Naphthoxazoles Derived from Lapachol as Fluorescent DNA Probes: Synthesis and Binding Studies with Calf Thymus DNA

Abstract

This study evaluated the potential of naphthoxazoles derived from lapachol as fluorescent deoxyribonucleic acid (DNA) probes. We synthesized three 2-substituted naphthoxazoles: one with a phenyl group (LOX1), and two with 2-hydroxyphenyl (LOX2) and 4-fluorophenyl (LOX3) groups, respectively. The compounds were synthesized using the Debus-Radziszewski reaction between the lapachol and aldehydes. The quantum yield (Φ_f) was determined using the classical relative method with 4’,6-diamidino-2-phenylindole (DAPI) as the standard, and the fluorescence lifetime (τ_f) was measured with a lifetime fluorescence spectrometer. The naphthoxazoles exhibited low fluorescence emission and consequently low values of Φ_f in their free form, demonstrating behavior similar to that of DAPI. These compounds displayed notably large Stokes shifts (approximately 340 nm) and the τ_f in the nanosecond range. DNA binding studies were conducted by monitoring changes in the absorption properties of naphthoxazoles in the absence and presence of calf thymus DNA at different concentrations. The results indicated that naphthoxazoles interacted with DNA through groove binding. LOX2 exhibited a binding constant (K_b) of 2.70 × 10⁴ M^-1, indicating strong binding to the biomolecule. These findings underscore the potential of LOX1, LOX2, and LOX3 as candidates for use as fluorescent DNA probes.

Keywords:
natural naphthoquinone; oxazole; DNA dye; groove binder; fluorescent marker

Introduction

Naphthoxazoles are fluorophores exhibiting promising photophysical characteristics, such as UV-Vis absorption and fluorescence emission, large Stokes shifts, increased fluorescence upon binding to biological targets, and high photostability.¹-⁴ Naphthoxazole derivatives result from substituting the oxazole ring at positions 4 and 5 with the naphthalene ring.⁵ These compounds can be obtained with high yields through one-pot synthesis protocols.¹,⁵

Fluorescent probes are extensively employed for deoxyribonucleic acid (DNA) studies.⁶,⁷ However, commonly used fluorescent DNA probes may be potentially toxic.⁸-¹⁰ Therefore, non-toxic and biocompatible organic fluorophores, with adjustable optical properties depending on the microenvironment, are valuable tools for probing interactions with DNA.¹¹

The literature⁵,¹²,¹³ reported the naphthoxazoles low toxicity; fluorophores derived from this class of compounds represent a sensitive and safe alternative to commercially available DNA fluorescent probes.

In this article, we present the synthesis, structural and photophysical characterization, and DNA interaction study of new naphthoxazole derivatives prepared from lapachol. We designed three 2-substituted naphthoxazoles derived from lapachol: one with a phenyl group (LOX1), and two with the 2-hydroxyphenyl (LOX2) and 4-fluorophenyl (LOX3) groups, respectively (Scheme 1).

Scheme 1
Synthesis reaction naphthoxazoles (LOX1, LOX2, and LOX3).

Experimental

General

In the synthesis experiments, reagents were purchased from Sigma-Aldrich (St. Louis, MO, USA) and used without preliminary purification, except lapachol, which was extracted from the wood of the yellow ipê (Tabebuia sp.). Thin-layer chromatography (TLC) (silica gel on aluminum chromatofoils with fluorescent indicator 254 nm) was purchased from Sigma-Aldrich (St. Louis, MO, USA) and was used to monitor the reaction progress. Column chromatography purification was carried out using silica gel pore size 60 Å, 230-400 mesh particle size were purchased from Sigma-Aldrich (St. Louis, MO, USA).

DNA sodium salt from calf thymus (CT-DNA type I, fibers form; Sigma-Aldrich, St. Louis, MO, USA) was employed in studies of interaction with the naphthoxazoles derivatives evaluated. A stock solution of CT-DNA was prepared by dissolving an appropriate amount of tris(hydroxymethylaminomethane)-HCl buffer (0.1 M, pH 7.4) and stored at 4 °C. The concentration of the stock solution of CT-DNA was determined by UV-Vis analysis using the Beer-Lambert equation with a molar absorptivity (ε) value of 6600 M^-1. To evaluate the purity of DNA against protein contamination, we used the ratio of absorbance at 260 and 280 nm (A₂₆₀/A₂₈₀); a value between 1.8 and 1.9 indicates that the DNA is protein-free.³,¹⁴,¹⁵

Instrumentation

All compounds were characterized by ¹H and ¹³C nuclear magnetic resonance (NMR) spectroscopy, infrared (IR) spectroscopy, and high-resolution mass spectrometry (HRMS). ¹H NMR and ¹³C NMR spectra were recorded on a Bruker Ascend^TM spectrometer (Zurich, Switzerland) (400 MHz for ¹H and 100 MHz for ¹³C) in deuterated dimethyl sulfoxide (DMSO-d₆). The chemical shifts (d) were reported in parts per million (ppm) and are referenced to either the residual solvent peak or tetramethylsilane (TMS) when possible. Coupling constants (J) were measured in hertz (Hz). Peaks are denoted as s (singlet), d (doublet), t (triplet) and m (multiplet). Fourier transform infrared (FTIR) spectra were acquired using the IRTracer-100 equipment (Shimadzu, Tokyo, Japan) with KBr pellets as the solid support. Scan were performed in the range of 4000 to 400 cm^-1 with a resolution of 8 cm^-1 and 45 scans. High-resolution mass spectrometry (HRMS) was obtained using a Q-TOF (ESI Q-TOF) mass spectrometer (Waters Synapt XS, Wilmslow, United Kingdom).

Absorption spectra were recorded using a Shimadzu UV 2600 absorption spectrometer (Tokyo, Japan) with 1.0 cm optical path length and covering the 200-800 nm region.

To determine the fluorescence quantum yield, samples were first evaluated using a UV-Vis spectrophotometer (Genesys 10S, Thermo Fisher Scientific, Madison, USA) within the 190-500 nm range, using a quartz cuvette to measure the maximum absorption wavelength with an absorbance below 0.1. The samples were then analyzed with a RF-6000 spectrofluorometer (Shimadzu Corporation, Nakagyo-ku, Kyoto, Japan), with excitation fixed at 345 nm and emission measured in the range of 640-800 nm, using slit opening of 10 nm for both excitation and emission, respectively.

To determine the fluorescence lifetime of the LOX1, LOX2, and LOX3 samples, a lifetime fluorescence spectrometer (EasyLife X, HORIBA Scientific, Sofia, Bulgaria) equipped with light emitting diode (LED) with excitation at 375 nm was used, following the protocol described in the equipment manual. All samples were prepared in DMSO.

Synthesis

Extraction and purification of lapachol

Lapachol was extracted from the heartwood of a Tabebuia genus and purified through recrystallization using ethanol.¹⁶ The heartwood of Tabebuia sp. was removed and cut into small, thin pieces (50 g). These pieces were then placed in 400 mL of a 1% (m/v) NaOH solution and left to stand for 24 h. The mixture was filtered, and the resulting dark red solution was acidified by slowly adding a 6.0 M HCl solution until a yellow solid precipitated. The solid was filtered and purified by recrystallization in hot ethanol. Lapachol was obtained as yellow crystals with a yield of 3.5% (m/m); melting point (mp) 138.3-140.3 °C.

General synthesis of the naphthoxazoles derived from lapachol (LOX1, LOX2, and LOX3)

The access was registered in the National System of Genetic Heritage and Associated Traditional Knowledge (SisGen) under the A5FDA89. A yellow solid was obtained with a yield of 1.5% (m/m) and mp 138.3-140.3 °C (lit.¹⁶ mp 138.3-140.3 °C).

To a solution of lapachol (1 mmol) in ethanol (6 mL), the appropriate aromatic aldehyde (2.5 mmol of benzaldehyde, salicylaldehyde, or 4-fluorobenzaldehyde) was added, and the mixture was heated in a glycerin bath at 70-75 °C; at this point, ammonium acetate (16.5 mmol) was added. The reaction was conducted in a closed system with a gas conditioning setup attached to the reaction flask. After the total consumption of lapachol, the reaction mixture was treated with 5% (m/v) sodium bisulfite solution at approximately 0 °C.¹⁶-¹⁸

4-(3-Methylbut-2-enyl)-2-phenylnaphtho[1,2-d]oxazol-5-ol (LOX1)

The complete consumption of lapachol occurred 2 h after the reaction started. After treatment with a sodium bisulfite solution, the reaction mixture produced a precipitate, which was purified by column chromatography. A white solid was obtained with a yield of 19.6% and a mp 165-167 °C (lit.¹⁷ mp 168 °C); ¹H NMR (400 MHz, DMSO-d₆) d 9.67 (s, 1H), 8.36 (m, 2H), 8.21 (m, 2H), 7.62 (m, 5H), 5.42 (m, 1H), 3.80 (d, 2H, J 7.3 Hz), 1.92 (s, 3H), 1.68 (s, 3H); ¹³C NMR (100 MHz, DMSO-d₆) d 159.94 (C), 148.22 (C), 131.81 (C), 130.89 (CH), 129.66 (C), 129.31 (CH), 127.08 (C), 126.71 (CH), 126.32 (CH), 124.60 (CH), 124.36 (C), 124.19 (C), 123.39 (CH), 121.56 (CH), 121.39 (CH), 109.28 (C), 25.49 (CH₃), 23.35 (CH₂), 17.83 (CH₃); IR (KBr) ν / cm^-1 3194 (O-H), 1639 (C=N), 1172 (C-O); HRMS (ESI) m/z, calcd. for C₂₂H₁₉NO₂: 329.1416, found: 330.1518 [M + H]⁺.

2-(2-Hydroxyphenyl)-4-(3-methylbut-2-enyl)naphtho[1,2-d]oxazol-5-ol (LOX2)

After 40 min from the start of the reaction, the product precipitated. The solid was filtered and washed with ethanol and sodium bisulfite solution. A white solid was obtained with a yield of 43.3% and mp 191-194 °C; ¹H NMR (400 MHz, DMSO-d₆) d 11.25 (s, 1H), 9.78 (s, 1H), 8.37 (t, 2H, J 8.6 Hz), 8.01 (d, 1H, J 7.5 Hz), 7.67 (t, 1H, J 7.4 Hz), 7.58 (t, 1H, J 7.5 Hz), 7.49 (t, 1H, J 7.6 Hz), 7.13 (m, 2H), 5.41 (t, 1H, J 6.5 Hz), 3.80 (d, 2H, J 7.1 Hz), 1.92 (s, 3H), 1.68 (s, 3H); ¹³C NMR (100 MHz, DMSO-d₆) d 159.73 (C), 156.74 (C), 148.64 (C), 146.87 (C), 132.82 (CH), 131.95 (C), 127.56 (C), 126.94 (CH), 126.44 (CH), 124.90 (CH), 124.22 (C), 123.46 (C), 123.40 (CH), 121.52 (CH), 121.42 (CH), 119.97 (CH), 117.08 (CH), 110.97 (C), 109.15 (C), 25.50 (CH₃), 23.29 (CH₂), 17.83 (CH₃); IR (KBr) ν / cm^-1 3429 (O-H), 1627 (C=N), 1195 (C-O); HRMS (ESI) m/z, calcd. for C₂₂H₁₉NO₃: 345.1365, found: 346.1472 [M + H]⁺.

2-(4-Fluorophenyl)-4-(3-methylbut-2-enyl)naphtho[1,2-d]oxazol-5-ol (LOX3)

The complete total consumption of lapachol occurred 1 h and 20 min after the onset of the reaction. After treatment with sodium bisulfite solution, the reaction mixture yielded a precipitate that was purified by column chromatography. A white solid was obtained with a yield of 20.4% and mp 158-159 °C; ¹H NMR (400 MHz, DMSO-d₆) d 9.66 (s, 1H), 8.34 (d, 2H, J 9.1 Hz), 8.23 (m, 2H), 7.65 (t, 1H, J 7.8 Hz), 7.56 (m, 1H), 7.47 (m, 2H), 5.41 (m, 1H), 3.78 (d, 2H, J 7.3 Hz), 1.91 (s, 3H), 1.68 (s, 3H); ¹³C NMR (100 MHz, DMSO-d₆) d 163.58 (d, ¹J (¹⁹F, ¹³C) 249.2 Hz, C4’), 159.15 (C), 148.24 (C), 148.21 (C), 131.81 (C), 129.62 (C), 128.79 (d, ³J (¹⁹F, ¹³C) 8.9 Hz, C2’ and C6’), 126.70 (CH), 124.61 (CH), 124.33 (C), 124.18 (C), 123.75 (d, ⁴J (¹⁹F, ¹³C) 2.8 Hz, C1’), 123.38 (CH), 121.55 (CH), 121.37 (CH), 116.49 (d, ²J (¹⁹F, ¹³C) 22.3 Hz, C3’ and C5’), 109.27 (C), 25.48 (CH₃), 23.33 (CH₂), 17.82 (CH₃); IR (KBr) ν / cm^-1 3352 (O-H), 1608 (C=N), 1230 (C-F), 1153 (C-O); HRMS (ESI) m/z, calcd. for C₂₂H₁₈FNO₂: 347.1322, found: 348.1426 [M + H]⁺.

Fluorescence quantum yields (Φf) determination

The fluorescence quantum yield (Φ_f) (λ_exc = 345 nm) of the LOX1, LOX2, and LOX3 samples was determined using the methodology described by Zhang et al.,¹⁹ which is based on the classical relative method. All samples were prepared in DMSO, while the 4’,6-diamidino-2-phenylindole (DAPI) standard (Φ_f = 0.021), was prepared in water, as previously described by Prajapati et al.²⁰ The fluorescence quantum yields were then calculated using equation 1:

(1) $${\mathrm{\Phi }}_{\mathrm{f}}={\mathrm{\Phi }}_{\mathrm{f}}^{\mathrm{r}}\times \frac{{\mathrm{F}}_{\mathrm{s}}}{{\mathrm{F}}_0}\times \frac{{\mathrm{A}}_0}{{\mathrm{A}}_{\mathrm{s}}}\times \frac{{\mathrm{n}}_{\mathrm{s}}^2}{{\mathrm{n}}_0^2}$$

where, Φ_f denotes the fluorescence quantum yield, F represents the integrated fluorescence intensity, A signifies the absorbance at the excitation wavelength and n indicates the refractive index of the solvent used. The subscript 0 refers to the reference, and the subscript s refers to the sample.

DNA binding study by UV-Vis absorption analysis

Absorption spectra of LOX1, LOX2, and LOX3 both in the absence of and in the presence of increasing concentrations of CT-DNA were obtained at room temperature in DMSO (5%)/Tris-HCl buffer (0.1 M, pH 7.4) mixture solution in the 200-800 nm range, and DMSO (5%)/Tris-HCl buffer (0.1 M, pH 7.4) was used as a reference medium. The concentrations naphthoxazoles were fixed at 10 μM, while CT-DNA concentrations varied from 0 to 320 μM. The naphthoxazole-DNA solutions were allowed to equilibrate at room temperature for 5 min before absorption spectra were recorded. The spectra were acquired using a rectangular quartz cuvette with a 1 cm optical path length. The percentage of hypochromism or hyperchromism (H) was calculated using the equation H (%) = (A_free - A_bound)/A_free × 100, where A_free is the absorbance before interaction with DNA, and A_bound is the absorbance after interaction with DNA. Bathochromic or hypsochromic shift was quantified by the equation ∆λ = λ_final - λ_initial, λ_initial is the wavelength of maximum absorption before interaction with DNA, and λ_final is the wavelength of maximum absorption after interaction with DNA.²,²¹ Data analysis and calculations were performed using Microsoft Excel (Microsoft Corporation, Redmond, WA, USA).²²

Results and Discussion

Synthesis and structural characterization

The naphthoxazoles (LOX1, LOX2, and LOX3) were synthesized using the Debus-Radziszewski reaction in a one-pot process involving lapachol and the corresponding aldehyde, with ammonium acetate serving as the source of ammonia (Scheme 1).¹⁶-¹⁸

The Debus-Radziszewski reaction typically employs an α-dicarbonyl compound, an aldehyde, and a nitrogen source. This reaction is one of the most commonly used methodologies for obtaining 2,4,5-substituted imidazoles. However, subsequent studies have reported that, in some cases, the Debus-Radziszewski methodology allows for the formation of two isosteric azoles: imidazole and oxazole.⁵,²³

In this study, the previously mentioned reaction predominantly yielded oxazole derivatives, leading to cleaner reactions. According to Del Rio et al.,¹⁸ since lapachol possesses only one free carbonyl group, the formation of an imidazole derivative, a keto-enol tautomerization of its phenolic hydroxyl would be necessary to generate an o-quinonimine intermediate. However, this process results in the loss of ring aromaticity, which disfavors the formation of the keto tautomer and prevents the nucleophilic attack of the amine. Consequently, the synthetic route to the imidazole is hindered (Figure 1).

Figure 1
Proposed mechanism for the Debus-Radziszewski reaction from lapachol.

The synthetic methodologies used for producing LOX1, LOX2, and LOX3 are adaptations, incorporating principles of green chemistry, from previously described methodologies.¹⁶-¹⁸ The use of ethanol as the solvent in these reactions not only provides environmental and economic benefits but also results in shorter reaction times (40 min to 2 h) and satisfactory yields (19.6 to 43.3%).

Among the three compounds synthesized in this study, only LOX1 has been previously described in the literature.¹⁷,¹⁸ LOX2 and LOX3 represent novel structures. LOX1, LOX2, and LOX3 were elucidated through combined analysis of one-dimensional (1D) and two dimensional (2D) NMR spectra and were confirmed by IR and HRMS techniques.

It is known that the synthesis of naphthoxazoles from lapachol typically yields two derivatives: a minor product, which is naphthoxazole derivative of β-xyloidone, and a major product, which is the naphthoxazole with the isoprenyl side chain of lapachol.¹⁷,¹⁸ According to Del Rio et al.¹⁸ the formation of the naphthoxazole derivative of β-xyloidone occurs with longer reaction times.

Given the characteristics described in the literature for the naphthoxazolic derivatives of β-xyloidone, we recognize that these compounds may form as secondary products of the reactions. However, due to their low yield and purity, structural analysis was not conducted.

The signals observed in the NMR spectra of LOX1 are consistent with the literature data for this compound (Table S1, Supplementary Information (SI) section). Additionally, IR and HRMS spectroscopy analyses confirmed the molecular structures of the three naphthoxazoles (LOX1, LOX2, and LOX3) (Figures S13-S16, SI section).

The ¹H NMR spectrum of LOX2 exhibited a singlet at d_H 11.25 ppm (1H). This signal, with a high chemical shift value, is characteristic of the formation of an intramolecular hydrogen bond between the phenolic substituent at position 2 of the oxazole nucleus and its nitrogen,¹¹Figure 2 (Figure S1, SI section).

Figure 2
Correlations confirming the presence of (a) LOX2 and (b) LOX3. Double arrows represent correlations between vicinal hydrogens observed in the ¹H-¹H COSY spectrum, while single arrows represent ²,³J_H-C correlations through the ¹H-¹³C HMBC spectrum.

The number of hydrogens obtained from signal integration and their spin-spin coupling corresponds to the structure of LOX2. The spectra of the other naphthoxazoles also exhibited integration and multiplicity consistent with their molecular structures.

The ¹³C NMR spectrum of LOX2 (Figure S2, SI section) exhibited 22 peaks corresponding to the carbons present in its molecular structure. The signals at d_C 156.74 and 148.64 ppm characterized two oxygenated carbons with sp² hybridization, corresponding to the carbons of the two hydroxyls of LOX2. The signal at d_C 148.64 ppm was confirmed as C10 by the heteronuclear 2D ¹H ¹³C heteronuclear multiple bond correlation (HMBC) spectrum (Figure S4, SI section). In this spectrum, the correlation between the signals at d_H 8.37 and 3.80 ppm with the carbon C10 (d_C 148.64 ppm) was observed. According to the heteronuclear 2D ¹H-¹³C heteronuclear single quantum coherence spectroscopy (HSQC) spectrum (Figure S5, SI section), the signals at d_H 8.37 and 3.80 ppm correspond to hydrogens H9 and H12, respectively (Figure 2).

The signal at d_H 8.37 ppm (2H) in the homonuclear 2D ¹H-¹H correlation (COSY) spectrum (Figure S6, SI section), which exhibits coupling between hydrogen atoms, correlates with the signal at d_H 7.67 ppm, corresponding to either hydrogen H8 or H7. This suggests that the signal at d_H 8.37 ppm corresponds to hydrogen atoms H9 and H6 (Figure 2).

Hydrogen H6 (d_H 8.37 ppm), in turn, correlates with the signal at d_C 127.56 ppm (C5). Furthermore, the signal at d_H 3.80 ppm exhibits a ³J_H-C correlation with the signal at d_C 146.87 ppm (C4), confirming the presence of the oxazole nucleus in LOX2 (Figure 2). The chemical shifts of C4 and C5 are similar to those observed by Dias et al.²⁴ for their 2-substituted naphthoxazoles with a phenol group. Figure 2 presents the main chemical shifts and correlations that confirmed the presence of the oxazole nucleus, as discussed earlier.

Another significant aspect to consider is the chemical shift of carbon C2. As reported by Dias et al.,²⁴ carbon C2 demonstrates chemical shift values ranging from 159.0 to 160.2 ppm. LOX2 exhibited a chemical shift for C2 of 159.73 ppm. The signal at d_C 159.73 ppm displays a ³J_H-C correlation with the signal at d_H 8.01 ppm. Simultaneously, the signal at d_H 8.01 ppm also demonstrates homonuclear correlation with the signal at d_H 7.13 ppm (m, 2H) and heteronuclear ³J_H-C correlation with the signal at d_C 156.74 ppm. Lastly, to confirm the presence of LOX2, the signal at d_H 7.13 ppm also correlates ³J_H-C with the signal at d_C 110.97 ppm, which is characteristic of carbon C1’ (see Figure 2). The chemical shift value obtained for C1’ also corresponds with the values reported by Dias et al.²⁴ (111.4-111.5 ppm for uncoordinated naphthoxazoles).

The structure of LOX3 was also elucidated through combined analysis of 1D and 2D NMR spectra (Figures S7 S12, SI section). Initially, two characteristic signals of oxygenated carbons with sp² hybridization were observed at d_C 148.24 and 148.21 ppm, corresponding to carbons C10 and C4 of LOX3. Although the values might be interchanged due to the similarity of their chemical environment, the chemical shifts of carbons C10 and C4 were confirmed by the ³J_H-C correlation between signals at d_H 8.34 and 3.78 ppm with both signals at d_C 148.24 and 148.21 ppm (Figure 2). The signals at d_H 8.34 and 3.78 ppm correspond to hydrogens H9 and H12, respectively. The signal at d_H 8.34 ppm (2H), as observed for LOX2, corresponds to hydrogens H9 and H6, as confirmed through homonuclear correlations between the signal at d_H 8.34 ppm and signals at d_H 7.65 and 7.56 ppm corresponding to hydrogens H7 and H8. Hydrogen H6 (d_H 8.34 ppm), in turn, correlates with the signal at d_C 129.62 ppm (C5), confirming the presence of the oxazole nucleus in LOX3 (Figure 2). The chemical shifts of C4 and C5 are similar to those reported by Aljaar et al.²⁵ for their 2-substituted naphthoxazole with a fluorobenzene group (d_C 148.0 and 137.6 ppm).

Carbon C2 of LOX3 exhibited a chemical shift value of 159.15 ppm, which is similar to that observed for LOX2 and characteristic of this carbon in naphthoxazole derivatives, as discussed previously.

Additionally, the signal at d_C 159.15 ppm exhibits a ³J_H-C correlation with the signal at d_H 8.23 ppm. Simultaneously, the signal at d_H 8.23 ppm exhibits a homonuclear correlation with the signal at d_H 7.47 ppm (m, 2H) and heteronuclear ³J_H-C correlation with the signal at d_C 163.58 ppm (C4’). Finally, the signal at d_H 7.47 ppm also shows a ³J_H-C correlation with the signal at d_C 123.75 ppm, characteristic of carbon C1’, and ²J_H-C with the signal at d_C 163.58 ppm (Figure 2). The chemical shifts at d_C 123.75 and 163.58 ppm for C1’ and C4’, respectively, are consistent with those reported by Aljaar et al.²⁵ (d_C 123.7 and 164.5 ppm).

Furthermore, the ¹³C NMR spectrum of LOX3 (Figure S8, SI section), also exhibited ¹³C-¹⁹F couplings, which provided crucial information for the assignment of the previously discussed chemical shifts. According to Pavia et al.,²⁶ when an organic compound has a fluorine atom attached to a carbon-13 atom, a heteronuclear ¹³C ¹⁹F coupling is observed. As stated by Branco et al.,²⁷ it is more common to obtain ¹³C NMR spectra with ¹³C ¹⁹F couplings for organofluorides than to obtain decoupled spectra.

In the ¹³C NMR spectrum of LOX3 (Figure S8, SI section), doublets were observed, reflecting the presence of a fluorine atom in the structure, resulting from ¹³C ¹⁹F couplings via ¹J_C-F up to ⁴J_C-F. The doublet at d_C 163.58 ppm (the value corresponding to the true chemical shift of the carbon atom, at the center of the doublet) corresponds to direct coupling via a bond between the fluorine and C4’ (ipso carbon) with a high coupling constant, J 249.2 Hz. This high coupling constant value is typical in direct fluorine couplings with a carbon-13 atom (¹J_C-F).²⁵-²⁷

Subsequently, the doublet at d_C 116.49 ppm, with J 22.3 Hz, corresponds to the coupling between the fluorine and the ortho carbons, C3’ and C5’. As expected, coupling constants decrease with increasing distance between the carbon and the fluorine.²⁵-²⁷ Meanwhile, the doublet at d_C 128.79 ppm, with J 8.9 Hz, corresponds to the coupling between the fluorine and the meta carbons, C2’ and C6’.

Furthermore, the doublet at d_C 123.75 ppm, with J 2.8 Hz, which corresponds to the coupling between fluorine and carbon C1’ via ⁴J_C-F, as also reported by Aljaar et al.²⁵ for their 2-substituted naphthoxazole with a fluorobenzene group. It is worth noting that the chemical shift values and coupling constants were all similar to those observed by Aljaar et al.,²⁵ confirming the synthesis of LOX3.

Photophysical properties

The naphthoxazoles are fluorophores that exhibit increased emission intensity upon binding to DNA.²,³ Therefore, the synthesis of naphthoxazoles with diverse structural characteristics, which could potentially influence their optical properties, should be explored to optimize the efficiency of fluorescent sensors.

LOX2 is a fluorophore capable of excited-state intramolecular proton transfer (ESIPT), as it features a proton donor group (hydroxyl group -OH) and a proton acceptor group (imine nitrogen -N=) in close proximity, forming an intramolecular hydrogen bond in the ground electronic state (Figure 3).²⁸,²⁹ On the other hand, LOX3 is a D-π-A conjugated system, containing an electron donor group (D) separated from an electron acceptor group (A) at the opposite ends other of the molecule by a π-linker bridge.³⁰,³¹ Hence, LOX3 is representative of a class of fluorophores commonly known as push-pull systems (Figure 3).³⁰,³²,³³

Figure 3
Molecular structure of the naphthoxazoles LOX1, LOX2, and LOX3, highlighting the structural features of LOX2 and LOX3.

In general, fluorophores with these characteristics exhibit large Stokes shifts and sensitivity to polarity, with results in changes to their emission profile in response to environmental conditions, among other properties. These properties are desirable for fluorescent molecules used in bioimaging.²⁸,³¹,³³

Large Stokes shifts eliminate spectral overlap between fluorophore absorption and emission, thereby contributing to a clearer fluorescence signal and reduced background interference.³⁴-³⁶ The optical characteristics of the naphthoxazoles LOX1, LOX2, and LOX3 were studied and are summarized in Table 1. LOX1, LOX2, and LOX3 exhibited large Stokes shifts, comparable to DAPI (Table 1). DAPI is a DNA fluorescent probe that binds to the minor groove.⁶,³⁷

Table 1
Optical properties of LOX1, LOX2, and LOX3 in DMSO, using DAPI in water as the standard, as described by Prajapati et al.²⁰

Compound	λabsmax / nm	λemmax / nm	ΔS / nm (cm-1)	Φf	τf / ns
DAPI	345	693	348 (14576)	0.0210	2.78a
LOX1	348	690	342 (14242)	0.0018	14.44
LOX2	353	690	337 (13835)	0.0020	1.45
LOX3	347	690	343 (14325)	0.0014	8.08

• ^a
  Berezin and Achilefu.³⁸ DAPI: 4’,6-diamidino-2-phenylindole;
• λ_abs^max: maximum emission wavelength; ∆S: Stokes shift; Φ_f: fluorescence quantum yield; τ_f: fluorescence lifetime.

Each compound fluorescence quantum yields (Φ_f) were calculated using the classical relative method, with DAPI as the reference standard (Table 1). The Φ_f is defined as the ratio between the number of photons emitted and the number of photons absorbed by the fluorophore.¹⁹,³⁷,³⁹ The relative method involves evaluating the absorption profile of the molecule followed by fluorescence emission, and then mathematically determining the quantum yield from the data obtained.¹⁹,³⁹

Φ_f is a parameter highly sensitive to the concentration of the sample and the excitation wavelength due to the self-absorption effect, wherein the molecule absorbs its own emitted energy.³⁹ To prevent this effect, the absorbance of the samples should be kept below 0.1 as described by Eaton.⁴⁰ Additionally, the chemical environment in which the molecule is solubilized directly impacts the fluorescence emission profile and consequently the fluorescence quantum yield.³⁷ Substances with a high fluorescence quantum yield, close to 1, exhibit an intense fluorescence emission profile, as nearly all absorbed energy is dissipated as photon emission.³⁷

DAPI has a low Φ_f; however, it is well known that this radiative effect increases significantly when bound with DNA molecules.²⁰ Similarly, we can consider that the LOXs behave analogously to DAPI, and exhibit reduced fluorescence in their free form, as demonstrated by the fluorescence quantum yield study (Figure 4).

Figure 4
UV-Vis 2D fluorescence emission spectra (a) and normalized spectra (b) of LOX1, LOX2, and LOX3 in DMSO, with DAPI in water as a standard compound. λ_exc = 345 nm, with excitation and emission bandwidths of 10 nm each.

Fluorescence lifetime (τ_f) refers to the duration for which an electron remains excited before returning to its ground state after absorbing light. During this return to the ground state, the absorbed energy is dissipated through fluorescence emission and other non-radiative processes, collectively known as quenching.³⁷,³⁸

The τ_f does not depend on conditions such as the excitation wavelength and the duration of exposure to light. Additionally, it does not depend on fluorescence intensity or fluorophore concentration. As the process of fluorescence emission time is associated with an energetically unstable state, the duration of fluorescence can be sensitive to a wide variety of internal factors such as the structure of the fluorophore, and external factors such as temperature, polarity, and the presence of fluorescence quenchers.³⁸,⁴¹

In this context, τ_f can be influenced by reversible changes in electron distribution occurring in the excited state. These changes, which are related to the structure of the fluorophore, are generally less susceptible to environmental factors. Significant processes in this category include excited-state charge transfers, such as electron transfer (ESET) and proton transfer (ESPT), as well as intersystem crossings.³⁸

The τ_f of the LOXs in their free state was typically in the nanosecond range, as expected for fluorescent organic dyes.¹¹,³⁷ Naphthoxazole LOX2 exhibited the shorter τ_f, at 1.45 ns. This shorter value, in comparison to the naphthoxazoles LOX1 and LOX3 (Table 1), suggests the influence of ESIPT on the loss of energy from the excited state, as this process may compete with the radiative process.³⁸,⁴¹ The long lifetimes of LOX1 and LOX3 are useful, as long τ_f are used in biological systems to completely eliminate autofluorescence background.³⁸

The increase in τ_f of fluorophores upon binding to DNA is well-documented. For example, the τ_f of ethidium bromide (EB) is approximately 1.7 ns in the free form and increases to about 20 ns after binding to DNA.³⁷,³⁸ This effect results from the restriction of fluorophore mobility upon interaction with DNA, leading to a decrease in non-radiative processes and consequently an increase in τ_f and fluorescence intensity.²,³⁸

DNA binding study

UV-Vis absorption spectroscopy is the most commonly employed technique for studying DNA interactions with small molecules.¹⁰ This investigation can be conducted by monitoring changes in the absorption properties of either the ligand molecule or the DNA.¹⁰,⁴²,⁴³

Generally, molecules used as DNA ligands exhibit absorption bands in the visible region, which are easily distinguishable from the absorption bands of DNA. DNA does not absorb in the visible region; its maximum absorption band is in the ultraviolet region at 260 nm.¹⁰,¹⁴ Therefore, examining changes in the intensity and position of the absorption bands of the ligand after interaction with DNA, compared to when the ligand molecule is free in solution, can indicate binding between DNA and the molecule.¹⁰,¹¹,¹⁴,¹⁵,⁴²,⁴⁴ The extent of this change can be interpreted as an indication of the strength of the interaction between DNA and the ligand.¹⁰,⁴⁵,⁴⁶

Small aromatic molecules can bind to DNA through three main modes of interaction: intercalation between base pairs, binding to minor and major DNA grooves, or electrostatic interactions between phosphate groups and charged species.¹⁰,¹¹

Compounds that bind to DNA through intercalation typically result in hypochromism and a bathochromic shift (red shift).²,¹⁰,⁴⁵ In contrast, compounds that interact by binding to the grooves or to phosphate groups often exhibit a hyperchromic effect which may or may not be accompanied by slight bathochromic or hypsochromic shifts (blue shift).¹⁰,⁴⁷,⁴⁸ It is important to note that in weak interactions, only hypochromic or hyperchromic effects are observed, that is, without significant changes in the spectral profile.¹⁰,⁴²

The naphthoxazoles (LOX1, LOX2, and LOX3) exhibited similar absorption profiles in DMSO (5%)/Tris-HCl buffer (pH 7.4) (Figure 5). Each compound displays three main absorption bands of high intensity, the most intense band is in the ultraviolet region, while the remaining bands are in the visible region. The wavelengths of maximum absorption (λ_abs) for LOX1, LOX2, and LOX3 are in Figure 5.

Figure 5
UV-Vis absorption spectra of LOX1, LOX2, and LOX3 in DMSO (5%)/Tris-HCl buffer (pH 7.4).

Aggregation assays determine the maximum concentration a molecule can reach before it aggregates in solution. The formation of aggregates causes bathochromic or hypsochromic shifts in the absorption spectrum, depending on the type of aggregate.⁴⁹,⁵⁰ The aggregation behavior of LOX2, selected as the representative compound, was evaluated using UV-Vis spectroscopy in DMSO(5%)/Tris-HCl buffer (pH 7.4). Aggregation was ruled out as no significant shifts in the maximum absorbance wavelengths were observed. A linear increase in the absorbance was noted as the concentration variation from 5.0 to 22.5 µM (Figure 6).

Figure 6
Aggregation study of LOX2 in DMSO (5%)/Tris-HCl buffer (pH 7.4) (inset: plot of the linear relationship of absorbance at 359 nm as a function of the concentration 5.0-22.5 µM).

The naphthoxazoles (LOX1, LOX2, and LOX3) were evaluated for their potential interaction with DNA. DNA binding studies involved monitoring changes in the absorption properties of LOX1, LOX2, and LOX3 both in the absence and presence of CT-DNA at various concentrations (Figure 7).

Figure 7
Absorption spectra of LOX1, LOX2, and LOX3 upon additions of CT-DNA, in DMSO (5%)/Tris-HCl buffer (pH 7.4). (a), (c), and (e) display the full spectrum, and (b), (d), and (f) highlight the main absorption bands in the visible region of LOX1, LOX2, and LOX3, respectively.

Addition of aliquots of CT-DNA (0-320 µM) to LOX1 resulted in significant changes to its absorption profile. Initially, a decrease in the absorbance of the main band at 360 nm was observed after the addition of the first aliquots of CT-DNA (10-40 µM), indicative of a hypochromic effect (2.95-8.87%). As the of CT-DNA concentration increased from 10 to 40 µM, the extent of hypochromism decreased from 8.87 to 2.95% (Figure 7).

However, at concentrations above 80 µM of CT DNA, a gradual increase in the absorbance of the main band was observed, indicative of a hyperchromic effect (1.18-10.65%). Additionally, increasing the CT-DNA concentration from 80 to 320 µM resulted in the appearance of a single broad absorption band with a blue shift of 1 6 nm (Figure 7).

Felouat et al.¹¹ observed a variation in the intensity of the bands during the DNA interaction assay for an ESIPT emitter benzoxazole. Initially, the authors noted a gradual increase in the absorbance of the compounds with the addition of the first aliquots of DNA. However, as the increase in DNA concentration, they observed pronounced hypochromism of the main absorption bands of benzoxazole. According to Felouat et al.¹¹ this significant alteration at higher DNA concentrations may be indicative of a second binding mode occurring after the initial mode of interaction.

The absorption profile of LOX1 at different concentrations of CT-DNA suggests a potential interaction between LOX1 and DNA via groove binding, given that this compound does not possess charged groups capable of interacting with the phosphate groups of DNA.¹⁰,⁴⁷,⁴⁸ This type of interaction is consistent with the molecular structure of LOX1, as aromatic rings connected by bonds that allow free rotation facilitate DNA groove binding.¹⁰

Similar to what was observed for LOX1, addition of aliquots CT-DNA (0-320 µM) to LOX2 resulted in changes in its absorption profile. A gradual increase in the absorbance of the main band (359 nm) was observed after the addition of CT-DNA aliquots (10-320 µM), characteristic of a hyperchromic effect (1.57-11.51%). This hyperchromism was accompanied by a blue shift (1 2 nm) (Figure 7).

It was also noticeable that, at the highest concentration of CT-DNA (320 µM) in the assay, LOX2 exhibited a smaller blue shift (1 nm) compared to other concentrations. Additionally, at 320 µM of CT-DNA, LOX2 demonstrated a lower hyperchromism rate compared to the previous CT DNA concentration (160 µM), indicating the interaction limit threshold between LOX2 and DNA (Figure 7).

When assessing the effect of increasing DNA concentrations on the absorption spectrum of EB, Waring⁵¹ observed a progressive bathochromic shift of the maximum absorption band (479 nm) towards a limit that, according to the author, represents the spectrum of EB in a fully complexed form. At this limit, EB exhibits the greatest bathochromic shift, but the hypochromism rate is not the highest.⁵¹ Beyond this limit, there appears to be a trend towards an increase in the band intensity compared with the intensity at lower DNA concentrations.

Thus, the hyperchromism and slight blue shift observed suggest a potential interaction between LOX2 and DNA via grooves binding, as its molecular structure, similar to LOX1, also lacks charged groups capable of interacting with the phosphate groups of DNA.¹⁰,⁴⁷,⁴⁸,⁵²

Except for the initial and final absorbance values of the assay, the DNA binding constant (K or K_b) of LOX2 was determined from the plot of [DNA]/(ε_a - ε_f) versus [DNA], where [DNA] is the DNA concentration (M^-1), ε_a is the apparent molar absorptivity (M^-1 cm^-1; corresponding to the ratio of observed absorbance to species concentration), and ε_f is the free molar absorptivity (M^-1 cm^-1) of the chemical species, i.e., in the absence of DNA. The value of K_b was calculated from the ratio of the slope and intercept coefficients (Figure 8).³,¹⁰,²¹

Figure 8
Plot of [DNA]/(ε_a - ε_f) vs. [DNA] in the main absorption band of the visible region. [LOX2] = 10 µM and [CT-DNA] = 20 to 160 µM.

The K_b value of LOX2 was 2.70 × 10⁴ M^-1. This K_b value indicates that LOX2 can strongly bind to the biomolecule.⁵⁰ The DNA binding constant of LOX2 was higher than the values obtained by Wang et al.²,³ for most of their evaluated naphthoxazoles (3.58 × 10³-5.29 × 10⁴ M^-1).

In contrast to LOX1 and LOX2, adding of aliquots of CT-DNA (0-320 µM) to LOX3 has resulted in minor changes in its absorption profile. Initially, a decrease in the absorbance of the main band (346 nm) was observed after the addition of the first aliquots of CT-DNA (10 40 µM), which is characteristic of a hypochromic effect (0.87 3.50%). As the concentration of CT-DNA increased (10 → 40 µM), the rate of hypochromism also increased from 0.87 to 3.50% (Figure 7).

However, similar to what was observed for LOX1, a gradual increase in the absorbance of the main band was noted above 80 µM of CT-DNA, indicating a hyperchromic effect (0.87-8.77%). Additionally, as the concentration of CT-DNA increased from 80 to 320 µM, a single broad absorption band appeared. No significant deviations in the λ_abs for LOX3 were observed (Figure 7).

These observations also suggest a potential interaction between LOX3 and DNA via groove binding, as LOX3 lacks charged groups that would facilitate binding to the phosphate groups of DNA.¹⁰,⁴⁷,⁴⁸ The absence of bathochromic or hypsochromic shifts suggests a weak interaction with the biomolecule.¹⁰,⁴²

Conclusions

The naphthoxazoles derived from lapachol (LOX1, LOX2, and LOX3) were synthesized using an economically and environmentally viable synthesis protocol. The range of yields of LOX1, LOX2, and LOX3 varied from 19.6 to 43.3%. These compounds exhibited large Stokes shifts, reduced fluorescence, low values of fluorescence quantum yield in their free form, demonstrating behavior similar to that of DAPI. Furthermore, they exhibited typical lifetimes in the nanosecond range. LOX1 and LOX3 showed long lifetimes of 14.44 and 8.08 ns, respectively. UV-Vis absorption spectroscopy of DNA binding studies revealed that the naphthoxazoles interact with DNA, as indicated by changes in the absorption properties with increasing concentrations of CT-DNA. The results suggest that LOX1, LOX2, and LOX3 bind to DNA via groove binding. Notably, LOX2 showed a pronounced K_b value, indicating strong binding to DNA. These findings suggest that LOX1, LOX2, and LOX3 may be suitable candidates as DNA fluorescent probes.

Supplementary Information

Supplementary data are available free of charge at http://jbcs.sbq.org.br as PDF file.https://minio.scielo.br/documentstore/1678-4790/yV4KqQVJp9PqC7FRvDDzmKN/0a08bba74add4077aff5854097f1b856e53ae027.pdf

Acknowledgments

The authors would like to thank the FACEPE for financial support IBPG-1482-4.03/19 and APQ-0788-1.06/22, CAPES and the National Laboratory of Bio-Renewables (LNBR) at the National Center for Energy and Materials Research (CNPEM) for the HRMS mass analysis.

References

• ¹ Slovesnova, N. V.; Minin, A. S.; Smolyuk, L. T.; Taniya, O. S.; Tsmokalyuk, A. N.; Kim, G. A.; Kovalev, I. S.; Pozdina, V. A.; Kopchuk, D. S.; Krinochkin, A. P.; Zyryanov, G. V.; Petrov, A. Y.; Charushin, V. N.; Dyes Pigm. 2022, 204, 110410. [Crossref]
  » Crossref
• ² Wang, X.-Z.; Yao, J.-H.; Xie, Y.-Y.; Lin, G.-J.; Huang, H.-L.; Liu, Y.-J.; Inorg. Chem. Commun. 2013, 32, 82. [Crossref]
  » Crossref
• ³ Wang, X.-Z.; Jiang, G.-B.; Xie, Y.-Y.; Liu, Y.-J.; Spectrochim. Acta, Part A 2014, 118, 448. [Crossref]
  » Crossref
• ⁴ Zhang, J.; Chen, R.; Zhu, Z.; Adachi, C.; Zhang, X.; Lee, C.-S.; ACS Appl. Mater. Interfaces 2015, 7, 26266. [Crossref]
  » Crossref
• ⁵ Santos, V. L. A.; Guimarães, D. G.; Nishimura, R. H. V.; Rolim, L. A.; Gonsalves, A. A.; Araújo, C. R. M.; Quim. Nova 2022, 45, 560. [Crossref]
  » Crossref
• ⁶ Bertozo, L. C.; Tutone, M.; Pastrello, B.; da Silva-Filho, L. C.; Culletta, G.; Almerico, A. M.; Ximenes, V. F.; J. Photochem. Photobiol., A 2023, 444, 114944. [Crossref]
  » Crossref
• ⁷ Osadchii, S. A.; Shubin, V. G.; Kozlova, L. P.; Varlamenko, V. S.; Filipenko, M. L.; Boyarskikh, U. A.; Russ. J. Appl. Chem. 2011, 84, 1541. [Crossref]
  » Crossref
• ⁸ El-din, A. E-D. S.; Abdullah, S.; Sayed, A. E.-D. H.; Sci. Afr. 2021, 13, e00961. [Crossref]
  » Crossref
• ⁹ Soriano, E.; Holder, C.; Levitz, A.; Henary, M.; Molecules 2016, 21, 23. [Crossref]
  » Crossref
• ¹⁰ Sirajuddin, M.; Ali, S.; Badshah, A.; J. Photochem. Photobiol., B 2013, 124, 1. [Crossref]
  » Crossref
• ¹¹ Felouat, A.; Massue, J.; Ulrich, G.; Dyes Pigm. 2021, 185, 108895. [Crossref]
  » Crossref
• ¹² Liu, Q.-Q.; Lu, K.; Zhu, H.-M.; Kong, S.-L.; Yuan, J.-M.; Zhang, G.-H.; Chen, N.-Y.; Gu, C.-X.; Pan, C.-X.; Mo, D.-L.; Su, G.-F.; Eur. J. Med. Chem. 2019, 165, 293. [Crossref]
  » Crossref
• ¹³ Deng, C.; Yan, H.; Wang, J.; Liu, B-s.; Liu, K.; Shi, Y-m.; Arabian J. Chem. 2022, 15, 104242. [Crossref]
  » Crossref
• ¹⁴ Santos-Junior, P. F. S.; Nascimento, I. J. S.; da Silva, E. C. D.; Monteiro, K. L. C.; de Freitas, J. D.; de Lima Lins, S.; Maciel, T. M. S.; Cavalcante, B. C.; Neto, J. B. V.; de Abreu, F. C.; Fegueiredo, I. M.; Santos, J. C. C.; Pessoa, C. Ó.; da Silva Júnior, E. F.; de Araújo-Júnior, J. X.; de Aquino, T. M.; New J. Chem. 2021, 45, 13847. [Crossref]
  » Crossref
• ¹⁵ Silva, M. M.; Savariz, F. C.; da Silva-Júnior, E. F.; de Aquino, T. M.; Sarragiotto, M. H.; Santos, J. C. C.; Figueiredo, I. M.; J. Braz. Chem. Soc. 2016, 27, 1558. [Crossref]
  » Crossref
• ¹⁶ Santos, V. L. A.; Gonsalves, A. A.; Guimarães, D. G.; Simplicio, S. S.; de Oliveira, H. P.; Ramos, L. P. S.; da Costa, M. P.; de Oliveira, F. C. E.; Pessoa, C.; Araújo, C. R. M.; Molecules 2023, 28, 3008. [Crossref]
  » Crossref
• ¹⁷ de Moura, K. C. G.; Emery, F. S.; Neves-Pinto, C.; Pinto, M. C. F. R.; Dantas, A. P.; Salomão, K.; Castro, S. L.; Pinto, A. V.; J. Braz. Chem. Soc. 2001, 12, 325. [Crossref]
  » Crossref
• ¹⁸ Del Rio, K. P.; de Moura, K. C. G.; Pinto, M. C. F. R.; Leitão, G. G.; J. Liq. Chromatogr. Relat. Technol. 2015, 38, 1479. [Crossref]
  » Crossref
• ¹⁹ Zhang, X.-F.; Zhang, J.; Liu, L.; J. Fluoresc. 2014, 24, 819. [Crossref]
  » Crossref
• ²⁰ Prajapati, R.; Chatterjee, S.; Kannaujiya, K. K.; Mukherjee, T. K.; Nanoscale 2016, 8, 13006. [Crossref]
  » Crossref
• ²¹ Acunha, T. V.; Chaves, O. A.; Iglesias, B. A.; J. Porphyrins Phthalocyanines 2021, 25, 75. [Crossref]
  » Crossref
• ²² Microsoft Excel, Office Home and Student; Microsoft Corporation, Redmond, WA, USA, 2021.
• ²³ Santos, V. L. A.; Gonsalves, A. A.; Araújo, C. R. M.; Quim. Nova 2020, 43, 1344. [Crossref]
  » Crossref
• ²⁴ Dias, G. G.; Rodrigues, B. L.; Resende, J. M.; Calado, H. D. R.; de Simone, C. A.; Silva, V. H. C.; Neto, B. A. D.; Goulart, M. O. F.; Ferreira, F. R.; Meira, A. S.; Pessoa, C.; Correa, J. R.; da Silva Júnior, E. N.; Chem. Commun. 2015, 51, 9141. [Crossref]
  » Crossref
• ²⁵ Aljaar, N.; Malakar, C. C.; Conrad, J.; Frey, W.; Beifuss, U.; J. Org. Chem. 2013, 78, 154. [Crossref]
  » Crossref
• ²⁶ Pavia, D. L.; Lampman, G. M.; Kriz, G. S.; Vyvyan, J. R.; Introdução à Espectroscopia, 4^th ed.; Cengage Learning: São Paulo, Brazil, 2010.
• ²⁷ Branco, F. S. C.; Silva, B. V.; do Rio, G. F.; Santana, M. J.; Queiroz Jr., L. H. K.; Pinto, A. C.; Boechat, N.; Lião, L. M.; Quim. Nova 2015, 38, 1237. [Crossref]
  » Crossref
• ²⁸ Klinhom, N.; Saengsuwan, N.; Sriyab, S.; Prompinit, P.; Hannongbua, S.; Suramitr, S.; Spectrochim. Acta, Part A 2019, 206, 359. [Crossref]
  » Crossref
• ²⁹ Purkayastha, P.; Chattopadhyay, N.; J. Mol. Struct. 2002, 604, 87. [Crossref]
  » Crossref
• ³⁰ Khaled, M. M.; Ismail, M. A.; Medien, H. A. A.; Abdel-Shafi, A. A.; Abdel-Samad, H. S.; Spectrochim. Acta, Part A 2023, 288, 122090. [Crossref]
  » Crossref
• ³¹ Cao, C.; Cao, C.; J. Lumin. 2021, 239, 118382. [Crossref]
  » Crossref
• ³² Bureš, F.; RSC Adv. 2014, 4, 58826. [Crossref]
  » Crossref
• ³³ Yin, G.; Li, Y.; Li, S.; Xu, B.; Yang, Q.; Zhang, Y.; Zhao, J.; Cao, X.; Dyes Pigm. 2022, 198, 110013. [Crossref]
  » Crossref
• ³⁴ Le, B. H.; Nguyen, T.-V. T.; Joo, H. N.; Seo, Y. J.; Bioorg. Med. Chem. 2018, 26, 4881. [Crossref]
  » Crossref
• ³⁵ Dai, L.; Zhang, Q.; Ma, Q.; Lin, W.; Coord. Chem. Rev. 2023, 489, 215193. [Crossref]
  » Crossref
• ³⁶ Abeywickrama, C. S.; Wijesinghe, K. J.; Stahelin, R. V.; Pang, Y.; Sens. Actuators, B 2019, 285, 76. [Crossref]
  » Crossref
• ³⁷ Lakowicz, J. R.; Principles of Fluorescence Spectroscopy, 3^rd ed.; Springer: New York, USA, 2006.
• ³⁸ Berezin, M. Y.; Achilefu, S.; Chem. Rev. 2010, 110, 2641. [Crossref]
  » Crossref
• ³⁹ Nawara, K.; Waluk, J.; Anal. Chem. 2017, 89, 8650. [Crossref]
  » Crossref
• ⁴⁰ Eaton, D. F. Pure Appl. Chem. 1988, 60, 1107. [Crossref]
  » Crossref
• ⁴¹ Valeur, B.; Molecular Fluorescence: Principles and Applications, 1^st ed.; Wiley-VCH Verlag: Weinheim, Federal Republic of Germany, 2001.
• ⁴² Lavanya, K.; Babu, P. V.; Bodapati, A. T. S.; Reddy, R. S.; Madku, S. R.; Sahoo, B. K.; Int. J. Biol. Macromol. 2023, 244, 125301. [Crossref]
  » Crossref
• ⁴³ Ressler, A. J.; Frate, M.; Hontoria, A.; Ream, A.; Timms, E.; Li, H.; Stettler, L. D.; Bollinger, A.; Poor, J. E.; Parra, M. A.; Ma, H.; Seeram, N. P.; Meschwitz, S. M.; Henry, G. E.; Bioorg. Med. Chem. 2023, 90, 117369. [Crossref]
  » Crossref
• ⁴⁴ Khan, R.; Tariq, M.; Shaaban, I. A.; Assiri, M. A.; Bhatti, M. H.; Asif, H. M.; Polyhedron 2023, 243, 116529. [Crossref]
  » Crossref
• ⁴⁵ Manna, A.; Chakravorti, S.; J. Phys. Chem. B 2012, 116, 5226. [Crossref]
  » Crossref
• ⁴⁶ Swathi, M.; Shankar, D. S.; Daravath, S.; Ganji, N.; Lakshmi, P. V. A.; Shivaraj; Inorg. Chem. Commun. 2023, 153, 110826. [Crossref]
  » Crossref
• ⁴⁷ Neha, K. N.; Anal. Biochem. 2023, 675, 115216. [Crossref]
  » Crossref
• ⁴⁸ Yildiz, U.; Coban, B.; J. Serb. Chem. Soc. 2019, 84, 563. [Crossref]
  » Crossref
• ⁴⁹ Wurthner, F.; Kaiser, T. E.; Saha-Moller, C. R.; Angew. Chem., Int. Ed. 2011, 50, 3376. [Crossref]
  » Crossref
• ⁵⁰ Tisoco, I.; Köhler, M. H.; Nogara, P. A.; Rocha, J. B. T.; do Nascimento, C. S.; Garcia, R. Q.; De Boni, L.; Iglesias, B. A.; Inorg. Chim. Acta 2023, 556, 121639. [Crossref]
  » Crossref
• ⁵¹ Waring, M. J.; J. Mol. Biol. 1965, 13, 269. [Crossref]
  » Crossref
• ⁵² Sahoo, D.; Bhattacharya, P.; Chakravorti, S.; J. Phys. Chem. B 2010, 114, 2044. [Crossref]
  » Crossref

Edited by

• Editor handled this article: Fernando C. Giacomelli (Associate)

Publication Dates

• Publication in this collection
  07 Oct 2024
• Date of issue
  2025

History

• Received
  09 Apr 2024
• Accepted
  20 Sept 2024