DNA binding, molecular docking and apoptotic inducing activity of nickel(II), copper(II) and zinc(II) complexes of pyridine-based tetrazolo[1,5-a]pyrimidine ligands

Abstract

Six mononuclear copper(II), nickel(II) and zinc(II) complexes, [ML¹Cl₂] (1–3) and [M(L²)₂Cl₂] (4–6), of two biologically active tetrazolo[1,5-a]pyrimidine core ligands, ethyl 5-methyl-7-pyridine-2-yl-4,7-dihydrotetrazolo[1,5-a]pyrimidine-6-carboxylate (L¹) and ethyl-5-methyl-7-pyridine-4-yl-4,7-dihydrotetrazolo[1,5-a]pyrimidine-6-carboxylate (L²), have been synthesized and characterized. The molecular structure of the ligands (L^1&2) and complex 6 were determined by single crystal X-ray diffraction. The X-ray crystal structure of 6 confirms that it has a distorted tetrahedral structure with a ZnN₂Cl₂ coordination environment. All of the complexes exhibit an unusually strong luminescence at room temperature. Electroparamagnetic resonance spectra of copper(II) complexes (2 and 5) show four lines, characteristic of square planar geometry, with nuclear hyperfine spin 3/2. DNA binding studies of complexes with calf-thymus DNA suggest that complexes 2 and 5 bind in the grooves of the DNA. These results were further supported by molecular docking studies. In vitro cytotoxic activities of the ligands (L^1&2) and complexes (1–6) against human cancer cell lines such as lung (A549), cervical (HeLa), colon (HCT-15) and a non-cancer human embryonic kidney cell line revealed that the complexes selectively inhibit the growth of cancer cells and are inactive against non-cancer cell lines, whereas the ligands were found to be inactive with both cancer and non-cancer cell lines. The IC₅₀ values of the complexes revealed that the copper(II) complexes (2 and 5) exhibit high cytotoxic activity against colon (HCT-15) cells when compared to the standard drug cisplatin. Furthermore, the live cell and fluorescent imaging of cancer cells show that complexes 2 and 5 induce cell death through apoptosis.

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Introduction

A great deal of attention has been devoted towards the synthesis of functional compounds containing polyazole rings, particularly tetrazoles and their derivatives. The synthesis of tetrazoles by traditional cycloaddition reactions requires the use of expensive and toxic metal–organic azides and suffers from severe water sensitivity, or uses hydrazoic acid, which is extremely toxic, volatile, explosive and needs a long reaction time at 100–125 °C.^1–3 A safe, convenient, and environmentally friendly synthetic route to synthesize 5-substituted 1H-tetrazoles using water as a solvent and zinc salts as a catalyst was achieved by Demko and Sharpless.⁴ Putting aside the mechanism of [2 + 3] cycloaddition and the in situ hydrothermal method, the tetrazole ligand has been shown to be able to participate in at least nine distinct types of coordination modes with metal ions in the construction of metal–organic frameworks.⁵ Tetrazole with a pyridyl functional group has attracted a growing amount of attention in recent years in coordination chemistry because of the aromaticity of the pyridyl group and the excellent coordination ability of the four nitrogen atoms of the tetrazole ring, which may act as either a multidentate or a bridging block in supramolecular assemblies.^6,7

Nowadays considerable attention has also been diverted towards the synthesis of pyrimidines and related N-containing heterocyclic derivatives such as tetrazolopyrimidines. Pyrimidines and fused pyrimidines are an integral part of DNA and RNA, and thus, play an essential role in several biological and pharmacological agents such as antibiotics, antibacterial, antitumor and cardiovascular drugs and so on.^8,9 The formation of tetrazolopyrimidines derived from the condensation of β-diketones and β-keto esters with 5-aminotetrazole was first described by Bulow.¹⁰ The tetrazolopyrimidines have been reported to be used in the treatment of obesity, diabetes, atherosclerosis, hypertension, coronary heart disease, hypercholesterolemia, hyperlipidemia, thyroid cancer, hypothyroidism, depression, glaucoma, cardiac arrhythmias and congestive heart failure.¹¹ In addition to this, the interaction of tetrazolo metal complexes with DNA has been extensively studied because of their site specific binding properties and many applications in cancer therapy. These coordination compounds were suitable for DNA secondary structure probes, photocleavers and antitumor drugs.¹² Other than the medicinal applications, tetrazolate-based complexes (5-R-tetrazoles, R = alkyl, pyridyl, phenyl) with transition metals (i.e., Cu, Cd and Zn) also play an important role in various applications such as metal organic coordination polymers (MOCPs), electroluminescence materials (LEDs) and in nonlinear optics.^13–15 Although many investigators are interested in the synthesis of new tetrazolopyrimidine derivatives, there are few publications concerning the anti-tumor effects of complexes of tetrazolopyrimidines.

In this context, the coordination of metal ions with tetrazolopyrimidines may be expected to take place in various common coordination modes as shown in Scheme 1.¹⁶ To expand this area, we are interested in investigating the use of new tetrazolopyrimidine complexes as potential therapeutic agents, especially as anticancer agents. In this paper, we describe the synthesis, structural characterization, DNA-binding, molecular docking, cytotoxic and apoptotic activities of a new class of mononuclear copper(II), nickel(II) and zinc(II) complexes of two different tetrazolo[1,5-a]pyrimidine core ligands, ethyl 5-methyl-7-pyridine-2-yl-4,7-dihydrotetrazolo[1,5-a]pyrimidine-6-carboxylate (L¹) and ethyl 5-methyl-7-pyridine-4-yl-4,7-dihydrotetrazolo[1,5-a]pyrimidine-6-carboxylate (L²).

Scheme 1 Possible coordination modes of tetrazolo[1,5-a]pyrimidines with metal ions.

Results and discussion

Synthesis of ligands and complexes

The target ligands L^1&2 were synthesized by following the procedure described in the literature.¹⁷ All the complexes (1–6) were obtained by the metalation of ligands L^1&2 with CuCl₂, NiCl₂ or ZnCl₂ in methanol. The proposed structure of the complexes, as shown in Scheme 2, was based on spectroscopic investigation as well as elemental analysis and magnetic susceptibility determination.

Scheme 2 Schematic route for synthesis of ligands and metal(II) complexes.

Description of the crystal structures

The structure of the ligands L^1&2 and complex 6 were determined by X-ray analysis, and the Oak Ridge Thermal Ellipsoid plot (ORTEP) diagrams are given in Fig. 1–3. The crystal refinement detail is listed in Table S1.† Single crystals of L^1&2 were obtained by slow crystallization from ethanol. The ligand L¹ crystallized in the triclinic space group P1̄ and L² crystallized in the orthorhombic space group pbca with inversion centers that exhibit planar structure; a similar arrangement of the ligand has been reported in the literature.¹⁸ Selected bond lengths and bond angles are given in Tables S2 and S4.† The NH proton in the pyrimidine ring is involved in intermolecular hydrogen bonding with the nitrogen atom of the tetrazole ring of the neighboring molecule for the ligand L¹, and with the nitrogen atom of the neighboring pyridine ring for the ligand L² (Tables S3 and S5†). For the ligand L¹, the pyrimidine ring adopts a flattened-boat conformation, with atoms N5 and C2 deviating from the N1/C1/C3/C4 plane by −0.0899 (16) Å and −0.1472 (17) Å, respectively. The N1–N4/C1 and C6–C10/N6 planes form dihedral angles of 5.93 (0.06)° and 86.01 (0.06)°, respectively, with the N1/C1/C3/C4 plane. For the ligand L², the pyrimidine ring also adopts a flattened-boat conformation, with atoms N5 and C7 deviating from the N5/C6/C4/C5 plane by −0.029 (2) Å and −0.049 (2) Å, respectively. The N2–N5/C6 and C8–C12/N6 planes form dihedral angles of 1.45 (0.08)° and 80.97 (0.08)°, respectively, with the N5/C6/C4/C5 plane (Fig. S1–S4†).

Fig. 1 Single crystal X-ray structure of ligand L¹.

Fig. 2 Single crystal X-ray structure of ligand L².

Fig. 3 Single crystal X-ray structure of complex 6.

Small colourless rectangular crystals of complex 6 were obtained by slow evaporation of a methanolic solution of the complex. Selected bond angles and bond lengths are given in Table 1. The crystal system of complex 6 belongs to the monoclinic space group C2/c with an inversion centre, and the Zn(II) atoms exhibit a distorted tetrahedral (ZnN₂Cl₂) coordination environment, which is formed by the coordination of two Cl atoms, with a Cl–Zn–Cl^# bond angle of 119.03 (15)°. More interestingly the zinc atom coordinates to one nitrogen N₁ atom of both the pyridine rings of pyridyl tetrazole which acts as a monodentate ligand with a bond angle N(1)–Zn–Cl(1)^# of 109.41 (17)° and N(1)^#–Zn–Cl(1) of 109.40 (17)°.

Table 1 Selected bond lengths (Å) and bond angles (°) for complex 6^a

Bond lengths (Å)
a Symmetry transformations used to generate equivalent atoms: #1 − x + 2, y, −z + 1/2.
Cl(1)–Zn(1)	2.195(2)	C(4)–C(5)	1.395(9)
Zn(1)–N(1)^#	2.030(5)	C(5)–N(1)	1.335(7)
Zn(1)–Cl(1)^#	2.195(2)	C(6)–N(2)	1.460(7)
N(1)–Zn(1)	2.030(5)	C(7)–N(5)	1.309(8)
C(1)–N(1)	1.326(8)	C(8)–N(6)	1.358(8)
C(1)–C(2)	1.363(9)	C(11)–O(1)	1.206(8)
C(2)–C(3)	1.367(7)	C(11)–O(2)	1.320(8)
C(3)–C(4)	1.329(8)	C(12)–C(13)	1.526(19)
C(3)–C(6)	1.538(8)	C(12)–O(2)	1.427(12)
Bond angles (°)
C(1)–N(1)–Zn(1)	119.4(4)	N(1)–Zn(1)–Cl(1)^#	109.41(17)
C(5)–N(1)–Zn(1)	122.7(5)	N(1)^#–Zn(1)–Cl(1)	109.40(17)
N(1)–Zn(1)–N(1)^#	106.0(3)	N(1)–Zn(1)–Cl(1)	106.18(18)
N(1)^#–Zn(1)–Cl(1)^#	106.18(18)	Cl(1)–Zn(1)–Cl(1)^#	119.03(15)

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The donor N-atom forms strong bonds with the zinc atom: the Zn–N(1)^# bond length is 2.030(5) Å whereas the Zn–Cl(1)^# bond length is 2.195(2) Å and the C–C and N–N bond distances are comparable to those in analogous systems reported in the literature.^19,20 However, there is a significant twist within the ligand so that the planes of the tetrazole and pyridine rings form dihedral angle of 88.13 (2)°, in the two independent ligands. The complex forms intermolecular hydrogen bonds involving the oxygen atom of the carboxylate group O1 of one layer and the hydrogen atom of C1 of the pyridine group of the neighboring layer (Fig. 4 and 5) with a C(1)–H(1)⋯O(1)^# distance of 2.65 Å. The nitrogen atom of the tetrazole ring N(5)^# is connected by hydrogen bonding with the hydrogen atom of N6 of the pyridine ring of a neighboring ligand N(6)–H(6)⋯N(5)^# with a distance of 2.10 Å (Table 2).

Fig. 4 View of crystal lattice packing showing the internuclear hydrogen bonding of complex 6.

Fig. 5 Crystal packing diagram of complex 6 projecting along the crystallographic b-axis.

Table 2 Selected interatomic distance (Å) and angles (°) for complex 6^a

D–H⋯A	d(D–H) (Å)	d(H⋯A) (Å)	d(D⋯A) (Å)	<(DHA) (°)
a Symmetry transformations used to generate equivalent atoms: #1 − x + 2, y, −z + 1/2 #2 − x + 3, −y + 1, −z #3 x − 1/2, y − 1/2, z.
N(6)–H(6)⋯N(5)^#	0.82	2.10	2.884(9)	156.2
C(1)–H(1)⋯O(1)^#	0.93	2.65	3.310(10)	128.2

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FT-IR spectra

The Fourier-transform infrared (FT-IR) spectra of ligands L^1&2 show the prominent bands for ν(C=N) of the pyrimidine ring and ν(N=N) of the tetrazole ring at 1580 and 1553 cm⁻¹, and 1447 and 1433 cm⁻¹, respectively. The ligands also show bands at 3246 and 3217 cm⁻¹, and 1701 and 1721 cm⁻¹ assigned for the amino and ν(C=O) of the acetyloxy groups, respectively.^21,22 The complexation was found to result in significant changes in the absorption bands of both ν(C=N) of the pyridine ring and ν(N=N) of the tetrazole ring. In the spectra of complexes 1–3, the observed stretching frequencies ν(C=N) of the pyridine ring at 1559–1560 cm⁻¹ and ν(N=N) of the tetrazole ring at 1420–1429 cm⁻¹ are lower than those of the ligands indicating the coordination of the metal ions through the nitrogen of pyridyl and the N=N of tetrazole rings. For complexes 4–6, the observed stretching frequency of pyridine ν(C=N) at 1502–1530 cm⁻¹ is lower than that of ligands, whereas the ν(N=N) stretching frequency of the tetrazole ring remains unchanged. This may be because of the coordination of metal ions with the nitrogen of the pyridyl substituent and not with the N=N of the tetrazole ring (Fig. S5–S8†). Furthermore, significant changes are obvious in the region of stretching and stretching-deformation vibrations of the tetrazole ring (1100–900 cm⁻¹). The assignment of bands in the region 420–490 cm⁻¹ can be attributed to the coordination of N atoms of both tetrazole and pyrimidine rings ν(M–N). No significant changes in the stretching frequency of ν(C=O) is observed for ligands and complexes, indicating that the carbonyl of the acetyloxy group is not bound to the metal ion.

NMR spectra

The newly synthesized ligands L^1&2 and their zinc(II) complexes (3 and 6) were characterized by nuclear magnetic resonance (NMR) spectroscopy (Fig. S9–S16†). The ¹H-NMR spectrum of complex 6 shows considerable changes in the chemical shifts of the signals of the protons when compared to that of free ligand L², whereas complex 3 shows no change in the proton signals with respect to their free ligand L¹. This observation also supports the proposed coordination mode of the ligand L¹ through the N-atom of the tetrazole ring and the N-atom of the pyridine ring to the zinc(II) metal center for complex 3 and the ligand L² through the N-atom of the pyridine ring but not the N-atom of the tetrazole ring for complex 3.^23,24 Thus, ¹H-NMR spectra support the proposed coordination modes of the ligands with the zinc(II) metal center. The ¹³C-NMR of the compounds confirmed the presence of the tetrazole ring with the signal in the region of 150.1–148.8 ppm.

UV-Vis spectra

The electronic absorption studies allowed an insight into the possible stereochemistry of these complexes. Electronic spectra of complexes (1–6) were obtained in acetonitrile (CH₃CN) solution (Fig. S17†). The electronic spectra of nickel(II) complexes (1 and 4) display a band around 249 and 274 nm because of an intraligand transition (π–π*) or ligand–metal charge transfer transition (n–π*). The bands observed at 462 and 470 nm are attributable to the d–d transition for complexes 1 and 4, respectively. This confirms the distorted tetrahedral geometry of the nickel(II) complexes.^25,26 The copper(II) complexes (2 and 5) also exhibit bands at 259 and 260, and 307 and 310 assigned to the intraligand (π–π*) and ligand–metal charge transfer transitions (n–π*), respectively. The absorption bands at 464 and 472 nm are because of d–d transitions for complexes 2 and 5, respectively. These assignments suggest a square planar geometry for the Cu(II) complexes under investigation.²⁷ The zinc(II) complexes (3 and 6) exhibit only a high intensity band because of a strong charge transfer transition at 275 and 290 nm, respectively, assigned as a ligand–metal transition, which indicates the tetrahedral geometry around Zn(II) ions.²⁸ The absence of a band above 400 nm may be because of the lack of a metal–ligand charge transfer transition, because of the d¹⁰ electronic configuration of the zinc(II) ion. On the basis of infrared (IR), NMR, electronic spectral data and X-ray studies, distorted tetrahedral geometry is proposed for nickel(II) and zinc(II) complexes, whereas square planar geometry is proposed for copper(II) complexes.

Luminescence properties

In general, the combination of tetrazole core ligands and transition-metal centers in the coordination framework can be viewed as an efficient method for obtaining new types of luminescent materials.²⁹ In this research, the luminescence properties of complexes (1–6) were investigated in a liquid state at room temperature by using CH₃CN (Fig. 6). Complexes 1 and 2 exhibit peaks at 409 (3.03 eV) and 400 nm (3.10 eV), respectively. The emission is neither metal-to-ligand charge transfer (MLCT) nor ligand-to-metal transfer in nature and can be assigned to intraligand fluorescent emission, because a similar emission (λ_max = 380–400 nm) is also observed for free ligand L¹.

Fig. 6 Emission spectra of complexes 1 and 4 at room temperature.

Complexes 4 and 5 display strong emission peaks at 334 (3.71 eV) and 385 nm (3.22 eV), respectively, which may be assigned to intraligand fluorescent emission with an enhanced fluorescent intensity that is almost twice that of free ligand L² (λ_max = 380 nm). The emission behavior of these complexes is likely to be because of (π–π*) and (n–π*) transitions of the pyridyl ring of the ligands and suggests direct bonding of the metal with the C=N of the pyridyl ring, and thus, a less rigid structure. In the case of complexes of L¹, the metal binds with both the C=N of the pyridine ring and the N=N of the tetrazole ring, thus, a comparatively more rigid structure may be assigned. This may be probably due to ligand conformational rigidity, thereby reducing the non-radiative decay of the intraligand (π–π*) excited state of the nickel(II) and copper(II) complexes (4 and 5) of L².^30,31

It is universally acknowledged that metal coordination frameworks with a d¹⁰ configuration possess excellent luminescence properties. The emission spectra of zinc(II) complexes (3 and 6) have their emission peaks centered at 342 (3.62 eV) and 381 nm (3.25 eV), respectively. As we know, it is usually believed that the energy transition of d¹⁰ complexes is tentatively attributed to the ligation of the ligand to the metal center and not to MLCT.³²

EPR spectral data

The solid state electroparamagnetic resonance (EPR) spectra of copper(II) complexes (2 and 5) were recorded in the X-band region at room temperature (25 °C). Fig. S18† shows the EPR spectrum of complex 2 and the data are summarized in Table 3. Complexes 2 and 5, exhibit g_∥ values of 2.20 and 2.16 and g_⊥ values of 2.09 and 2.05, respectively. These values indicate that the ground state of Cu(II) is predominantly d_{x² − y²}.

Table 3 EPR spectral assignments for complexes 2 and 5 at room temperature

Complexes	g∥	g⊥	gav	A∥ × 10^−4 (cm^−1)	F (cm)	α^2	G
2	2.20	2.09	2.12	184.33	119	0.785	2.25
5	2.16	2.05	2.08	183.02	118	0.725	3.30

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The geometric parameter G, which is a measure of the exchange interaction between the copper centers in the polycrystalline compound, is calculated using the equation:

G = (g_∥ − 2.0023)/(g_⊥ − 2.0023)

According to Hathaway and Tomlinson,³³ if G > 4.0, considerable exchange interaction is negligible because the local tetragonal axes are aligned parallel or slightly misaligned. If G < 4.0, exchange is considerable and the local tetragonal axes are misaligned. The observed G values of complexes 2 and 5 (2.25 and 3.30, respectively) suggest that there is no exchange interaction in the copper(II) complexes. The covalency parameter α² is calculated using the following equation:^34,35

α² = (A_∥/0.036) + (g_∥ − 2.0023) + 3/7(g_⊥ − 2.0023) + 0.04

If the value of α² = 0.5, it indicates complete covalent bonding, whereas if the value of α² = 1.0 it suggests complete ionic bonding. The observed values of α² (0.725 and 0.785) for the complexes are less than unity, which indicates that the complexes have covalent character in the ligand environment.

The two parameters g_∥ and A_∥ are to some extent correlated by the empirical factor f = g_∥/A_∥.^36,37 Within a related series of compounds, an increasing covalency of the metal–ligand bonds leads to a decrease in g_∥ and an increase in A_∥. As the covalency increases, the energies of the excited states rise, so that the orbital contribution to g_∥ becomes less effective and the g_∥ becomes closer to the free-electron value. The effect of covalency on the A_∥ values is impossible to predict, because there are so many factors involved, but theoretical values also suggest that there is a considerable dependence on A_∥ and g_∥. The best discrimination is obtained from the parallel values, because these show the widest variation.³⁸ All square planar compounds fall in the region shown in Fig. 7, with the complexes of softer ligands lying towards the upper part of the band. Distortion towards tetrahedral geometry gives parameters which lie below this band. The ratio g_∥/A_∥ suggests that the Cu(II) complexes lie towards the upper part of the band, suggesting square planar geometry.

Fig. 7 Correlation between g_∥ and A_∥ for complexes 2 and 5.

Magnetic susceptibility data

The measurement of molar magnetic susceptibility (χ_M) allows us to determine the number of unpaired electrons associated with the transition metal in the compound and is useful in understanding the compound's bonding, and its magnetic and spectral characteristics. The effective magnetic moments of complexes 2 and 5 are 1.88 and 1.93 BM, respectively, slightly higher than the spin only values (1.73 μ_eff) expected for a d⁹ system with one unpaired electron.³⁹ The magnetic moments of Ni(II) complexes 1 and 4 are 3.3 and 3.5 BM, respectively, which indicates a paramagnetic nature because of the tetrahedral geometry around the metal ion.⁴⁰ The Zn(II) complexes (3 and 6) under study were found to be diamagnetic in nature because of the d¹⁰ system, as expected.

Interaction with calf thymus DNA

Absorption spectral studies. Electronic absorption spectroscopy is widely employed to determine the DNA binding affinity of metal complexes. The absorption spectra of the Cu(II) complexes (2 and 5) in the absence and presence of DNA are shown in Fig. 8. The absorption bands were affected by increasing concentrations of DNA. A strong hyperchromism together with a minor blue shift indicates a strong interaction of complexes with calf thymus–DNA (CT–DNA) mainly through groove binding.⁴¹ DNA possesses several hydrogen bonding sites which are accessible both in the major and minor grooves.⁴² It is well known that the interactions of chemical species with the minor groove of DNA differ from those occurring in the major groove, both in terms of electrostatic potential and steric effects, because of the narrow shape of the former. For the minor groove, the backbones of the DNA are closer together with an 8.2 Å depth, whereas the major groove has the backbones far apart with an 11.6 Å width and an 8.5 Å depth, offering easy access to bulky molecules.^43,44 Thus, it is easier for certain metal complexes to interact with the bases on the major groove side because the backbone is not in the way. In this context complex 2 may interact with DNA in the minor groove because it has little steric interference compared to complex 5 which may bind to DNA in major groove fashion.⁴⁵ In order to compare the quantitative DNA-binding affinities of these complexes, their intrinsic binding constants were obtained by monitoring the changes in absorption at intraligand bands (π–π*) with increasing concentration of DNA. When the concentration of CT–DNA was increased, a strong hyperchromic effect was observed for complexes 2 and 5, although no appreciable change in the position of the intraligand band was observed. The binding constant (K_b) values are found to be 1.91 × 10⁵ M⁻¹ and 2.8 × 10⁵ M⁻¹ for complexes 2 and 5, respectively. The higher hyperchromism and K_b value obtained for complex 5 suggest it has higher binding affinity to DNA compared with that for the complex 2. DNA binding studies reveal that both the complexes prefer groove binding; complex 2 binds to CT–DNA through the minor groove and complex 5 through the major grooves of CT–DNA.

Fig. 8 Absorption spectra of complexes 2 (a) and 5 (b) (10 μM) in Tris–HCl buffer upon addition of CT–DNA (0–100 μM). Arrows indicate the changes in absorbance upon increasing DNA concentration. Inset: plots of [DNA]/(ε_a − ε_f) versus [DNA] for absorption titration of CT–DNA with complexes.

Fluorescence quenching studies. To further understand the mode of interaction between the copper(II) complexes (2 and 5) with CT–DNA, ethidium bromide (EB) fluorescence displacement experiments were carried out, by monitoring the changes in emission intensity of EB bound to DNA as a function of added complex concentration.⁴⁶ The emission spectra of EB bound to DNA in the absence and presence of complexes 2 and 5 are shown in Fig. 9. Upon addition of the complexes to DNA pretreated with EB, an appreciable reduction in the DNA-induced emission intensity of EB is caused, indicating the binding of complexes with DNA at the sites occupied by EB. The data were analyzed by using the Stern–Volmer equation. The quenching plots (inset of the respective figures) illustrate that the fluorescence quenching of EB bound to DNA by complexes 2 and 5 is in linear agreement with the Stern–Volmer equation, which confirms the interaction of the complexes with DNA.

Fig. 9 Fluorescence spectra of EB bound to DNA in the absence and presence of increasing amount of complexes 2 (a) and 5 (b) at 37 °C: [complex] = (0–20 μM), [DNA]/[EB] = 10 μM after excitation at λ_exc 459 and 393 nm. Inset: plots of I₀/I versus [Q] × 10⁴ M⁻¹ for the titration of complex with [DNA]/[EB].

The K_sv values for complexes 2 and 5 are 0.52 and 1.92 × 10⁻⁴ M⁻¹, respectively. These values prove the partial replacement of EB bound to DNA by the two complexes which results in a decrease in fluorescence intensity. The intrinsic binding parameters clearly show that complexes 2 and 5 strongly interact with CT–DNA. A large increase in emission intensity suggests a stronger binding propensity to CT–DNA.

Viscosity measurements. To further clarify the interaction mode of the complexes with DNA, viscosity measurements were carried out. In classical intercalation, the DNA helix lengthens as the base pairs are separated to accommodate the binding ligand, leading to an increase in DNA viscosity.⁴⁷ In contrast, a partial and/or non-classical intercalation of ligand would bend (or kink) the DNA helix, reducing its effective length, and thereby, its viscosity, while ligands that bind exclusively in the DNA grooves (e.g., netropsin, distamycin), under the same conditions, typically cause less pronounced changes (positive or negative) or no changes in the DNA viscosity.⁴⁸ The plots of relative viscosity (η/η₀)^1/3 versus binding ratio [complex]/[DNA] are shown in Fig. 10 for distamycin and complexes 2 and 5.

Fig. 10 Effect of the increase in amounts of distamycin (▼), and complexes 2 (○) and 5 (■) on the relative viscosity of CT–DNA at 28.0 ± 0.5 °C.

Distamycin, a well-known classical groove binder, caused a change in DNA viscosity upon complexation. Interestingly, complex 5 confers a greater enhancement in DNA viscosity than complex 2. The hydrophobic interaction of the ligand of the bulkier complex with the DNA major groove, followed by aggregation of the free complex in solution with the DNA-bound complex through the overlap of ligands, leads to untwisting of the DNA helix at the binding sites and thus, to an increase in overall DNA counter length.^49,50 In contrast, complex 2, which is expected to bind with DNA minor grooves, showed a moderate change in viscosity. The binding ability of complexes to increase the viscosity of the DNA varies in the order distamycin > 5 > 2, which parallels the hyperchromism and DNA binding affinities. The results of this study clearly show that complexes 2 and 5 preferentially bound to DNA through minor and major grooves, respectively. Thus, the viscosity measurements are consistent with the results of the spectral experiments. Therefore, we propose the groove modes of binding for the two complexes.

Molecular docking of the complexes with DNA

The molecular docking of the complexes (2 and 5) with the self-complementary DNA duplex of sequence dodecamer d(CGCGAATTCGCG)₂ demonstrates that the complexes are stabilized by additional electrostatic and hydrogen bonding interactions with the DNA (Fig. 11). The docked conformation of complexes 2 and 5 was analysed in terms of energy, hydrogen bonding, and hydrophobic interactions between the complexes and DNA. Detailed analyses of the complex–DNA interactions were carried out, and final coordinates of the complexes and receptor were saved. For display of the receptor with the complex binding site, PyMOL software was used. From the docking scores, the free energy of binding (FEB) of the compounds was calculated (Table 4). The complexes adopt a characteristic shape and are flexible enough to adopt a conformation which is complementary to the minor and major groove. The molecular docking results show that complexes 2 and 5 bind efficiently with the DNA receptor and exhibit FEB values −8.48 and −9.77 kcal mol⁻¹, respectively. The more negative relative binding energy of complex 5 indicated that it binds more strongly to the DNA than does complex 2. Because right handed DNA is complementary to complex 5, it exhibits strong interactions via symmetrical hydrogen bonding.⁵¹ The best possible conformation of the mononuclear copper(II) complexes 2 and 5 is through the interactions of the tetrazole, pyridine and acetyloxy groups of the ligand to DNA through the minor and major grooves, which is because of stabilization by hydrogen bonding. Furthermore, in complex 5, tetrazole nitrogens interact with DA-6, DC-21 and DT-21 residues, the pyridine nitrogen interacts with the DG-4 residue, and carbonyl oxygens of the ester substituents interact with DG-2 and DC-23 residues of DNA. In complex 2, the tetrazole nitrogen interacts with the DC-11 residue, carbonyl oxygens of the ester substituents interact with DA-16 and DA-17 residues and oxygen of the ester substituent interacts with the DG-10 residue of DNA. Thus, it can be concluded that molecular docking studies provide additional evidence for the preferred minor and major groove modes of binding with the copper(II) complexes.

Fig. 11 Molecular docked model of complexes 2 and 5 with DNA (PDB ID: 1BNA) dodecamer duplex of sequence d(CGCGAATTCGCG)₂.

Table 4 Molecular docking parameters of the complexes

Complex	Final intermolecular energy (kcal mol^−1)			Final total internal energy (kcal mol^−1) (2)	Torsional free energy (kcal mol^−1) (3)	Unbound system's energy [= (2)] (kcal mol^−1) (4)	Estimated free energy of binding [(1) + (2) + (3) − (4)] (kcal mol^−1)
	vdW + H bond + dissolving energy	Electrostatic energy	Total (1)
2	−8.19	−1.11	−9.30	−0.66	+0.82	−0.66	−8.48
5	−10.90	−1.62	−12.51	−3.03	+2.74	−3.03	−9.77

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Cytotoxicity evaluation

In vitro cytotoxic activity of all the ligands (L^1&2) and complexes (1–6) was done by MTT [3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide] reduction assays on three different cancer cell lines, namely lung (A549), cervical (HeLa) and colon (HCT-15), together with a normal human embryonic kidney (HEK-293) cell line. For cytotoxic activity, four different concentration of compounds (100, 250, 500 and 1000 nM) and cisplatin (positive control) were tested in triplicate for 48 h in two independent experiments. The results are expressed as the average ± standard deviation of two independent experiments. The inhibitory concentration (IC₅₀) of the complexes exhibited differential and dose-dependent inhibitory activity on lung (A549), cervical (HeLa), and colon (HCT-15) cancer cells (Table 5).

Table 5 IC₅₀^a values of ligands, complexes and cisplatin against lung (A549), cervical (HeLa) and colon (HCT-15) cancer cells

Compounds	Lung (A549)	Cervical (HeLa)	Colon (HCT-15)
a Na*: not active; inhibitory concentration (IC50) values are derived from dose–response curves obtained by measuring the percentage of viable cells relative to untreated and control after a 48 h exposure to test compounds using the MTT assay. Values represent means of two independent experiments.
DMSO	Na*	Na*	Na*
L^1	Na*	Na*	Na*
L^2	Na*	Na*	Na*
1	988 ± 0.14	>1000	120.5 ± 0.22
2	409.4 ± 0.12	521.5 ± 0.1	69.13 ± 0.13
3	438.4 ± 0.4	>1000	166.4 ± 0.26
4	>1000	318.4 ± 0.4	416.3 ± 0.17
5	715.3 ± 0.12	508.9 ± 0.3	33.51 ± 0.16
6	531.7 ± 0.6	978.4 ± 1.2	569.3 ± 0.76
Cisplatin	169 ± 0.8	160.8 ± 1.3	189.3 ± 0.9

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All the complexes were active against the tested cell lines but the ligands showed no activity. The complexes 1, 2, 3 and 5 exhibited excellent activity against the colon cancer cell line (HCT-15) when compared with the lung (A549) and cervical (HeLa) cancer cells with respect to the standard drug cisplatin. When comparing the inhibitory effect with respect to IC₅₀ values, complexes 2 and 5 showed very high activity compared with the other complexes (Fig. 12 and 13). However, complexes exhibited minimum (less than 20%) cytotoxic effect on normal human embryonic kidney cells (HEK-293).

Fig. 12 Cytotoxic activity of complex 2 against non-cancer and cancer cell lines with different concentrations of complex at 48 h.

Fig. 13 Cytotoxic activity of complex 5 against non-cancer and cancer cell lines with different concentrations of complex at 48 h.

Apoptosis activity of complexes 2 and 5 by live cell and fluorescent imaging

To further address the anticancer activity pattern, the cancer cell lines: lung (A549), cervical (HeLa) and colon (HCT-15) were treated with IC₅₀ concentrations of complexes 2 and 5, and the cells were observed by phase contrast microscope after staining with acridine orange – EB, which shows dead and floating cells. The acridine orange – EB staining shows the morphological aspect of chromatin condensation in the stained nucleus, allowing one to distinguish viable, apoptotic and necrotic cells. Viable cells possess a uniform green nucleus, early apoptotic cells show bright green areas of condensed or fragmented chromatin in the nucleus, and necrotic cells show a uniform bright orange nucleus. After staining with acridine orange – EB, the cancer cells treated with copper(II) complexes showed apoptotic characteristics such as changes in cell morphology, cell shrinkage and nuclear fragmentation, and an increase in permeability of the cell membrane (Fig. 14). The results of the staining assay demonstrated by the apoptotic features such as nuclear shrinkage, chromatin condensation and nuclear fragmentation after 48 h (Fig. 15) showed that complexes 2 and 5 exhibited higher activity against colon (HCT-15) cancer cells than lung (A549) and cervical (HeLa) cancer cell lines.

Fig. 14 Live cell imaging of lung (A549), cervical (HeLa) and colon (HCT-15) cancer cells treated with IC₅₀ concentrations of complexes 2 and 5.

Fig. 15 The morphological changes of lung (A549), cervical (HeLa) and colon (HCT-15) cancer cells treated with IC₅₀ concentrations of complexes 2 and 5 after staining with acridine orange – EB.

The mechanism of action of these complexes is still not completely elucidated. Efforts are continuing to establish the mechanism of action. However, there is evidence that supports the idea that these compounds are able to inhibit cell proliferation and produce dose-dependent cell death by apoptosis. Inhibition of cancer cells and induction of apoptosis might be efficient ways of treating cancer. Apoptosis or programmed cell death involves the activation of energy requiring intracellular machinery, which is tightly regulated and conserved throughout evolution.⁵² Apoptosis eliminates cells exposing the organism to danger. For example, virally infected cells or cells with damaged DNA will be removed by apoptosis.⁵³ Cancer cells evade apoptosis by several mechanisms. There are two types of apoptosis, the death receptor-dependent pathway, also called the extrinsic pathway, and the mitochondrial-dependent or intrinsic pathway. Therefore, these complexes have the potential to act as effective metal-based anticancer drugs.

Conclusions

Six new complexes of tetrazolo[1,5-a]pyrimidine core ligands of type [ML¹Cl₂] (1–3) and [M(L²)₂Cl₂] (4–6) have been synthesized and characterized by spectral methods. The complexes 1–3 and 4–6 were formed in 1 : 1 and 1 : 2 metal-to-ligand mole ratios, respectively. Based on the spectral data, square planar geometry was assigned for copper(II) complexes (2 and 5) while tetrahedral geometry was assigned for nickel(II) (1 and 4) and zinc(II) complexes (3 and 6). Single crystal X-ray diffraction (XRD) results for complex 6 revealed distorted tetrahedral geometry around the metal ion involving two Cl atoms and the N atom of the two pyridine rings. DNA binding studies revealed that the complexes show a preference for the mode of groove binding; complex 2 binds to CT–DNA through the minor groove and complex 5 binds to the major grooves of DNA, which was also supported by molecular docking experiments. The cytotoxicity results showed that the complexes selectively inhibited cancer cells whereas the ligands were found to be inactive against both cancer and non-cancer cell lines. The cytotoxicity of the complexes was arranged in the order 5 > 2 > 1 based on their inhibitory effects against colon cells. Furthermore, the apoptotic results of complexes 2 and 5 suggest that these complexes can act as promising anticancer agents for cancer therapy.

Experimental

Materials and methods

Metal(II) chlorides and other commercially available reagents (5-aminotetrazole, sodium thiosulfate, iodine and ethyl acetoacetate) were purchased from Aldrich and used without further purification. Solvents for synthesis were reagent grade or better and purified according to standard methods.⁵⁴ CT–DNA was purchased from Bangalore Genei (India). The melting points were determined using electrothermal capillary apparatus and they are reported as uncorrected values. Elemental analyses were performed using a Carlo Erba 1106 elemental analyzer. The FT-IR spectra were recorded on a Thermo Nicolet 6700 FT-IR spectrometer using KBr pellets in the range of 400–4000 cm⁻¹. The ¹H-NMR and ¹³C-NMR spectra were recorded on an Bruker Avance 400 MHz spectrometer using deuterated dimethyl sulfoxide (DMSO-d₆) and deuterated chloroform (CDCl₃) as solvent. The mass data of ligands and complexes were obtained on a Jeol DX-303 mass spectrometer and a Thermo Finnigan LCQ Advantage MAX 6000 electrospray ionisation (ESI) spectrometer, respectively. UV-Vis spectra were recorded on a Shimadzu UV-2450 spectrophotometer using CH₃CN as solvent. Fluorescence spectra were recorded on a Horiba Jobin Yvon FluoroLog SPEX-F311 spectrofluorometer. X-band EPR spectra were recorded at 25 °C on a Varian EPR E-112 spectrometer using 2,2′-diphenyl-1-picrylhydrazyl (DPPH) as the reference. Room temperature magnetic studies were performed on a PAR 155 vibrating sample magnetometer.

Caution! Metal tetrazolate complexes are potentially explosive. Only a small amount of material should be prepared and handled with caution.

DNA binding experiments

Absorption titrations. All the experiments involving the interaction of complexes with DNA were carried out in Tris–HCl/NaCl buffer (Tris–HCl 5 mM/NaCl 50 mM, pH 7.2). A solution of CT–DNA in the buffer gave an acceptable ratio of UV absorbance at 260 and 280 nm, indicating that the DNA was sufficiently free from proteins.⁵⁵ The DNA concentration was determined by absorption spectroscopy using the molar absorption coefficient 6600 M⁻¹ cm⁻¹ at 260 nm.⁵⁶ Absorption titration experiments were performed by keeping the complex concentration constant (10 μM) and varying the concentration of DNA (0–100 μM). While measuring the absorption spectra, equal amounts of DNA were added to both the complex solution and the reference solution to eliminate the absorbance of DNA itself. After the addition of DNA to the metal complexes, the resulting solution was allowed to equilibrate at room temperature for 5 min. Then, the sample solution was scanned in the range 200–350 nm. The binding constant (K_b) was determined from the spectroscopic titration data using the following equation:⁵⁷

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

where [DNA] is the concentration of CT–DNA in the base pairs. The apparent absorption coefficient (ε_a) was obtained by calculating A_obs/[complex]. The terms ε_f, and ε_b correspond to the extinction coefficient of free (unbound) and fully bound compound, respectively. A plot of [DNA]/(ε_a − ε_f) versus [DNA] will give a slope 1/(ε_b − ε_f) and an intercept 1/K_b(ε_b − ε_f). K_b is given by the ratio of the slope to the intercept.

Fluorescence spectral studies. For fluorescence quenching experiments, DNA was pretreated with 1% dimethyl formamide in Tris–HCl/NaCl (pH 7.2) buffer solution as a blank to make preliminary adjustments. The competitive binding experiments were carried out in the buffer by keeping the concentration of [DNA]/[EB] constant at 10 μM, and varying the concentration of the complexes (0–20 μM) at 37 °C. Before measurements, the mixture was shaken up and incubated at room temperature for 30 min. The fluorescence spectra of EB were measured using an excitation wavelength of 380–460 nm and the emission range was set between 300 and 650 nm. Relative binding of these complexes to CT–DNA is measured from the extent of reduction in emission intensity. The Stern–Volmer quenching constant value was calculated using the equation:⁵⁸

I₀/I = 1 + K_sv[Q]

where I₀ is the emission intensity of EB–DNA in the absence of complex, I is the emission intensity of EB–DNA in the presence of complex, and [Q] is the concentration of quencher. K_sv is the linear Stern–Volmer constant, obtained as a slope of intercept I₀/I versus [Q].

Viscosity experiments. Viscosity experiments were carried out on an Ostwald's viscometer at a constant temperature of 28.0 ± 0.5 °C in a thermostated bath. Flow time was measured with a digital stopwatch for different concentrations of complexes (10–50 μM), but maintaining the initial DNA concentration (100 μM). Each sample was measured three times and an average flow time was calculated. The DNA viscosity was calculated according to: η_i = (t_i − t₀)/t₀, where η_i is the corresponding values of DNA viscosity, t_i is the flow time of the solution in the presence or absence of the complexes, and t₀ is the flow time of buffer alone. Data were presented as (η/η₀)^1/3 versus binding ratio, where η is the viscosity of DNA in the presence of complex and η₀ is the viscosity of DNA alone.⁵⁹

Molecular docking studies. Molecular docking studies were carried out using the AutoDock Tools (ADT) version 1.5.6 and AutoDock version 4.2.5.1 docking programmes. This is an interactive molecular graphics program used to understand the drug–DNA interactions and investigate the potential binding mode of complexes. Docking studies were performed using an Intel® Core™ i5 CPU (2.53 GHz) with the Windows 7 operating system.

DNA preparation. The X-ray crystal structure of B-DNA (PDB ID: 1BNA) dodecamer d(CGCGAATTCGCG)₂ was obtained from the Protein Data Bank (http://www.rcsb.org/pdb). For the DNA, hydrogen were added before computing Gasteiger charges, and then non-polar hydrogen atoms were merged. The distance between donor and acceptor atoms forming a hydrogen bond was defined as 1.9 Å with a tolerance of 0.5 Å, and the acceptor–hydrogen–donor angle was not less than 120°. The structures were then saved in PDBQT file format, for further studies in ADT.

Complex preparation. The two-dimensional (2D) structures of complexes 2 and 5 were drawn using ChemDraw Ultra 12.0 (ChemOffice 2010) software. Chem3D Ultra 12.0 was used to convert the 2D structures into three-dimensional (3D) and the energy was minimized using the semi-empirical AM1 method. Energy minimization was performed with a minimum root mean squared gradient of 0.100 in each iteration. All structures were saved as pdb file format for input to ADT. The structures of complexes were then saved in PDBQT file format to carry out docking in ADT.

Grid formation. Preparation of parameter files for grid and docking was done using ADT. The copper parameters, van der Waals (vdW) radii of 0.96 Å and vdW well depth of 0.01 kcal mol⁻¹ were used in the docking calculation.⁶⁰ A grid box with dimensions of 60 × 60 × 110 ų with 0.375 Å spacing and centered on (x, y, z) 14.779, 20.976, 8.804 was created that included the whole DNA using ADT. The center of the box was set at the DNA center and grid energy calculations were carried out.

Synthesis of tetrazolo[1,5-a]pyrimidine core ligands

Ethyl 5-methyl-7-pyridine-2-yl-4,7-dihydrotetrazolo[1,5-a]pyrimidine-6-carboxylate (L¹). A mixture of 5-aminotetrazole (0.52 g, 5 mmol), 2-pyridinecarboxaldehyde (0.54 g, 5 mmol) and ethyl acetoacetate (0.65 g, 5 mmol) in iso-propyl alcohol (5 mL) was put in a round bottomed flask and iodine (0.13 g, 0.5 mmol) was added to the stirred mixture at a refluxing temperature (82–85 °C). After refluxing for 4 h, the whole mixture was cooled to room temperature and then a solution of sodium thiosulfate (10 M, 2 mL) was added to the mixture. The precipitated solid was filtered and washed with cold methanol (2 mL) followed by water. The crude product was dried and recrystallized from hot ethanol to give L¹ as white crystals.

Yield: (1 g, 69.9%). Melting point (mp) 260 °C. Analytical calculation for C₁₃H₁₄N₆O₂ (%): C, 54.57; H, 4.93; N, 29.37; found: C, 54.52; H, 4.89; N, 29.35. Selected IR data (KBr, ν, cm⁻¹): ν 3246–2937 (br, N–H), 1580 (s, C=N, pyridine ring), 1447 (s, N=N, tetrazole ring), 1701 (s, C=O). ¹H-NMR (400 MHz: DMSO-d₆ and CDCl₃: Me₄Si) δ (ppm): 1.03–1.07 (t, 3H, CH₃ (Et)), 2.42 (s, 3H, –CH₃), 3.93–4.01 (m, 2H, –CH₂Me), 6.63 (s, 1H, –CH–), 7.12–7.15 (t, J = 0.8 Hz, 1H, –C₅H₄N), 7.40–7.42 (d, J = 0.8 Hz, 1H, –C₅H₄N), 7.59–7.65 (m, 1H, –C₅H₄N), 8.35–8.37 (dd, J = 4.0, 0.8 Hz, 1H, –C₅H₄N), 10.91 (s, 1H, –NH). ¹³C-NMR (100 MHz: DMSO-d₆ and CDCl₃: Me₄Si) δ (ppm): 164.2, 157.4, 148.8, 148.6, 146.9, 135.8, 122.6, 121.5, 96.3, 59.3, 59.1, 18.2, 13.2. ESI-MS (m/z): 286 (M⁺, 100.0%), 259 (20), 214 (15).

Ethyl 5-methyl-7-pyridine-4-yl-4,7-dihydrotetrazolo[1,5-a]pyrimidine-6-carboxylate (L²). The procedure for synthesis was similar to that of L¹ except that 4-pyridinecarboxaldehyde was used in place of 2-pyridinecarboxaldehyde to give L² as white crystals.

Yield: (1.35 g, 94.0%). mp 195 °C. Analytical calculation for C₁₃H₁₆N₆O₃ (%): C, 51.30; H, 5.29; N, 27.61; found: C, 51.26; H, 5.25; N, 27.60. Selected IR data (KBr, ν, cm⁻¹): ν 3217–2981 (br, N–H), 1553 (s, C=N, pyridine ring), 1433 (s, N=N, tetrazole ring), 1721 (s, C=O). ¹H-NMR (400 MHz): (DMSO-d₆ and CDCl₃: Me₄Si) δ (ppm): 1.10–1.14 (t, 3H, CH₃(Et)), 2.65 (s, 3H, –CH₃), 4.04–4.12 (m, 2H, –CH₂Me), 6.69 (s, 1H, –CH–), 7.27–7.29 (dd, J = 4.8, 1.6 Hz, 2H, –C₅H₄N), 8.59–8.61 (d–d, J = 4.4, 1.6 Hz, 2H, –C₅H₄N), 11.47 (s, 1H, –NH). ¹³C-NMR (100 MHz: DMSO-d₆ and CDCl₃: Me₄Si) δ (ppm): 164.5, 150.1, 148.7, 148.3, 147.4, 122.2, 97.5, 60.8, 58.3, 19.4, 14.0. ESI-MS (m/z): 305.3 (M⁺ + H, 100.0%), 259 (25), 214 (15).

General procedure for synthesis of mononuclear copper(II), nickel(II) and zinc(II) complexes

To a methanolic solution (25 mL) of ligand: L¹ (1.36 mmol) or L² (2.72 mmol), was added a methanolic solution (25 mL) of metal(II) chloride (1.36 mmol) under constant stirring. Then, the reaction mixture was refluxed for 2 h, filtered hot and allowed to stand at room temperature for 7 days. The crystalline product obtained was washed with cold methanol and dried in vacuo.

(Ethyl 5-methyl-7-pyridine-2-yl-4,7-dihydrotetrazolo[1,5-a]pyrimidine-6-carboxylato)dichloronickel(II) complex [NiL¹Cl₂] (1). Yield: (0.36 g, 64.10%). Green solid. Analytical calculation for C₁₃H₁₄N₆O₂Cl₂Ni (%): C, 37.72; H, 3.40; N, 20.30; found: C, 37.69; H, 3.38; N, 20.29. Selected IR data (KBr, ν, cm⁻¹): ν 3249–2978 (br, N–H), 1560 (s, C=N, pyridine ring), 1429 (s, N=N, tetrazole ring), 1706 (s, C=O). UV-Vis [(λ_max (nm) (ε, dm³ mol⁻¹ cm⁻¹))] in CH₃CN: 249 (11, 400), 462 (590). ESI-MS (m/z): 415 ([NiL¹Cl₂]⁺; 40%), 380 ([NiL¹Cl]⁺; 60%), 344 ([NiL¹]⁺; 50%), 287 ([L¹]⁺ + H; 90%). μ_eff: 3.33 BM.

(Ethyl 5-methyl-7-pyridine-2-yl-4,7-dihydrotetrazolo[1,5-a]pyrimidine-6-carboxylato)dichlorocopper(II) complex [CuL¹Cl₂] (2). Yield: (0.40 g, 70.37%). Green solid. Analytical calculation for C₁₃H₁₄N₆O₂Cl₂Cu (%): C, 37.27; H, 3.36; N, 20.06; found: C, 37.24; H, 3.34; N, 20.05. Selected IR data (KBr, ν, cm⁻¹): ν 3096–2978 (br, N–H), 1559 (s, C=N, pyridine ring), 1420 (s, N=N, tetrazole ring), 1705 (s, C=O). UV-Vis [(λ_max (nm) (ε, dm³ mol⁻¹ cm⁻¹))] in CH₃CN: 259 (7816), 307 (7400), 464 (1580). g_∥: 2.20, g_⊥: 2.09 and A_∥: 184.33. ESI-Mass (m/z): 420.55 ([CuL¹Cl₂]⁺; 40%), 385 ([CuL¹Cl]⁺; 70%), 349 ([CuL²]⁺; 40%), 286 ([L¹]⁺; 75%). μ_eff: 1.88 BM.

(Ethyl 5-methyl-7-pyridine-2-yl-4,7-dihydrotetrazolo[1,5-a]pyrimidine-6-carboxylato)dichlorozinc(II) complex [ZnL¹Cl₂] (3). Yield: (0.30 g, 52.65%). Colourless solid. Analytical calculation for C₁₃H₁₄N₆O₂Cl₂Zn (%): C, 37.18; H, 3.36; N, 20.01; found: C, 37.15; H, 3.33; N, 20.00. Selected IR data (KBr, ν, cm⁻¹): ν 3246–2963 (br, N–H), 1560 (s, C=N, pyridine ring), 1429 (s, N=N, tetrazole ring), 1706 (s, C=O). ¹H-NMR (400 MHz: DMSO-d₆ and CDCl₃: Me₄Si) δ (ppm): 1.03–1.07 (t, 3H, CH₃(Et)), 2.41 (s, 3H, –CH₃), 3.93–4.03 (m, 2H, –CH₂Me), 6.69 (s, 1H, –CH–), 7.23–7.26 (t, J = 0.8 Hz, 1H, –C₅H₄N), 7.51–7.53 (d, J = 8.0 Hz, 1H, –C₅H₄N), 7.73–8.16 (m, 1H, –C₅H₄N), 8.40 (dd, J = 4.8, 0.8 Hz, 1H, –C₅H₄N), 11.16 (s, 1H, –NH). ¹³C-NMR (100 MHz): DMSO-d₆ and CDCl₃: Me₄Si) δ (ppm): δ 164.4, 158.2, 149.5, 149.2, 147.1, 136.6, 123.4, 122.3, 96.9, 59.5, 18.5, 13.8. UV-Vis [(λ_max (nm) (ε, dm³ mol⁻¹ cm⁻¹))] in CH₃CN: 275 (15 000). ESI-MS (m/z): 422.70 ([ZnL¹Cl₂]⁺; 40%), 387 ([ZnL¹Cl]⁺; 60%), 351 ([ZnL¹]⁺; 70%), 286 ([L¹]⁺; 65%).

Bis(ethyl 5-methyl-7-pyridine-4-yl-4,7-dihydrotetrazolo[1,5-a]pyrimidine-6-carboxylato)dichloronickel(II) complex [Ni(L²)₂Cl₂] (4). Yield: (0.75 g, 77.31%). Yellowish green solid. Analytical calculation for C₂₆H₂₈N₁₂O₄Cl₂Ni (%): C, 44.60; H, 4.03; N, 24.00; found: C, 44.56; H, 3.99; N, 23.99. Selected IR data (KBr, ν, cm⁻¹): ν 3010–2966 (br, N–H), 1520 (s, C=N, pyridine ring), 1430 (s, N=N, tetrazole ring), 1720 (s, C=O). UV-Vis [(λ_max (nm) (ε, dm³ mol⁻¹ cm⁻¹))] in CH₃CN: 274 (13 600), 470 (490). ESI-MS (m/z): 703 ([Ni(L³)₂Cl₂⁺ + 1H; 50%), 665 ([Ni(L³)₂Cl⁺; 95%), 629 ([Ni(L³)₂⁺; 15%). μ_eff: 3.35 BM.

Bis(ethyl 5-methyl-7-pyridine-4-yl-4,7-dihydrotetrazolo[1,5-a]pyrimidine-6-carboxylato)dichlorocopper(II) complex [Cu(L²)₂Cl₂] (5). Yield: (0.84 g, 86.59%). Green solid. Analytical calculation for C₂₆H₂₈N₁₂O₄Cl₂Cu (%): C, 44.28; H, 4.00; N, 23.83; found: C, 44.24; H, 3.97; N, 23.82. Selected IR data (KBr, ν, cm⁻¹): ν 3190–2928 (br, N–H), 1502 (s, C=N, pyridine ring), 1430 (s, N=N, tetrazole ring), 1719 (s, C=O). UV-Vis [(λ_max (nm) (ε, dm³ mol⁻¹ cm⁻¹))] in CH₃CN: 260 (7600), 310 (7500), 472 (1430). g_∥: 2.16, g_⊥: 2.05 and A_∥: 183.02. ESI-MS (m/z): 717 ([Cu(L³)₂Cl₂]⁺; 25%), 671 ([Cu(L³)₂Cl⁺; 636 ([Cu(L³)₂⁺; 25%). μ_eff: 1.93 BM.

Bis(ethyl 5-methyl-7-pyridine-4-yl-4,7-dihydrotetrazolo[1,5-a]pyrimidine-6-carboxylato)dichlorozinc(II) complex [Zn(L²)₂Cl₂] (6). Yield: (0.75 g, 76.49%). Colourless solid. Analytical calculation for C₂₈H₃₇N₁₂O_6.5Cl₂Zn (%): C, 43.00; H, 4.76; N, 21.49; found: C, 42.96; H, 4.73; N, 21.48. Selected IR data (KBr, ν, cm⁻¹): ν 3243–2934 (br, N–H), 1530 (s, C=N, pyridine ring), 1431 (s, N=N, tetrazole ring), 1720 (s, C=O). ¹H-NMR (400 MHz: DMSO-d₆ and CDCl₃: Me₄Si) δ (ppm): 1.07 (t, 3H, CH₃(Et)), 2.52–2.54 (s, 6H, –(CH₃)₂), 3.21 (s, 3H, solvent CH₃OH), 4.02–4.13 (m, 4H, –(CH₂Me)₂), 6.75 (s, 1H, –CH–), 7.41 (s, 2H, –C₅H₄N), 8.61 (s, 2H, –C₅H₄N), 11.45 (s, 1H, –NH). ¹³C-NMR (100 MHz: DMSO-d₆ and CDCl₃: Me₄Si) δ (ppm): 164.3, 150.1, 149.0, 148.4, 147.7, 122.3, 96.3, 59.8, 57.7, 48.6 (solvent CH₃OH), 18.5, 13.7. UV-Vis [(λ_max (nm) (ε, dm³ mol⁻¹ cm⁻¹))] in CH₃CN: 290 (11 900). ESI-MS (m/z): 782.1 ([Zn(L³)₂Cl₂: 2CH₃OH: 1/2H₂O]⁺; 60%), 673 ([Zn(L³)₂Cl]⁺; 40%), 637 ([Zn(L³)₂]⁺; 70%).

Cell culture

Human lung (A549), cervical (HeLa) and colon (HCT-15) cancer cell lines and human embryonic kidney (HEK) non-cancer cell lines were obtained from the National Centre for Cell Science (NCCS), Pune, India. The lung (A549), cervical (HeLa), colon (HCT-15) and HEK cells were maintained in Dulbecco's Modified Eagle's medium (DMEM) while the colon (HCT-15) was maintained in Roswell Park Memorial Institute Medium (RPMI) supplemented with 10% fetal bovine serum (FBS), 10 mg mL⁻¹ of penicillin, 10 mg mL⁻¹ of streptomycin, and 0.25 mg mL⁻¹ of amphotericin B at 37 °C and kept in an incubator under 5% CO₂.

MTT assay

The cytotoxicity or survival of cells in the presence or absence of the experimental agents was determined using the MTT assay as described previously by Mosmann.⁶¹ Briefly, human lung (A549), cervical (HeLa) and colon (HCT-15) cancer and HEK non-cancer cells were seeded at the density of 6000 cells per well in 96 well plates for 24 h, in 200 μL of DMEM supplemented with 10% FBS. Different concentrations of complexes (100–1000 nM) were treated in triplicates and incubated for 48 h at 37 °C under 5% CO₂ in an incubator. After treatment, cells were incubated with MTT (10 μL; 5 mg mL⁻¹) at 37 °C for 3 h and the formazan crystals which formed were dissolved in 80 μL of dimethyl sulfoxide (DMSO). The plates were read at 590 nm on a scanning multiwell spectrophotometer and the IC₅₀ (concentration of compounds found to kill 50% of the cells) values were obtained using dose–response curves using GraphPad Prism software. Graphs of percentage cell inhibition versus concentration were plotted using Origin 6.0 software. The statistical significance of the data was analysed by ANOVA with the level of significance, p < 0.05. To clarify any participating role of DMSO in the cancer and non-cancer cell lines, separate studies were carried out with the solution of DMSO and they showed no activity against any of the cancer and non-cancer cell lines.

Apoptosis assessment by live cell imaging and acridine orange-ethidium bromide staining

For live cell imaging, the cancer cell lines were treated with the appropriate IC₅₀ concentrations of copper(II) complexes (2 and 5) for 48 h and the morphology of cells was observed under an inverted phase contrast microscope. For fluorescent imaging, after treatment, the cancer cells were stained with freshly prepared acridine orange – EB staining solution (100 μg mL⁻¹) and observed under a Olympus BX-51 fluorescent microscope within 20 min.

X-ray crystallography

XRD studies for ligands (L^1&2) and complex 6 were carried out using a Bruker AXS Kappa APEX II single crystal charge-coupled detection diffractometer equipped with graphite-monochromated Mo (Kα) (λ = 0.71073 Å) radiation. The diffractometer has a four circle goniometer with φ, χ, ω and 2θ axes by which the crystal is rotated. The intensity data were collected using ω and φ scans with a frame width of 1°. For all the compounds, data collection and data reduction was done using APEX2, SAINT/XPREP software (version 7.06a). The multi-scan absorption correction was done using the SADABS program.⁶² The structures were solved by direct methods using SHELXS97 and refined using SHELXL97 by the methods of full-matrix least squares refinement.⁶³ The molecular graphics were done using ORTEP-3.⁶⁴

References

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Footnote

† Electronic supplementary information (ESI) available: Table S1 shows crystal data and structure refinement for ligands L^1&2 and complex 6, Tables S2–S5 show the data of bond length, bond angles and hydrogen bond parameters for ligands L^1&2, Fig. S1–S4 show crystal packing and hydrogen bonding parameters of ligands L^1&2, Fig. S5–S8 show FT-IR spectra of ligands L^1&2 and complexes 1 and 4. Fig. S9–S16 show ¹H-NMR and ¹³C-NMR spectra of ligands L^1&2 and complexes 3 and 6. Fig. S17 shows UV-Vis spectra of complexes 1 and 2 and Fig. S18 shows an X-band EPR spectrum of complex 2. CCDC 881456, 881461 and 873831. For ESI and crystallographic data in CIF or other electronic format see DOI: 10.1039/c4ra11197d

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