Bimodal sensor employing a novel approach for simultaneous selective detection of Ni²⁺ and biomolecules via turn-on fluorescence supported by DFT and molecular docking

Abstract

A bimodal sensor, (E)-2-(4-(diphenylamino)styryl)-1-methylquinolin-1-ium (DSM), was designed and synthesized for the simultaneous fluorescence turn-on detection of Ni²⁺ ion and biomolecules such as ct-DNA, BSA, and ovalbumin. Due to its distinct size and steric properties, DSM exhibits different binding modes when interacting with Ni²⁺ and DNA/proteins. The probe DSM possesses dual functionalities, allowing it to selectively detect Ni²⁺ at one binding site while interacting with ct-DNA, BSA, and ovalbumin at another. Thus, interactions of DSM with Ni²⁺ result in fluorescence enhancement at 377 nm and 400 nm, with a detection limit of 1.53 μM and binding constant of 1.2 × 10⁶ M⁻¹. Moreover, the binding of DSM with Ni²⁺ has been demonstrated via UV-vis, mass spectra, Jobs plots and DFT analysis. Conversely, binding of DSM with ct-DNA, ovalbumin and BSA led to an increase in the fluorescence at 425 nm and 435 nm, respectively, with the detection limit at micromolar (ct-DNA) and nanomolar (BSA and ovalbumin) levels. These interactions have been validated through UV-vis spectroscopy, fluorescence studies, and molecular docking analysis. Thus, this study underscores the potential of DSM as a versatile tool for simultaneous detection of both metal ions and biomolecules with a unique bimodal approach.

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1. Introduction

The development of various chemosensors to detect different metal ions has garnered notable attention due to their extensive relevance in medical, biological, and industrial applications.^1–3 To date, many researchers have investigated various analytical techniques for detecting different metal ions and anions, including atomic absorption spectroscopy, surface-enhanced Raman scattering, ion-selective electrodes, and inductively coupled plasma mass spectrometry.⁴ However, these techniques often involve complex procedures, demand skilled operators, and incur significant costs.⁵ Therefore, among the various analytical techniques, colorimetric and fluorescent assays are highly favoured as optical methods, as they provide numerous benefits, such as rapid response and high selectivity and sensitivity. Consequently, for the purpose of detecting different metal ions, a multitude of colorimetric and fluorescent sensors have been created.^6–10

Across various metal ions, nickel is a vital metal nutrient essential for sustaining human life through different functions involving several nickel-containing enzymes and coenzymes, such as urease, Ni–Fe hydrogenases, and F430.¹¹ Meanwhile, excess nickel accumulation can adversely affect the respiratory and immune systems, although the exact mechanism of nickel imbalance has not yet been identified. Selective fluorescent probes can be used as cell-imaging reagents to detect trace levels of Ni²⁺ in living systems.^12,13 Selective detection of Ni²⁺ is extremely important, but only a few Ni²⁺-selective peptide,^14,15 protein,¹⁶ polymer^17,18 and fluorescent sensors have been developed so far.^19–22 The development of chemosensors for selective detection of Ni²⁺ is challenging.

Current research is also focusing on the interactions of ligands with proteins and nucleic acids, as these molecules serve as a model for drug design and cancer treatment.²³ Thus, suitable dye molecules that bind to DNA and protein can be used for the detection of nucleic acids or proteins.²⁴ The binding interactions of the dye with nucleic acid can be easily understood by the change in the absorption or emission spectra. The relatively low demands and the high sensitivity of the equipment aid in promoting the fluorometric detection of biomacromolecules as the key method. In this regard, a variety of fluorescent probes are particularly helpful (“light-up probes”) since their emission intensity dramatically increases when they form complexes with proteins or DNA.²⁵

In this respect, various fluorescent probes whose emission intensity increases significantly upon complex formation with DNA or proteins are especially useful (“light-up probes”). This light up fluorescence technique is commonly applied in biology, biochemistry and also in medicine.²⁶ Along with the biomacromolecules, several small molecules turn out to be essential within cells, and their concentration in the cellular medium determine function or malfunction of the physiological processes. Thus, we herein developed a multifunctional styryl dye, (E)-2-(4-(diphenylamino)styryl)-1-methylquinolin-1-ium (DSM), for selective detection of the metal ion Ni²⁺ and biomacromolecules like ct-DNA, BSA and ovalbumin. In this scenario, it has been demonstrated that styryl dyes with a cationic hetarene unit with donor-acceptor sites exhibit favourable properties for DNA sensing.^27–29 Additionally, we have shown that donor (triphenyl amine)-substituted- with acceptor (quinolinium) derivatives DSM exhibit favorable photophysical properties, making them suitable as versatile building blocks for fluorescent light-up probes for DNA or protein.^30,31 Although there are reports for the detection of Ni²⁺ and DNA/proteins by developing individual sensors, the simultaneous detection of Ni²⁺ and DNA/proteins by designing a single multifunctional chemosensor like DSM is rare (Table S1†). Therefore, here we have combined two types of sensing phenomena in DSM that associates with DNA/proteins through the quaternary quinolinium motif and Ni²⁺ through the triphenyl amine part (Scheme 1).

Scheme 1 (a) Simplified cartoon diagram demonstrating the working principle of a dual-mode chemosensor in the present work compared with previous reports. The colored circles represents the fluorescence light-up effect of the molecule. (b) Design of a bimodal senor for simultaneous detection of metal and DNA/protein.

Herein, we have developed a multifunctional sensor (E)-2-(4-(diphenylamino)styryl)-1-methylquinolin-1-ium (DSM) using Knoevenagel-condensation³² of 1,2-dimethylquinolin-1-ium (1) with 4-(diphenylamino)benzaldehyde in 66% yield (Scheme 2). The chemical structure of DSM was confirmed using ¹H-NMR, ¹³C-NMR, and mass spectrometry (Fig. S9–S11, ESI†).

Scheme 2 (a) Methyl iodide and ethanol refluxed at 50 °C, 48 h. (b) Piperidine and ethanol refluxed at 70 °C, overnight.

2. Experimental

2.1 Materials and instrumentation

General. Chemicals, solvents including buffer solution and biomolecules like ct-DNA, BSA and ovalbumin were purchased from Sigma Aldrich. Melting points were determined by utilizing a hot-plate melting point equipment with an open-mouth capillary. On a Bruker Avance 400 MHz instrument, ¹H-NMR and ¹³C-NMR spectra were recorded in DMSO-d₆ solvent. The ¹H–¹H coupling constants are mentioned in Hz, and chemical shifts are specified in δ-units. UV-vis and fluorescence titration experiments were performed using a UV-spectrophotometer: PerkinElmer, Lambda 30 and a Shimadzu spectrofluorophotometer RF-6000 using a fluorescence cell of 10 mm path, respectively.

2.2 Synthesis and characterization of DSM

Synthesis of 1,2-dimethyl quinolinium iodide was carried out using the reported literature procedure.³³ 2-Methyl quinaldine (0.725 g, 4.5 mmol) was dissolved in ethanol and methyl iodide (9.3 mmol) was added to it. The reaction mixture was then heated to reflux for 48 hours at 80 °C. A yellow-colored precipitate was formed and the precipitate was filtered and dried for the next step without further purification. 1,2-Dimethyl quinolinium iodide (0.208 g, 0.7 mmol) and 4-diphenyl amino benzaldehyde (0.2 g, 7.31 mmol) were dissolved in ethanol (10 ml) and a catalytic amount of piperidine was added to the reaction mixture, which was further refluxed for 24 hours. Reaction progress was monitored by TLC and a red precipitate was formed by the addition of diethyl ether to the reaction mixture. The red precipitate was further purified via column chromatography using 2–3% methanol in chloroform as the eluent to obtain a pure red product.

Yield: 200 mg, 66%. Mp-135–140 °C. ¹H NMR (DMSO-d₆, 400 MHz): δ (ppm): 8.51 (d, 1H, J = 9.2 Hz), 8.33 (d, 2H, J = 8 Hz), 8.21 (d, 1H, J = 15.6 Hz), 8.16 (m, 2H, J = 15.6 Hz), 7.89 (m, 3H), 7.71 (d, 1H, J = 15.6 Hz), 7.42 (t, 4H, J = 7.6 Hz), 7.2 (q, 6H), 6.96 (d, 2H, J = 8.4 Hz), 4.51 (s, 3H). ¹³C NMR (DMSO-d₆, 100 MHz): δ (ppm): 156.6, 150.9, 147.6, 146.3, 143.6, 139.6, 135.1, 131.6, 130.4, 129.1, 127.9, 127.8, 126.3, 125.5, 121.2, 120.2, 119.6, 116.3. Mass (MS): M⁺ calculated for C₃₀H₂₅N₂⁺ is 413.201; found: 413.250.

3. Results and discussion

3.1 Binding study with Ni²⁺

The emission properties of DSM were studied by adding different interfering metal cations like Cu²⁺, Hg²⁺, Al³⁺, Ni²⁺, Mn²⁺, Zn²⁺, Co²⁺, Cd²⁺, Cr³⁺, Fe²⁺, Fe³⁺, and Pb²⁺ (c = 2 × 10⁻⁴ M) in CH₃CN-HEPES buffer (7 : 3, v/v, pH = 7.4). Initially, DSM exhibits two absorption peaks at 331 nm and 497 nm, and results a gradual decrease of absorbance with a moderate shift of wavelength with the incremental addition of Ni²⁺ (Fig. 1a). For DSM and DSM–Ni²⁺, the molar absorption coefficients at 497 nm were determined to be 5.3 × 10⁴ liter mol⁻¹ cm⁻¹ and 2.5 × 10⁴ liter mol⁻¹ cm⁻¹, respectively.

Fig. 1 (a) and (c) UV-vis and fluorescence titration of DSM (c = 2 × 10⁻⁵ M) upon the addition of Ni²⁺ (2 × 10⁻⁴ M, 10 equiv.) respectively. (b) and (d) changes in concentration vs. intensity of DSM for Ni²⁺ in UV-vis (λ_em = 497 nm) and fluorescence spectra (λ_em = 377 nm) respectively.

In the emission experiment, the ligand DSM itself exhibits a weak emission signal at 400 nm (λ_ex = 331 nm) with a quantum yield (Φ = 0.19). With further increase in the concentration of Ni²⁺, the emission intensity of DSM was notably enhanced by 6-fold by the generation of another blue shifted emission peak at 377 nm (Δλ = 23 nm) along with the ligand signal at 400 nm with a high quantum yield (Φ = 0.76) exhibiting naked eye fluorescence colour change from light orange to blue (Fig. 1c). The decrease of the absorbance and increase of the emission intensity of DSM varied up to the saturation level with increasing concentrations of Ni²⁺ (Fig. 1b and d).

3.2 Interference study

Fluorescence titration experiments were performed to evaluate the selective binding affinity of DSM towards Ni²⁺ in the presence of various metal ions such as Cu²⁺, Hg²⁺, Al³⁺, Ni²⁺, Mn²⁺, Co²⁺, Zn²⁺, Cd²⁺, Cr³⁺, Fe²⁺, Fe³⁺, and Pb²⁺ in CH₃CN–aqueous HEPES buffer (7/3, v/v, pH = 7.4). It is noted that the presence of Ni²⁺ only led to a detectable increase in the fluorescence intensity and no other significant equivalent emission enhancement was observed in the presence of other interfering metal ions (Fig. 2a and b). The bar diagram represents the fluorescence behaviour of DSM towards various metal ions with a remarkable selectivity towards Ni²⁺. The blue bar with the highest intensity indicates the fluorescence enhancement of DSM in the presence of Ni²⁺ and orange bars represent no considerable binding of DSM with other metal ions (Fig. 2c).

Fig. 2 (a) Fluorescence colour behaviour of probe DSM with various metal ions. b) Emission spectra of DSM upon the addition of different metal ions (10 equiv.) in CH₃CN–aqueous HEPES buffer solution (7/3, v/v, pH = 7.4) (λ_em = 377 nm). (c) Changes in the emission intensity of the sensor DSM with the addition of 10 equiv. of each interfering cation (c = 2 × 10⁻⁴ M, 10 equiv.).

3.3 Competition study

Fluorometric analysis was used to investigate the effect of competing metal ions on DSM binding to Ni²⁺ to better analyse the selectivity of DSM for Ni²⁺, where 10 equivalents of Ni²⁺ and other metal ions were used in a cross-contamination study. Notably, the fluorescence enhancement for Ni²⁺ binding remained unperturbed (red bars), and the other interfering metal ions did not affect the sensing of Ni²⁺ (black bars). This indicates that the receptor DSM is highly selective and sensitive for Ni²⁺ (Fig. 3).

Fig. 3 Change in fluorescence intensity of DSM (c = 2 × 10⁻⁵ M) on the addition of 10 equiv. of various metal ions (c = 2 × 10⁻⁴ M) (green bars); change in fluorescence with the addition of 10 equiv. of various other metal ions in the presence of 10 equiv. of Ni²⁺ at 377 nm (grey bars).

The detection limit of DSM for Ni²⁺ was calculated as 1.53 μM based on the fluorescence titration studies using the formula DL = K × Sb₁/S, where K = 3, Sb₁ is the standard deviation of the blank solution, and S is the slope of the calibration curve (Fig. S4†).³⁴ The binding stoichiometry was demonstrated through Job's plot analysis showing 1 : 1 binding of DSM with Ni²⁺ (Fig. S8†) and the association constant (K_a) of DSM with Ni²⁺ was calculated as 1.2 × 10⁶ M⁻¹ (error <10%) from the fluorescence titration experiment (Fig. S2†).³⁵

3.4 Reversibility test

To further explore the binding nature of DSM with Ni²⁺, a fluorescence titration experiment was performed. By adding excess Na₂EDTA (0–2.0 equivalents) to the DSM–Ni²⁺ solution, the fluorescence intensity was gradually decreased due to the removal of Ni²⁺ from the metal binding chamber of DSM by the strong binding of Na₂EDTA with Ni²⁺ (Fig. 4). The experiment was repeated for several cycles with alternate additions of Ni²⁺ and Na₂EDTA (Fig. S14†). A systematic switching on–off pattern was observed between DSM and DSM with Ni²⁺ followed by the addition of EDTA. Thus, the reversible binding pattern of Ni²⁺ with DSM has been demonstrated.

Fig. 4 Fluorescence changes of DSM + Ni ²⁺ complex solution (c = 2 × 10⁻⁵ M) upon the addition EDTA (c = 2 × 10⁻⁴ M).

3.5 Dipstick method

To investigate the practical application of sensor DSM towards the sensing of Ni²⁺, test strips were prepared by immersing the TLC plates into the receptor solution (c = 2 × 10⁻⁵ M) in CH₃CN–H₂O (7 : 3, v/v). When DSM–Ni²⁺ and DSM-coated test strips were exposed to UV light, there was no significant fluorescence observed for the DSM-coated test strip (Fig. S13,† left), but the DSM–Ni²⁺-treated test kit exhibited a noticeable turn-on blue fluorescence, which was also observed in the solution phase (Fig. S13,† right). This demonstrates that the test strips, also referred to as dipsticks, selectively detect Ni²⁺ by enhancement of fluorescence which can also be detectable with the naked eye in solid phase. Such dipsticks provide instant qualitative results, eliminating the need for complex instrumental analysis.

3.6 Probable binding mode in solution phase

Initially, DSM alone exhibited weak fluorescence, but a notable fluorescence enhancement was observed at 400 nm in the presence of Ni²⁺. Since DSM is a donor–acceptor type of molecule, there is no significant fluorescence in the absence of Ni²⁺ due to the highly efficient PET process from the donor triphenyl amine moiety to acceptor quinolinium backbone (Scheme 3) (off-state). Significantly, the lone pair of electrons on the nitrogen of the triphenyl amine moiety coordinates with Ni²⁺, which consequently suppresses the PET process and operates through the chelation enhanced fluorescence (CHEF) effect, which results in notable fluorescence enhancement (on-state).³⁶ The emission intensity of DSM is notably weak (Φ = 0.01), which is likely due to radiationless deactivation of the excited state caused by the isomerization process due to free rotation around the C=C bond, a general phenomenon commonly observed in styryl dyes. Due to co-ordination of Ni²⁺ with DSM, C=C bond isomerization was inhibited owing to steric hindrance, which resulted in increased fluorescence intensity of the ligand.³⁷ This observation can be verified by blocking C=C bond rotation by increasing the viscosity of the solution due to the inhibition of rotational relaxation of the excited molecule (Fig. S1, ESI†).³⁸ Notably, in highly viscous media with a high proportion of glycerol in methanol, a noticeable rise in fluorescence intensity was observed at 461 nm. Due to the suppression of twisting intramolecular charge transfer (TICT) brought on by conformational changes from rotation around the C=C bond, geometrical rearrangement in the excited state may slow down under high viscosity conditions, leading to enhanced fluorescence. The formation of a complex between DSM and Ni²⁺ has been verified through the emergence of a mass peak at m/z = 582.950, attributed to [DSM + Ni²⁺ + 2Cl⁻ + CH₃CN] (Fig. S12†), which has also been justified by a 1 : 1 binding stoichiometric ratio in the Jobs plot analysis and theoretical calculation.

Scheme 3 Probable binding mode of DSM with Ni²⁺ in solution phase.

3.7 Theoretical calculation for Ni²⁺ binding study

The theoretical calculation has been demonstrated through optimization of the DSM and DSM–Ni²⁺ structures through DFT and TD-DFT calculations. In the optimized structure of the DSM–Ni²⁺ complex, Ni²⁺ shows tetrahedral coordination with the charged ligand DSM employing four co-ordinations, consisting of one nitrogen from the triphenyl amine group of DSM, two from counter anions Cl⁻ and one from the nitrogen of the acetonitrile solvent. From the optimized structure of the DSM–Ni²⁺ complex, the calculated Ni–N (triphenyl amine and acetonitrile nitrogen) bond distances are 1.916 Å and 1.839 Å, respectively, and the Ni–Cl bond distance is 2.171–2.191 Å. Meanwhile, the bond angles of nitrogen and chlorine with Ni are estimated as 176.86° and 175.23°, respectively. Additionally, the HOMO–LUMO energy gap is found to be ΔE = 6.82 eV and 4.97 eV for DSM and the DSM–Ni²⁺ complex, respectively, indicating their contribution to the stabilization of complex formation (Fig. 5c). Based on the DFT-optimized structures, the total energies of DSM alone and the DSM–Ni²⁺ complex are 136.55 kcal mol⁻¹ and 225.59 kcal mol⁻¹, respectively. Therefore, the energy difference between the DSM–Ni²⁺ complex and DSM alone is calculated to be 89.04 kcal mol⁻¹. On the other hand, the changes in the absorbance of DSM occur upon binding with Ni²⁺ at 330 nm and 490 nm. From the theoretical study, the corresponding estimated absorption bands have been calculated at 329 nm (E = 4.760, f = 0.0079 nm) and 498 nm (E = 3.489, f = 0.1609). For the transition from S0 → S15 and S0 → S2, the experimental data are in good agreement with the theoretical computations; hence, the experimental data align well with the theoretical calculations.

Fig. 5 Geometrically optimized molecular structures of (a) DSM and (b) DSM–Ni²⁺ complex. (c) Frontier molecular orbital with the energy difference of DSM and the DSM–Ni²⁺ complex.

3.8 Biological applications

3.8.1 DNA binding study. Utilizing the UV-vis and fluorescence spectroscopy, the relative DNA binding characteristics with DSM were examined in Tris-HCl buffer at pH 7.2. Initially, DSM itself showed strong absorption signals at 330 nm and 490 nm. The increase in concentration of ct-DNA to DSM solution resulted in gradual decrease in absorbance at 330 nm and 490 nm (Fig. 6a). In the fluorescence spectra, DSM itself exhibited weak emission at 425 nm with low quantum yield (Φ = 0.01) (λ_ex = 330 nm) but progressive addition of ct-DNA to DSM caused a gradual increase in the fluorescence intensity at 425 nm by 5-fold with high quantum yield (Φ = 0.02) (Fig. 6c). The spectra undeniably depict the fluorescence turn on behaviour of DSM on the addition of ct-DNA. From UV-vis and fluorescence studies, the detection limits of DSM for ct-DNA were calculated as 2.79 μM and 1.01 μM, respectively (Fig. S5†). The binding constant (K_b) of 1.2 × 10⁴ M⁻¹ was determined using the non-linear binding isotherm of DSM towards ct-DNA obtained from the fluorometric titrations (Fig. S3a†). The binding study of DSM with ct-DNA conducted through photometric and fluorometric experiments unequivocally verifies the binding of these probes with DNA. First, UV-vis titrations revealed a moderate red shift of the absorption band with progressive titration with DNA.³⁹ The ligand DSM has very low quantum yields (Φ = 0.01) with mild fluorescence when DNA was not present in aqueous buffer solution due to radiationless deactivation by conformational relaxation of the excited state. The association with ct-DNA results in the enhancement of emission intensity (Fig. 6c). It has been shown that several DNA sensitive probes exhibit fluorescent light-up effects due to the result of the restricted conformational freedom of movement of the ligand within the binding site,³⁷ which is also comparable with the fluorescence enhancement observed in highly viscous media like glycerol.³⁸ Therefore, we propose that the emission enhancement of DSM in the presence of DNA is the result of restricted conformational flexibility that reduces the radiationless de-activation of the excited state, which has been demonstrated in a similar study using several styryl- and stilbene-type dyes.^27–29 The steric protection and delocalization of the amino group's lone pair limit its availability, making ligand DSM association primarily driven by attractive dispersion interactions like π stacking and van der Waals forces, along with the release of the thermodynamically favorable counter-ion from DNA. Additionally, the structural flexibility of DSM, with its hydrophobic triphenylamine and DNA-binding quinolinium backbone, may cause its bulky diaryl unit to point outside the binding site when the charged quinolinium moiety intercalates into DNA.³⁹

Fig. 6 (a) and (c) UV-vis and fluorescence titration of DSM (c = 2.0 × 10⁻⁵ M) upon incremental addition of ct-DNA (c = 2 mM in base pairs) in Tris-HCl buffer, pH 7.2. (b) and (d) Changes in concentration vs. intensity in UV-vis (λ_em = 490 nm) and fluorescence spectra (λ_em = 425 nm).

3.8.2 Ethidium bromide (EB) competition assay. To further investigate the interaction mode between ct-DNA and DSM, the quenching assay was employed mainly by the displacement of the intercalating dye (EB) from ct-DNA. When EB is intercalated into the base pair of ct-DNA, it shows a significant increase in the fluorescence intensity. However, the incorporation of a second ligand which can substitute the ct-DNA bound EB or break the secondary structure of DNA results in a decrease of the fluorescence intensity.⁴⁰

Herein, the EB replacement assay has been performed by the fluorescence experiment in Tris-HCl buffer, pH = 7.2. Initially, high fluorescence intensity has been observed for EB bound ct-DNA at 627 nm, which was significantly quenched with the increase in concentrations of the compound DSM. This fluorescence quenching is mainly due to the replacement of EB from EB bound ct-DNA complex by DSM. This confirms the intercalation binding mode of DSM with ct-DNA.⁴¹

By using the linear Stern–Volmer equation, the quenching of fluorescence in ct-DNA bound EB can be well understood, in which DSM is the quencher:⁴²

I₀/I = 1 + K_svQ

where I₀ and I indicate the fluorescence intensities in the presence and absence of the compound DSM (quencher), respectively; K_SV is the linear Stern–Volmer quenching constant; and Q is the concentration of DSM. The ratio of the slope to the intercept gives the K_SV value. The K_SV value was found to be 6.3 × 10³ M⁻¹. L, which explains the high quenching efficiency of DSM (Fig. 7).

Fig. 7 (a) Fluorescent changes that occur when the ct-DNA–ethidium bromide system is titrated with DSM (c = 2 × 10⁻⁵ M) in Tris-HCl buffer, pH = 7.2. (b) Plot of I₀/I vs. [DSM] for the titration of the compound to the ct-DNA–EB complex.

3.8.3 Protein binding study. Using UV-vis and fluorescence investigations, the binding capacity of DSM with bovine serum albumin (BSA) and ovalbumin were examined in Tris-HCl buffer at pH 7.2.

Initially, DSM exhibits strong absorption signals at 330 nm and 490 nm. However, increasing concentrations of BSA and ovalbumin results in significant decreases in the corresponding absorption signals (Fig. 8a and 9a). In the fluorescence experiment, DSM exhibited weak emission signal in the absence of BSA and ovalbumin at 425 nm with very low quantum yield (Φ₀ = 0.01). However, incremental increases in the concentration of BSA and ovalbumin to the DSM solution resulted in the enhancement of fluorescence intensities by 3-fold at 435 nm and 425 nm with comparatively high fluorescence quantum yields at Φ = 0.02 and Φ = 0.13, respectively (Fig. 8c and 9c). The detection limits of DSM for BSA and ovalbumin calculated from UV-vis and fluorescence measurements are 7 nM, 5 nM and 17 nM, 5 nM, respectively (Fig. S6 and S7†). The binding constants of DSM with BSA and ovalbumin obtained from non-linear fitting curves from spectrofluorometric titrations are 1.2 × 10⁴ M⁻¹ and 5.4 × 10⁵ M⁻¹, respectively (Fig. S3b and c†).

Fig. 8 (a) and (c) UV-vis and fluorescence of DSM (c = 2.0 × 10⁻⁵ M) upon titration with BSA (c = 7.4 μM) in Tris-HCl buffer at pH 7.2 respectively. (b) and (d) Changes in concentration vs. intensity in UV-vis (λ_em = 490 nm) and fluorescence spectra (λ_em = 425 nm) respectively.

Fig. 9 (a) and (c) UV-vis and fluorescence of DSM (c = 2.0 × 10⁻⁵ M) upon titration with ovalbumin (c = 4.24 μM) in Tris-HCl buffer, with pH = 7.2. (b) and (d) Changes in concentration vs. intensity in UV-vis (λ_em = 490 nm) and fluorescence spectra (λ_em = 425 nm).

The same mechanism that explains the fluorescence light-up effect in the presence of proteins also explains the association of DSM with nucleic acids. This mechanism involves a combination of restricted conformational freedom of the molecule and suppressed formation of an ICT or CS excited state within the binding site. The variation of absorbance and the fluorescence response of DSM on binding with DNA and proteins is due to the different polarity of the binding cavities and the CS or ICT state is stabilized more proficiently inside the more polar DNA binding cavity compared with the less polar proteins (BSA and ovalbumin) binding pocket.^43,44 The comparatively significant light-up effect in the presence of proteins resulted in a decrease of the CS or ICT state within the binding site.

3.8.4 In silico molecular docking studies. The binding interactions of DSM with BSA protein and ct-DNA were further supported by molecular docking studies to predict the binding affinity and sites of DSM with biomolecules such as proteins and ct-DNA. The analysis was conducted using Autodock Vina technology (ESI). In silico molecular docking is a powerful tool that complements experimental techniques, enhancing the understanding and development of fluorescent ligands for DNA and protein binding. Thus, the molecular docking result showed that the selected ligand DSM could bind strongly with the target protein bovine serum albumin (BSA) (binding energy: −8.6 kcal mol⁻¹) and the DNA molecule (binding energy: −8.5 kcal mol⁻¹). The interaction between the protein and DSM was stabilized through SER104, ASP107, ASP108, TYR147, CYS199, ALA200, VAL461, and LYS465 (Fig. 10a1–c1). Furthermore, the ligand DSM formed a complex with the DNA through an intercalation mode and interacted with deoxyribose cytosine (817) and thymine residues (906) (Fig. 10a2–c2), which was also proven by the ethidium bromide (EB) competition assay.

Fig. 10 Possible binding mechanisms of DSM with BSA (1), and ct-DNA (2). Three-dimensional surface (a1 and b2) and ribbon (b1 and b2) views of the complex of DSM with BSA and ct-DNA, respectively; c1 and c2 represent two-dimensional interaction of BSA and ct-DNA with DSM, respectively.

4. Conclusion

The multifunctional sensor DSM was successfully designed and synthesized for the simultaneous fluorescence turn-on detection of metal ion Ni²⁺ and biomolecules such as ct-DNA, BSA, and ovalbumin. The unique structural characteristics of DSM allow it to exhibit different binding modes due to its distinct size and steric demands. The bimodal functionalities of DSM enable selective detection of Ni²⁺ at one binding site while interacting with ct-DNA, BSA, and ovalbumin at another. The interaction of DSM with Ni²⁺ results in fluorescence enhancement, with a specific detection limit and binding constant. Similarly, binding investigations with ct-DNA, BSA, and ovalbumin also show increased fluorescence with the detection limit at the micromolar (ct-DNA) and nanomolar (BSA and ovalbumin) levels. The binding of DSM with Ni²⁺ was confirmed through mass spectrometry, Job's plots, and DFT analysis, while the binding interactions with ct-DNA, BSA, and ovalbumin were demonstrated using UV-vis, fluorescence, and docking studies. This study highlights the potential of DSM as a versatile probe for detecting both metal ions and biomolecules.

Data availability

The data supporting this article have been included as part of the ESI.†

Conflicts of interest

There are no conflicts of interest to declare.

Acknowledgements

The authors would like to thank Christ University, Bengaluru, for the research facilities and Centre for Research, Christ University, for the seed money grant (grant approval number CU-ORS-SM-24/09). Avijit Kumar Das specially acknowledges State University Research Excellence (SERB-SURE) of the Science and Engineering Research Board (SERB) (File Number: SUR/2022/002461) under the Department of Science and Technology, Government of India, for financial support by the research grant and research fellowship for Hazeena Shinziya.

References

1. V. Raju, R. SelvaKumar, S. K. AshokKumar, Y. Tharakeswar and S. K. Sahoo, Inorg. Chem. Commun., 2019, 101, 74–80.
2. M. Yang, L. Ma, J. Li and L. Kang, RSC Adv., 2019, 9, 16812–16818.
3. W. Sik Na, P. Raj, N. Singh and D. O. Jang, Tetrahedron Lett., 2019, 39, 151075.
4. C. Gao, X. Liu, X. Jin, J. Wu, Y. Xie, W. Liu, X. Yao and Y. Tang, Sens. Actuators, B, 2013, 185, 125–131.
5. H. H. Hammud, S. E. Shazly, G. Sonji, N. Sonji and K. H. Bouhadir, Spectrochim. Acta, Part A, 2015, 150, 94–103.
6. (a) D. Maity, D. Karthigeyan, T. K. Kundu and T. Govindaraju, Sens. Actuators, B, 2013, 176, 831–837; (b) A. K. Das and S. Goswami, Sens. Actuators, B, 2017, 245, 1062–1125.
7. (a) W. Du, R. J. Liu, J. Fang, H. Gao, Y. W. Wang and Y. Peng, Tetrahedron, 2019, 75, 130477; (b) S. Goswami, S. Maity, A. C. Maity, A. k. Das, B. Pakhira, K. Khanra, N. Bhattacharyya and S. Sarkar, RSC Adv., 2015, 5, 5735–5740; (c) M. S. Kumar, A. K. Das, Y. Bylappa and A. Nag, RSC Adv., 2025, 15, 6708–6717.
8. (a) T. B. Wei, B. R. Yong, L. R. Dang, Y. M. Zhang, H. Yao and Q. Lin, Dyes Pigm., 2019, 171, 107707; (b) S. Vishnu, A. K. Das, Y. Bylappa, A. Nag and M. Dolai, Anal. Methods, 2024, 16, 8164–8178.
9. Z. Gu, H. Cheng, X. Shen, T. He, K. Jiang, H. Qiu, Q. Zhang and S. Yin, Spectrochim. Acta, Part A, 2018, 203, 315–323.
10. S. C. Lee and C. Kim, Anal. Sci., 2019, 35, 1189–1193.
11. A. Sigel, H. Sigel and R. K. O. Sigel, Nickel and Its Surprising Impact in Nature, John Wiley & Sons, 2007, vol. 2.
12. D. W. Domaille, E. L. Que and C. J. Chang, Nat. Chem. Biol., 2008, 4, 168–175.
13. E. L. Que, D. W. Domaille and C. J. Chang, Chem. Rev., 2008, 108, 1517–1549.
14. D. A. Pearce, G. K. Walkup and B. Imperiali, Bioorg. Med. Chem. Lett., 1998, 15, 1963–1968.
15. A. Torrado, G. K. Walkup and B. Imperiali, J. Am. Chem. Soc., 1998, 3, 609–610.
16. L. L. Salins, E. S. Goldsmith, M. C. Ensor and S. Daunert, Anal. Bioanal. Chem., 2002, 372, 174–180.
17. B. Wang, Y. Hu and Z. Su, React. Funct. Polym., 2008, 7, 1137–1143.
18. (a) B. Y. Wang, X. Y. Liu, Y. L. Hu and Z. X. Su, Polym. Int., 2009, 6, 703–709; (b) S. Goswami, S. Chakraborty, A. K. Das, A. Manna, A. Bhattacharyya, C. K. Quah and H.-K. Fun, RSC Adv., 2014, 4, 20922–20926.
19. (a) L. Fabbrizzi, M. Licchelli, P. Pallavicini, A. Perotti, A. Taglietti and D. Sacchi, Chem. – Eur. J., 1996, 21, 75–82; (b) R. K. Pathak, J. Dessingou and C. P. Rao, Anal. Chem., 2012, 84, 8294–8300.
20. C. Bargossi, M. C. Fiorini, M. Montalti, L. Prodi and N. Zaccheroni, Coord. Chem. Rev., 2000, 208, 17–32.
21. (a) L. J. Jiang, Q. H. Luo, Z. L. Wang, D. J. Liu, Z. Zhang and H. W. Hu, Polyhedron, 2001, 20, 2807–2812; (b) R. Joseph, B. Ramanujam, H. Pal and C. P. Rao, Tetrahedron Lett., 2008, 49, 6257–6261.
22. H. Wang, D. Wang, Q. Wang, X. Li and C. A. Schalley, Org. Biomol. Chem., 2010, 8, 1017–1026.
23. Molecular Aspects of Anticancer Drug–DNA Interactions, ed. S. Neidle and M. Waring, CRC, Boca Raton, FL, 1993.
24. A. Granzhan, H. Ihmels and K. Jäger, Chem. Commun., 2009, 1249–1251.
25. (a) G. Cosa, K. S. Focsaneanu, J. R. N. McLean, J. P. McNamee and J. C. Scaiano, Photochem. Photobiol., 2001, 73, 585–599; (b) H. Shinziya, R. S. Menon and A. K. Das, RSC Adv., 2024, 14, 30631–30646.
26. D. Monchaud and M. P. Teulade-Fichou, Org. Biomol. Chem., 2008, 4, 627–636.
27. (a) R. W. Dirks and H. J. Tanke, Chem. Biol., 2006, 13, 559–561; (b) C. V. Kumar, R. S. Turner and E. H. Asuncion, J. Photochem. Photobiol., A, 1993, 74, 231–238; (c) A. K. Das, S. I. Druzhinin, H. Ihmels, M. Müller and H. Schönherr, Chem. – Eur. J., 2019, 25, 12703–12707.
28. (a) V. B. Kovalska, D. V. Kryvorotenko, A. O. Balanda, M. Y. Losytskyy, V. P. Tokar and S. M. Yarmoluk, Dyes Pigm., 2005, 67, 47–54; (b) J.-S. Lee, Y. K. Kim, M. Vendrel and Y.-T. Chang, Mol. BioSyst., 2009, 5, 411–421; (c) D. V. Berdnikova, O. A. Fedorova, E. V. Tulyakova, H. Li, S. Kölsch and H. Ihmels, Photochem. Photobiol., 2015, 91, 723–731.
29. (a) Q. Li, Y. Kim, J. Namm, A. Kulkarni, G. R. Rosania, Y. H. Ahn and Y. T. Chang, Chem. Biol., 2006, 13, 615–623; (b) M. Q. Wang, S. Liu, C. P. Tang, A. Raza, S. Li, L. X. Gao, J. Sun and S. P. Guo, Dyes Pigm., 2017, 136, 78–84; (c) A. Manna and S. Chakravorti, J. Phys. Chem. B, 2012, 116, 5226–5233.
30. (a) R. W. Sinkeldam, N. J. Greco and Y. Tor, Chem. Rev., 2010, 110, 2579–2619; (b) T. Deligeorgiev, A. Vasilev, S. Kaloyanova and J. J. Vaquero, Color. Technol., 2010, 126, 55–80; (c) S. Vishnu, A. Nag and A. K. Das, Anal. Methods, 2024, 16, 5263–5271.
31. (a) H. Özhalici-Ünal, C. L. Pow, S. A. Marks, L. D. Jesper, G. L. Silva, N. I. Shank, E. W. Jones, J. M. Burnette, P. B. Berget and B. A. Armitage, J. Am. Chem. Soc., 2008, 130, 12620–12621; (b) A. K. Das, H. Ihmels and S. Kölsch, Photochem. Photobiol. Sci., 2019, 18, 1373.
32. L. F. Tietze and U. Beifuss, Angew. Chem., Int. Ed. Engl., 1993, 32, 131–163.
33. V. F. Traven, A. V. Manaev, A. Yu. Bochkov, T. A. Chibisova and I. V. Ivanov, Russ. Chem. Bull., 2012, 61, 1342–1362.
34. M. Shortreed, R. Kopelman, M. Kuhn and B. Hoyland, Anal. Chem., 1996, 8, 1414–1418.
35. (a) H. A. Benesi and J. H. Hildebrand, J. Am. Chem. Soc., 1949, 71, 2703–2707; (b) Y. Shiraishi, Y. Kohno and T. Hirai, Ind. Eng. Chem. Res., 2005, 44, 847–851.
36. H. Niu, J. Liu, H. M. O'Connor, T. Gunnlaugsson, T. D. James and H. Zhang, Chem. Soc. Rev., 2023, 52, 2322–2357.
37. R. N. Dsouza, U. Pischel and W. M. Nau, Chem. Rev., 2011, 111, 7941–7980.
38. M. A. Martin, M. Ballesteros and B. D. Castillo, Anal. Chim. Acta, 1985, 170, 95–100.
39. W. Sbliwa, G. Matusiak and B. Bachowska, Croat. Chem. Acta, 2006, 79, 513.
40. (a) J. H. Zhou, S. Q. Xia, J. R. Chen, X. S. Wang, B. W. Zhang, H. J. Zhang, P. Zou, X. Cheng Ai and J. P. Zhang, J. Photochem. Photobiol., A, 2004, 165, 143–147; (b) Q. Zhang, F. Zhang, W. Wang and X. Wang, J. Inorg. Biochem., 2006, 100, 1344–1352; (c) Y. F. Song and P. Yang, Polyhedron, 2001, 20, 501–506.
41. D. J. Patel, Acc. Chem. Res., 1979, 12, 118–125.
42. Y. Chen, M. Cai, Y. Zhang and W. Zheng, Study on The Mechanism of Chitosan Complex Formation with PEGFPC∼ 3 DNA, Pharmaceutical Biotechnology Beijing, 2005, vol. 5, p. 291.
43. A. Cuervo, P. D. Dans, J. L. Carrascosa and L. Fumagalli, Proc. Natl. Acad. Sci. U. S. A., 2014, 111, 3624–3630.
44. J. Eden, P. R. C. Gascoyne and R. Pethig, J. Chem. Soc., Faraday Trans., 1980, 76, 426–434.

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Footnote

† Electronic supplementary information (ESI) available. See DOI: https://doi.org/10.1039/d5sd00028a

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