 3, 109–113.
29. N. S. Ng, P. Leverett, D. E. Hibbs, Q. Yang, J. C. Bulanadi, M. J. Wua and J. R. Aldrich-Wright, Dalton Trans., 2013, 42, 3196–3209.
30. I. Yousuf, F. Arjmand, S. Tabassum, L. Toupet, R. A. Khan and M. A. Siddiqui, Dalton Trans., 2015, 44, 10330–10342.
31. F. Arjmand, I. Yousuf, M. Afzal and L. Toupet, Inorg. Chim. Acta, 2014, 421, 26–37.
32. F. Arjmand, I. Yousuf, T. b. Hadda and L. Toupet, Eur. J. Med. Chem., 2014, 81, 76–88.
33. P. S. Subramanian, E. Suresh, P. Dastidar, S. Waghmode and D. Srinivas, Inorg. Chem., 2001, 40, 4293–4301.
34. F. Arjmand, I. Yousuf, Y. Zaidi and L. Toupet, RSC Adv., 2015, 5, 16250–16264.
35. M. Chauhan, K. Banerjee and F. Arjmand, Inorg. Chem., 2007, 46, 3072–3082.
36. S. Dhar, M. Nethaji and A. R. Chakravarty, Inorg. Chem., 2006, 45, 11043–11050.
37. (a) M. A. Neelakantana, F. Rusalraj, J. Dharmaraja, S. Johnsonraja, T. Jeyakumar and M. Sankaranarayana Pillai, Spectrochim. Acta, Part A, 2008, 71, 1599–1609; (b) F. Arjmand, M. Muddassir and R. H. Khan, Eur. J. Med. Chem., 2010, 45, 3549–3557.
38. (a) D. inci, R. Aydın, Ö. Vatan, D. Yılmaz, H. Mutlu Gençkal, Y. Zorlu and T. Cavaş, Spectrochim. Acta, Part A, 2015, 145, 313–324; (b) A. K. Patra, M. Nethaji and A. R. Chakravarty, Dalton Trans., 2005, 2798–3280.
39. A. Wolfe, G. H. Shimer Jr and T. Meehan, Biochemistry, 1987, 26, 6392–6396.
40. H. Xu, Y. Liang, P. Zhang, F. Du, B.-R. Zhou, J. Wu, J.-H. Liu, Z.-G. Liu and L.-N. Ji, JBIC, J. Biol. Inorg. Chem., 2005, 10, 529–538.
41. X.-L. Liang and L.-F. Tan, Aust. J. Chem., 2010, 63, 1453–1461.
42. K. A. Meadows, F. Liu, J. Sou, B. P. Hudson and D. R. McMillin, Inorg. Chem., 1993, 32, 2919–2923.
43. F. Cui, Y. Yan, Q. Zhang, J. Du, X. Yao, G. Qu and Y. Lu, Carbohydr. Res., 2009, 344, 642–647.
44. J.-B. LePecq and C. Paoletti, J. Mol. Biol., 1967, 27, 87–106.
45. (a) M. Vorlickova, Biophys. J., 1995, 69, 2033–2043; (b) J. Kypr and M. Vorlickova, Biopolymers, 2002, 67, 275–277.
46. E. Froehlich, E. J. S. Mandeville, D. Arnold, L. Kreplak and H. A. Tajmir-Riah, Biomacromolecules, 2012, 13, 282–287.
47. (a) A. Raja, V. Rajendiran, P. U. Maheswari, R. Balamurugan, C. A. Kilner, M. A. Halcrow and M. Palaniandavar, J. Inorg. Biochem., 2005, 99, 1717–1732; (b) B. C. Baguley and M. LeBert, Biochemistry, 1984, 23, 937–943.
48. L. A. Gugliotti, D. L. Feldheim and B. E. Eaton, J. Am. Chem. Soc., 2005, 127, 17814–17818.
49. X. D. Liu, H. Y. Diao and N. Nishi, Chem. Soc. Rev., 2008, 37, 2745–2757.
50. D. Agudelo, P. Bourassa, M. Beauregard, G. Berube and H.-A. Tajmir-Riahi, PLoS One, 2013, 8, e69248.
51. A. Altomare, M. C. Burla, M. Camalli, G. L. Cascarano, C. Giocavazzo, A. Guagliardi, A. G. C. Moliterni, G. Polidori and S. Spagna, J. Appl. Crystallogr., 1999, 32, 115–119.
52. G. M. Sheldrick, SHELX-97, Program for Crystal Structure refinement, University of Göttingen, Germany, 1997.
53. A. L. Spek, PLATON Procedure, A multipurpose Crystallographic Tool, Utrecht University, Utrecht, The Netherlands, 1998.
54. J. R. Lakowicz and G. Webber, Biochemistry, 1973, 12, 4161–4170.
55. A. Wolfe, G. H. Shimer Jr and T. Meehan, Biochemistry, 1987, 26, 6392–6396.
56. F. Arjmand, M. Muddassir, Y. Zaidi and D. Ray, MedChemComm, 2011, 14, 394–405.
57. (a) D. Mustard and D. W. Ritchie, Proteins: Struct., Funct., Bioinf., 2005, 60, 269–274; (b) G. Macindoe, L. Mavridis, V. Venkatraman, M. D. Devignes and D. W. Ritchie, Nucleic Acids Res., 2010, 38, 445–449.

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Footnote

† Electronic supplementary information (ESI) available. CCDC 1405339 and 1472957. For ESI and crystallographic data in CIF or other electronic format see DOI: 10.1039/c6ra14503e

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