Green-light responsive fluorescein-based photoremovable protecting group: nanoparticle formulation for controlled release of bioactive molecules with real-time-monitoring ability

Abstract

Dye-based photoremovable protecting groups (PRPGs) are explored for biological applications because they release bioactive molecules by absorbing light at higher wavelengths, and their self-fluorescent properties make them suitable for cellular imaging and image-guided photorelease inside the cells. Henceforth, we modified fluorescein dye to a cinnamyl-based PRPG for the release of alcohols to overcome the limitations of multiple photoproduct formation. The carboxylic acid group at C1 and the phenolic-OH group at the C6 positions in the fluorescein PRPG resulted in interesting pH-sensitive photophysical properties due to their existence in different forms (lactone, quinoid, monoanionic, dianionic) at different pHs, which is well supported by theoretical studies. Caged esters (3a–e) of fluorescein-based PRPG released the corresponding alcohols with good chemical yields and moderate photouncaging quantum yields upon exposure to green light. To enhance the biological utility, our developed fluorescein PRPG was formulated as nanoparticles (Nano-3d) having better cell penetration and accumulation. Interestingly, the fluorescein-based PRPG exhibited a change in fluorescence after photorelease ensuring its real-time monitoring ability in biological media. Furthermore, green light (525 ± 5 nm) exposure of our prepared nanoparticles (Nano-3d) released the bioactive molecule menthol within the MCF-7 breast cancer cell line causing effective cytotoxicity after photorelease. Hence, this development of a fluorescein-based PRPG can contribute to advancements in dye-based image-guided nanodrug delivery systems.

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Introduction

Photoremovable protecting groups (PRPGs) have gained significant attention in the areas of bioscience and material science^1–3 due to their ability to release bioactive molecules in a spatio-temporal controlled manner. In recent times, researchers have made reasonable contributions to improve the suitability of PRPGs for the release of bioactive molecules like drugs, neurotransmitters, gasotransmitters and nucleic acids at longer wavelengths.^4,5 In particular, PRPGs integrated with the ability to sense metal ions,⁶ pH,⁷ peroxide,⁸ and biological markers,⁹ have further amplified their potential, making them even more intriguing for biological applications. In the quest for promising chromophores as PRPGs, dye molecules are recognised for their ability to absorb light of longer wavelengths and for sensing applications due to the presence of in-built functionalities like carboxylic acid, phenolic-OH, or amine groups on their structures.^10–13 In addition, the intense fluorescence of these dyes also makes them useful in cellular imaging and as contrasting agents.^14,15 Interestingly, to date, only a few dyes have been reported as PRPGs, which include BODIPY,¹⁶ rhodamine,¹⁷ cyanin,¹⁸ naphthyridine-BODIPY,¹⁹ and xanthene.²⁰

After its synthesis by Bayer in 1871, fluorescein dye received great attention because of its versatile applicability in cellular imaging due to its promising fluorescent properties.²¹ The extensive application of fluorescein as a fluorescent probe highlights its biocompatibility.^22–24 The pH sensitivity of fluorescein makes it sensational, as changes in the pH of the solvent medium lead to the existence of different forms, a phenomenon first reported by Zanker and Peter.²⁵ The carboxylic acid and phenol groups at the C1 and C6 positions of the fluorescein molecule allow it to adopt various forms due to lactonization of the –COOH group and the protonation or deprotonation of both the carboxyl and phenolic-OH groups. Depending on the pH of the solvent medium, fluorescein dye exists in lactone, quinoid, monoanionic, and dianionic forms (Scheme 1).²⁶ Also, the non-emissive lactone and monoanionic forms, and emissive quinoid and dianionic forms of fluorescein dye show pH-dependent fluorescence on–off properties. Because of the above-mentioned unique fluorescent properties, fluorescein dye has been utilised for sensitive detection, precise measurements, and real-time monitoring of chemical processes, making it useful for different techniques such as colourimetry,²⁷ titrimetry,²⁸ and sensing.²⁹ In particular, green light absorption and less photobleaching make fluorescein dye more biocompatible. The green light absorption minimises the cell damage due to light, and the mitigation of photobleaching reduces the cytotoxicity.³⁰ The above-discussed interesting photophysical properties of fluorescein inspired us to utilise them as a green light-activated PRPG.

Scheme 1 Different forms of fluorescein dye depending on pH.

Previously, Sebej et al. reported a fluorescein analogue as a photoremovable group for carboxylates and phosphates (Fig. 1a).³¹ However, their synthesised fluorescein PRPG was a complex with DDQ instead of the native form, and photolysis resulted in two photoproducts along with the release of caged molecules, limiting its applicability in biological studies. Porter and his group first reported o-hydroxy cinnamyl-based photorelease for the direct photouncaging of alcohols and amines (Fig. 1b).^32,33 This photorelease mechanism resulted in the formation of a single photoproduct with a moderate to high photouncaging quantum yield, which improves its suitability for biological applications. Hence, we engineered the fluorescein chromophore by introducing a cinnamyl ester at the C5 position, enabling the direct release of caged alcohols upon green light activation via the o-hydroxy cinnamyl mechanism (Fig. 1c). In addition, we explored both experimentally and theoretically the pH dependency of the absorption and emission spectra of our model caged ester 3c. To expand its therapeutic utility, we formulated organic nanoparticles (Nano-3d) of caged ester 3d as a delivery system for the release of bioactive molecules such as menthol to check its cellular internalization and cell viability.

Fig. 1 (a) Sebej's work about a fluorescein analogue-based photoremovable protecting group. (b) Porter's work based on the o-hydroxy cinnamyl photorelease mechanism. (c) The present work based on fluorescein as a photoremovable protecting group.

Experimental section

General procedure for the synthesis of caged esters (3a–e)

Compound 2 (100 mg, 0.28 mmol) was dissolved in dry THF. In the solution, RCOCH=PPh₃ was added, and the reaction mixture was stirred at room temperature overnight. Then, the product was purified by column chromatography to obtain red-coloured solid products.

(E)-2-(5-(3-Ethoxy-3-oxoprop-1-en-1-yl)-6-hydroxy-3-oxo-3H-xanthen-9-yl)benzoic acid (3a)

To a solution of compound 2 (100 mg, 0.28 mmol) in 15 mL dry THF, ethyl 2-(triphenyl-λ⁵-phosphaneylidene)acetate (111 mg, 0.34 mmol) was added at room temperature and stirred overnight. The product was purified using column chromatography (EtOAc : hexane = 1 : 1) to get a red coloured solid product (yield 52%). ¹H NMR (500 MHz, DMSO) δ 11.11 (s, 1H), 10.19 (s, 1H), 8.16 (d, J = 16.3 Hz, 1H), 8.00 (d, J = 7.6 Hz, 1H), 7.80 (t, J = 7.4 Hz, 1H), 7.72 (t, J = 7.5 Hz, 1H), 7.31 (d, J = 7.6 Hz, 1H), 7.06 (d, J = 16.3 Hz, 1H), 6.82–6.69 (m, 2H), 6.64 (d, J = 8.8 Hz, 1H), 6.59 (s, 2H), 4.24 (q, J = 7.1 Hz, 2H), 1.31 (t, J = 7.1 Hz, 3H). ¹³C NMR (126 MHz, DMSO) δ 168.5, 167.1, 159.5, 159.3, 152.2, 151.2, 150.4, 135.6, 134.2, 130.4, 130.1, 128.9, 126.0, 124.6, 124.0, 120.9, 113.2, 112.4, 109.9, 109.3, 108.6, 102.1, 82.7, 60.0, 14.3. HRMS (ESI) m/z: [M + H]⁺ calcd for C₂₅H₁₈O₇ 430.1053; found 430.1049.

(E)-2-(5-(3-(Cyclohexyloxy)-3-oxoprop-1-en-1-yl)-6-hydroxy-3-oxo-3H-xanthen-9-yl)benzoic acid (3b)

To a solution of compound 2 (100 mg, 0.28 mmol) in 15 mL dry THF, cyclohexyl 2-(triphenyl-λ⁵-phosphaneylidene)acetate (129 mg, 0.34 mmol) was added at room temperature and stirred overnight. The product was purified using column chromatography (EtOAc : hexane = 1 : 1) to get a red coloured solid product (yield 53%). ¹H NMR (400 MHz, DMSO) δ 11.11 (s, 1H), 10.21 (s, 1H), 8.12 (d, J = 16.3 Hz, 1H), 7.96 (d, J = 7.6 Hz, 1H), 7.76 (t, J = 7.2 Hz, 7.6 Hz, 1H), 7.68 (t, J = 7.2 Hz, 7.6 Hz, 1H), 7.27 (d, J = 7.6 Hz, 1H), 6.98 (d, J = 16.2 Hz, 1H), 6.69 (d, J = 8.8 Hz, 2H), 6.66–6.51 (m, 3H), 4.80 (d, J = 4.0 Hz, 1H), 1.77 (d, J = 63.1 Hz, 4H), 1.51–1.22 (m, 6H). ¹³C NMR (101 MHz, DMSO) δ 168.7, 166.6, 159.6, 159.4, 152.4, 151.3, 150.4, 135.7, 134.1, 130.5, 130.2, 129.1, 126.1, 124.7, 124.1, 121.4, 113.3, 112.5, 109.9, 109.3, 108.6, 102.1, 82.8, 72.0, 31.3, 25.0, 23.4. HRMS (ESI) m/z: [M + H]⁺ calcd for C₂₉H₂₄O₇ 484.1522; found 484.1517.

(E)-2-(5-(3-(4-Bromophenoxy)-3-oxoprop-1-en-1-yl)-6-hydroxy-3-oxo-3H-xanthen-9-yl)benzoic acid (3c)

To a solution of compound 2 (100 mg, 0.28 mmol) in 15 mL dry THF, 4-bromophenyl 2-(triphenyl-λ⁵-phosphaneylidene)acetate (153 mg, 0.34 mmol) was added at room temperature and stirred overnight. The product was purified using column chromatography (EtOAc : hexane = 1 : 1) to get a red coloured solid product (yield 60%). ¹H NMR (400 MHz, DMSO) δ 11.30 (s, 1H), 10.17 (s, 1H), 8.29 (d, J = 16.2 Hz, 1H), 7.96 (d, J = 7.5 Hz, 1H), 7.76 (t, J = 7.6 Hz, 6.8 Hz, 1H), 7.68 (t, J = 7.6 Hz, 6.8 Hz, 1H), 7.61 (d, J = 8.2 Hz, 2H), 7.30–7.19 (m, 4H), 6.76 (s, 1H), 6.72 (d, J = 8.7 Hz, 1H), 6.65 (d, J = 8.8 Hz, 1H), 6.55 (s, 2H). ¹³C NMR (126 MHz, DMSO) δ 168.7, 165.8, 159.9, 159.7, 152.4, 151.2, 150.7, 150.0, 136.5, 135.8, 132.4, 131.2, 130.3, 129.1, 126.1, 124.8, 124.4, 124.1, 119.4, 118.2, 113.4, 112.6, 109.9, 109.3, 108.5, 102.3, 99.6. HRMS (ESI) m/z: [M + H]⁺ calcd for C₂₉H₁₇BrO₇ 556.0158; found 556.0150.

(E)-2-(6-Hydroxy-5-(3-((2-isopropyl-4-methylcyclohexyl)oxy)-3-oxoprop-1-en-1-yl)-3-oxo-3H-xanthen-9-yl)benzoic acid (3d)

To a solution of compound 2 (100 mg, 0.28 mmol) in 15 mL dry THF, 2-isopropyl-4-methylcyclohexyl 2-(triphenyl-λ⁵-phosphaneylidene)acetate (148 mg, 0.34 mmol) was added at room temperature and stirred overnight. The product was purified using column chromatography (EtOAc : hexane = 1 : 1) to get a red coloured solid product (yield 55%). ¹H NMR (500 MHz, DMSO) δ 8.09 (d, J = 16.2 Hz, 1H), 7.94 (d, J = 7.5 Hz, 1H), 7.69 (t, J = 7.3 Hz, 1H), 7.62 (t, J = 7.4 Hz, 1H), 7.20 (d, J = 7.3 Hz, 1H), 7.01 (d, J = 16.1 Hz, 1H), 6.71–6.59 (m, 2H), 6.54 (d, J = 13.2 Hz, 3H), 1.91 (d, J = 11.7 Hz, 1H), 1.82 (d, J = 7.6 Hz, 1H), 1.61 (d, J = 10.2 Hz, 2H), 1.48–1.38 (m, 2H), 1.09 (d, J = 6.4 Hz, 4H), 0.87–0.81 (m, 6H), 0.73 (d, J = 6.9 Hz, 3H). ¹³C NMR (126 MHz, DMSO) δ 168.5, 166.6, 159.5, 159.3, 152.3, 151.3, 150.3, 135.5, 134.1, 130.2, 130.1, 128.9, 126.0, 124.6, 124.0, 121.2, 113.2, 112.4, 109.9, 109.3, 108.6, 102.1, 73.3, 46.6, 40.7, 33.7, 30.9, 26.3, 23.5, 21.8, 20.3, 16.7. HRMS (ESI) m/z: [M + H]⁺ calcd for C₃₃H₃₂O₇ 540.2148; found 540.2182.

(E)-2-(6-Hydroxy-5-(3-((13-methyl-17-oxo-7,8,9,11,12,13,14,15,16,17-decahydro-6H-cyclopenta[a]phenanthren-3-yl)oxy)-3-oxoprop-1-en-1-yl)-3-oxo-3H-xanthen-9-yl)benzoic acid (3e)

To a solution of compound 2 (100 mg, 0.28 mmol) in 15 mL dry THF, 13-methyl-17-oxo-7,8,9,11,12,13,14,15,16,17-decahydro-6H-cyclopenta[a]phenanthren-3-yl 2-(triphenyl-λ⁵-phosphaneylidene)acetate (186 mg, 0.34 mmol) was added at room temperature and stirred overnight. The product was purified using column chromatography (EtOAc : hexane = 1 : 1) to get a red coloured solid product (yield 49%). ¹H NMR (500 MHz, DMSO) δ 8.32 (d, J = 16.2 Hz, 1H), 8.02 (d, J = 7.6 Hz, 1H), 7.80 (t, J = 7.1 Hz, 1H), 7.73 (t, J = 7.5 Hz, 1H), 7.36 (d, J = 8.6 Hz, 1H), 7.32 (d, J = 7.6 Hz, 1H), 7.28 (d, J = 16.2 Hz, 1H), 7.00 (dd, J = 8.4, 2.2 Hz, 1H), 6.96 (s, 1H), 6.79 (s, 1H), 6.73 (d, J = 8.9 Hz, 1H), 6.67 (d, J = 8.9 Hz, 1H), 6.61 (s, 2H), 2.93–2.87 (m, 2H), 2.29 (t, J = 8.8 Hz, 1H), 2.08 (m, J = 18.4, 8.6 Hz, 1H), 2.01–1.96 (m, 2H), 1.80 (d, J = 11.7 Hz, 1H), 1.62–1.56 (m, 2H), 1.55–1.46 (m, 2H), 1.43 (d, J = 10.9 Hz, 2H), 1.11 (t, J = 5.9 Hz, 2H), 0.86 (s, 3H). ¹³C NMR (126 MHz, DMSO) δ 169.0, 166.6, 155.5, 151.9, 151.5, 149.0, 138.3, 137.6, 136.6, 135.8, 133.6, 132.5, 132.0, 131.9, 131.3, 130.6, 129.5, 129.3, 129.2, 126.8, 126.5, 125.4, 124.8, 122.1, 119.8, 119.4, 110.2, 110.0, 109.0, 102.7, 50.1, 47.8, 44.1, 38.0, 35.9, 31.8, 29.4, 26.3, 25.9, 21.6, 14.0. HRMS (ESI) m/z: [M + H]⁺ calcd for C₅₁H₅₄O₉ 810.3768; found 810.3917.

Preparation of nanoparticles

The nanoparticles of caged ester 3d were prepared by following the reprecipitation technique. 1.1 mg of caged ester 3d was dissolved in 2 ml of DMSO to obtain 1 mM stock solutions. To 4 mL of distilled water, 40.5 μL of DMSO solution of 3d was added dropwise for 30 min with constant sonication to get a 10⁻⁵ M solution of nanoparticles of the caged ester (3d), Nano-3d.

Cell culture

Human breast cancer cells (MCF-7) and human embryo kidney cells (HEK293) were procured from National Center for Cell Sciences (Pune, India) and cultured in RPMI medium supplemented with 10% fetal bovine serum (FBS; Life Technologies), and 1% penicillin/streptomycin (Life Technologies) at 37 °C in a humidified atmosphere of 5% CO₂.

Cellular internalisation study of Nano-3d in MCF-7 cells

MCF-7 cells were seeded at a density of 5000 cells per cover glass bottom dish one day before the treatment. Then, the cells were treated with Nano-3d (10 μM) for 24 hours. The cells were washed with fresh medium followed by irradiation with a green LED (525 ± 5 nm) for 10 min. There was no light treatment in the without light treatment group. Both the cells with and without light treatment were imaged in confocal laser fluorescence scanning microscopy (CLFSM).

Cell viability study

Cell viability of the Nano-3d and menthol were measured using the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay. 5 × 10³ cells per well were seeded in a 96-well plate overnight. Then, different concentrations of the compounds were treated for 10 min with and without green LED (525 ± 5 nm) light irradiation. After that, MTT solutions were treated for 4 hours. The crystallized formazan was dissolved in a 1 : 1 mixture of dimethyl sulfoxide and methanol, and the absorbance was measured at 595 nm with an ELISA plate reader. The results are expressed as percent viability = [(A₅₉₅(treated cells) − background)/(A₅₉₅(untreated cells) − background)] × 100.

Results and discussion

The caged esters of the fluorescein PRPG were synthesised in two steps. At first, fluorescein aldehyde (2) was synthesised from commercially available fluorescein dye (1) by the Reimer–Tiemann reaction in methanol at 55 °C.²⁷ In the second step, the Wittig reaction was performed on 2 in dry THF at room temperature, resulting in the desired aliphatic caged esters (3a, 3b, 3d) and aromatic caged esters (3c, 3e) (Fig. 2a). Furthermore, to confirm the isomeric form (E or Z) of our synthesised caged esters (3a–e), we calculated the coupling constant values of the alkene protons of the caged esters. As a representative example, the alkene protons of caged ester 3c (Fig. 2b) showed a doublet with the chemical shifts at 8.31–8.27 ppm and 7.24–7.20 ppm with coupling constant values of 16 Hz, confirming that the caged ester 3c existed in its (E)-isomer. Detailed synthetic procedures and the characterization of all caged esters (3a–e) by ¹H NMR, ¹³C NMR, and HRMS are provided in the ESI† (Fig. S1–S12).

Fig. 2 (a) Synthetic scheme of caged esters (3a-e). (b) ¹H NMR of caged ester 3c (400 MHz, DMSO-d₆).

According to the literature, fluorescein dye exists in different forms depending on the pH of the solution, where the neutral forms (L, Q) are predominant at pH 4.6, the monoanionic form (M) is predominant at pH 6.5, and the dianionic form (D) is predominant at pH 8 (Scheme 1).²⁹ This prompted us to investigate the pH-dependent absorption and emission spectra of our newly developed caged esters of fluorescein PRPG.

As a representative example, the pH-dependent absorption and emission of the caged ester 3c [10⁻⁵ M solution of ACN/PBS buffer (1 : 1)] at three different pHs 4.6, 6.5, and 8 were recorded (Fig. 3a-b) and tabulated (Table 1). Also, the steady-state absorption and emission spectra of all the caged esters (3a–e) are presented in the ESI† (Fig. S13 and Table S1).

Fig. 3 (a) Absorption, and (b) emission spectra (λ_ex = 480 nm) of caged ester 3c in ACN : PBS buffer = 1 : 1, 10⁻⁵ M solution at different pHs.

Table 1 Photophysical properties of 3c at pH 4.6, 6.5, and 8

pH	Absorption		Emission
	λ max (nm)	(εmax × 10^4)^b	λ max (nm)	Stokes shift^d (nm)	(Φf)^e
a Maximum absorption wavelength. b Molar absorption coefficient (M^−1 cm^−1). c Maximum emission wavelength (λex = 480 nm). d Difference between maximum absorption wavelength and maximum emission wavelength. e Fluorescence quantum yield (error limit within ±10%).
4.6	311	1.89	—	—	—
	480	0.21
6.5	317	1.68	543	33	0.26
	381	1.20
	510	4.36
8	317	1.85	543	33	0.32
	390	1.31
	510	5.87

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We observed that 3c showed three absorption bands, 317 nm, 381 nm and 510 nm, at pH 6.5 and pH 8. Additionally, 3c showed two absorption bands, 311 nm and 480 nm, at pH 4.6 (Fig. 3a). The recorded emission spectra of 3c (λ_ex = 480 nm) showed only one emission maximum (542 nm) at pH 6.5 and 8 (Fig. 3b). At pH 4.6, 3c was found to be non-emissive (Fig. S14, ESI†).

Furthermore, to support our experimental observations, we calculated the theoretical absorption and emission spectra of 3c by TD-DFT calculation using ORCA 5.0.3 software. TD-DFT calculations revealed that the absorption band at 311 nm in pH 4.6 appeared due to the predominant existence of the lactone (L) form (Fig. S26a, ESI†), whereas the absorption band at 480 nm suggested the presence of the quinoid (Q) form (Fig. S26b, ESI†). On the other hand, at pH 6.5 and pH 8, the absorption bands at 317 nm, 381 nm, and 510 nm indicated the existence of monoanionic (M) and dianionic (D) forms, respectively (Fig. S26c and d, ESI†). Again, the computationally simulated emission spectrum for dianionic species (D) of 3c, around 527 nm, closely matches the recorded emission spectra (Fig. S27, ESI†). The results signified the possibility of the existence of a dianionic (D) form in the excited state that causes the emission at 525 nm (at pHs 6.5 and 8).³⁴

Next, 10⁻⁵ M solutions (ACN/PBS buffer of pH 6.5) of caged esters (3a–e) were irradiated with a 525 ± 5 nm LED lamp (40 W) to check the photouncaging ability of fluorescein PRPG. The photouncaging processes for caged esters (3a–c) were monitored by ¹H NMR spectroscopy (Fig. S17, ESI†). The uncaging quantum yield was calculated using a relative method with fulgide as an actinometer (Fig. S15, ESI†).³⁵ In all cases, photo-uncaging of alcohols was achieved with a high chemical yield of 70–95% and a moderate photouncaging quantum yield ranging from 0.002 to 0.025 (Table 2).

Table 2 Photochemical properties of caged esters 3a–e at pH 6.5

Caged esters	(Φu)^a	Uncaging yield^b (%)
a Photorelease quantum yield (error limit within ±10%). b Percentage of uncaging of caged alcohols from caged esters (3a–e).
3a	0.003	90
3b	0.003	86
3c	0.020	95
3d	0.003	78
3e	0.009	72

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As a representative example, the photouncaging ability of 3c (10⁻⁵ M solution) at pH 6.5 was monitored by reverse-phase high-performance liquid chromatography (RP-HPLC) at different time intervals. The RP-HPLC profile depicted that with the increase in the irradiation time, the peak at retention time, 9.7 min, corresponding to 3c, gradually decreased. On the contrary, two new peaks at retention times 4.7 min and 5 min (Fig. 4a and Fig. S16a, ESI†) gradually appeared. The peak at 4.7 min retention time corresponding to p-bromophenol was confirmed by injecting the authentic sample. The second peak at the retention time of 5 min corresponded to photoproduct (4), further characterised by ¹H-NMR and HRMS (Fig. S21 and S22, ESI†) after isolation from the photolysate. We further quantified the photouncaging process by RP-HPLC chromatogram peak area at regular time intervals and found that the photouncaging process followed first-order rate kinetics with a rate constant of 2.03 × 10⁻³ s⁻¹ (Fig. S16b, ESI†). We also quantified the photouncaging of 3c by measuring the decrease in the peak area at a retention time of 9.7 min and the increase in the peak area at a retention time of 5 min in the RP-HPLC chromatogram (Fig. 4c).

Fig. 4 Monitoring of photorelease for 3c using (a) RP-HPLC (mobile phase was ACN/water 3 : 2 (v/v) and UV detector 220 nm) and (b) ¹H NMR spectra (1.5 mg in 0.6 ml CD₃CN and 0.05 ml PBS buffer of pH 6.5). (c) Quantification of photouncaging of 3c during the course of photolysis. (d) Real-time monitoring study of 3c by emission spectra (10⁻⁵ M solution in ACN/PBS buffer 1 : 1) (inset: change in fluorescence colour before and after irradiation at 525 ± 5 nm). (e) Possible photorelease mechanism for fluorescein-based PRPG.

Furthermore, the photorelease capability of 3c was monitored using a time-dependent ¹H NMR study by irradiating 1.5 mg of 3c in CD₃CN/PBS buffer of pH 6.5 (0.6 ml/0.05 ml). With the increase of irradiation time, the intensity of the peaks corresponding to the (E)-olefinic protons of 3c, i.e., H_a and H_b at 8.1 ppm and 6.9 ppm (J = 16 Hz), gradually decreased with the simultaneous increase in the intensity of the new peaks at 8.2 ppm and 6.2 ppm (J = 10 Hz) corresponding to the (Z)-olefinic protons of the photoproduct (4) i.e., H_c and H_d. Also, the newly formed peaks at 7.0 ppm and 6.4 ppm indicated the release of caged alcohol (p-bromophenol) from 3cvia an (E)–(Z) photoisomerization process (Fig. 4b).

Interestingly, during the photolysis of 3c, we noted a prominent change in fluorescence from bright greenish yellow to less intense green, which is due to the formation of the photoproduct (4). Hence, we monitored the photolysis of 3c by emission spectroscopy (Fig. 4d), which showed that the intensity of the emission maximum of 3c at 542 nm decreased gradually, and the intensity of the newly formed emission maximum at 500 nm increased, and we also noted an isosbestic point at 512 nm.

Furthermore, the light ‘ON–OFF’ study was performed on 3c and monitored by RP-HPLC to check the temporal control over the release of caged alcohols (Fig. S18, ESI†). We found that light showed precise control over the release.

From all the experimental evidence, theoretical calculations, and literature studies,^32,33 we proposed the possible photorelease mechanism of caged esters (3a–e) (Fig. 4e). In the ground state, the caged esters (3a–e) at pH 6.5 existed as monoanionic forms (3a–e (M)). The quenching study using singlet state quencher perylene demonstrates that the photochemistry of 3a–e occurs from the singlet excited state (Fig. S19, ESI†) [the average lifetime of the singlet excited state of 3c measured by the time-correlated single photon counting (TCSPC) experiment was 2.73 ns (Fig. S20, ESI†)]. So, on irradiation with 525 ± 5 nm LED light, the monoanionic form of the caged esters (3a–e (M)) gets excited to their singlet excited state, and via the proton transfer process, these caged esters form an excited state (E) isomer of dianionic form (3a–e (E)). Then, (E)/(Z)-photoisomerization occurs, resulting in the conversion of (E)-isomers (3a–e (E)) into their corresponding (Z)-isomers (3a–e (Z)). These (Z)-isomers then undergo a thermally driven lactonisation process via nucleophilic attack of the C3 phenolic-OH. This lactonisation process forms the tetrahedral intermediates (3a–e (T)), which further release the caged alcohols along with the formation of photoproduct (4) confirmed by time-dependent ¹H NMR study. The photoproduct (4) was isolated and characterised (Fig. S21 and S22, ESI†).

The formation of a single photoproduct (4) with moderate uncaging quantum yield leads us to the synthesis of caged ester 3d, which can release the bioactive molecule menthol, known for anticancer activity,³⁶ upon irradiation with light. To improve the biological applicability (better cellular internalisation), we formulated caged ester 3d as nanoparticles (Nano-3d) using the reprecipitation technique.³⁷ We performed a high-resolution transmittance electron microscopy (HRTEM) to study the size and shape of Nano-3d and found nanoparticles were globular in shape with a diameter of 47 ± 10 nm (Fig. 5a). Dynamic light scattering (DLS) analysis suggested that the average hydrodynamic diameter for Nano-3d was in the range of 21–91 nm (PDI = 0.17 ± 0.05) (Fig. S23, ESI†). Then, to check the stability of the formulated nanoparticles of Nano-3d, the surface charge was determined by measuring the zeta potential (ζ), which was −41.9 ± 0.5 mV (Fig. 5b). Such a high negative value of zeta potential (ζ) indicated the presence of the hydroxyl and carboxylic acid groups on the surface and confirmed the stability of the Nano-3d. After that, we recorded the absorption and emission spectra for Nano-3d. The Nano-3d showed a decrease in intensity of the absorption band, whereas an increase in intensity for the emission band was observed compared to the bulk (Fig. 5c and d). Also, to monitor the change in shape during photolysis, we irradiated our Nano-3d with 525 ± 5 nm (40 W LED). From the HRTEM image, we found an increase in the particle size of Nano-3d with a diameter of 56 nm (Fig. 5a). This indicated the photodissociation of Nano-3d. Furthermore, to monitor the pH dependency of the nanoparticle formulation, we prepared the Nano-3d in different pH solutions (4.6, 6.5 and 8) and characterised them using DLS (Fig. S23b–d, ESI†), where the average hydrodynamic diameter and Z_avg increased with the increase of pH.

Fig. 5 (a) Schematic representation of the formulation of nanoparticles, and HR-TEM images of Nano-3d before and after irradiation with 525 ± 5 nm LED light for 30 min. (b) Measurement of zeta potential (ζ) for Nano-3d. (c) Absorption, and (d) emission spectra of Nano-3d and 3d with 10⁻⁵ M concentration.

To check the cellular internalisation of Nano-3d, we recorded confocal images of the MCF-7 breast cancer cell line after incubation with Nano-3d for 24 h. The cellular uptake was confirmed by observing the green fluorescence in the green channel, indicating the internalisation of Nano-3d in the MCF-7 breast cancer cell line (Fig. 6a). Irradiation with a green LED (525 ± 5 nm) for 5 min and 10 min resulted in a decrease in the green fluorescence (in the green channel) and an increase in the fluorescence (in the blue channel). This confirmed the release of the bioactive molecule menthol inside the MCF-7 breast cancer cell by Nano-3d upon exposure to green light.

Fig. 6 (a) Cellular internalisation studies of Nano-3d in the MCF-7 breast cancer cell line at different irradiation times (0, 5, 10 min) with a green LED (525 ± 5 nm) (scale bar: 10 μm): (i) bright field images, (ii) cells observed in the blue channel, (iii) cells observed in the green channel. (b) Cell viability studies of Nano-3d in the HEK293 normal cell line and MCF-7 breast cancer cell line before and after light irradiation (525 ± 5 nm).

After that, to check the biocompatibility of Nano-3d, we performed an MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay on the HEK293 normal cell line and the MCF-7 breast cancer cell line. Before irradiation, the Nano-3d showed 5% cell death in the HEK293 normal cell line, which indicated the biocompatibility of Nano-3d towards normal cells (Fig. 6b). Also, the calculated IC₅₀ values for the MCF-7 breast cancer cell line before and after green light irradiation were found to be 117 ± 18 μM and 64 ± 6 μM, respectively. This decrease in IC₅₀ value implied enhanced cytotoxicity of Nano-3d in cancer cells only upon light exposure.

Conclusion

We engineered fluorescein dye to a PRPG by converting it to a cinnamyl ester for the release of aliphatic and aromatic alcohols. The presence of –COOH and phenolic-OH groups at the C1 and C6 positions in fluorescein dye caused interesting pH-dependent photophysical properties by forming different forms (L, Q, M, and D). Our developed caged esters (3a–e) released caged alcohols upon irradiation with green light (525 ± 5 nm) with moderate photouncaging quantum yield via an (E)–(Z) photoisomerization process. Also, the single photoproduct formation after photorelease made the PRPG suitable for biological applications by minimizing the side effects. Interestingly, the change in fluorescence from bright greenish yellow to less intense green made the PRPG appropriate for image-guided delivery of bioactive molecules inside the cells. Hence, we prepared Nano-3d to release a bioactive molecule, menthol, to demonstrate its therapeutic utility inside cells. In vitro study confirmed the cellular internalisation and enhanced cytotoxicity upon light exposure.

To date, only a few dyes have been reported as PRPGs, highlighting their potential biological applications. Potential applications of these dye-based PRPGs can cover a broad area, such as targeted drug delivery systems for the controlled release of bioactive molecules in cancer treatment and wavelength-selective release for synergistic effects. Also, modifying NIR light-activated dyes like Si-rhodamine, Nile Red, and Nile Blue can engrave the pathway to real-time therapy in living organisms.

Author contributions

S. Pal conceptualized and designed the research project, synthesized the photocages, investigated photophysical and photochemical properties, conducted various experiments, and wrote the full manuscript. N. D. P. Singh supervised, validated all data, and assisted in manuscript writing. S. Paul conducted TD-DFT calculations. S. Biswas assisted in conducting the experiments. B. Jana carried out in vitro studies and contributed to data analysis. N. D. P. Singh was responsible for securing the funds.

Data availability

Data supporting this article have been uploaded as the ESI.†

Conflicts of interest

There are no conflicts to declare.

Acknowledgements

We express our gratitude to DST SERB for the financial support (Grant No. CRG/2022/000067) and DST (SR/FST/CSII-026/2013) for providing access to the 500 and 400 MHz NMR spectrometers. Our appreciation extends to the supercomputing facility “PARAM Shakti” at IIT Kharagpur, which was established under the National Supercomputing Mission (NSM) by the government of India. We acknowledge SATHI (Sophisticated Analytical Technical Help Institute) for providing access to HRTEM. We thank Professor Tomas Slanina for the fulgide compound that was used to determine the photon flask.

Notes and references

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Footnote

† Electronic supplementary information (ESI) available: Synthetic details; ¹H NMR, ¹³C NMR, and HRMS spectra; photophysical properties of caged esters; measurement of fluorescence quantum yields; measurement of photochemical quantum yields; photochemical rate constant determination; photorelease study of caged esters by ¹H NMR; fluorescence lifetime measurements; characterization of the photoproduct; singlet quenching study; DLS measurement; cell imaging; and computational data. See DOI: https://doi.org/10.1039/d5tb00388a

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