Push–pull fluorophores based on NHS esters of bithiophene for labelling of biomolecules containing primary amines

Abstract

Fluorescent labelling is a versatile tool to visualize biomolecules containing primary amines in their cellular environment, allowing the study of their function or interactions. Here, three organic fluorophores that can irreversibly bind to the primary amine group on the target biomolecule are reported. They consist of push-pull heterocyclic dyes based on bithiophene and incorporating a terminal N-hydroxysuccinimidyl ester as a reactive group for labelling primary amine groups from biomolecules as (poly)amines, peptides or proteins. Their potential as chemosensors for primary amines, using Nα-Boc protected amino acid l-lysine as a model, was assessed through UV- Visible, fluorescence and 1H NMR titrations.

Headnote

Keywords:

fluorescent labelling, push-pull heterocyclic fluorophores, bithiophene, Boc-lysine, primary amine chemosensor

Fluorescent labelling is a versatile tool to visualize biomolecules containing primary amines in their cellular environment, allowing the study of their function or interactions. Here, three organic fluorophores that can irreversibly bind to the primary amine group on the target biomolecule are reported. They consist of push-pull heterocyclic dyes based on bithiophene and incorporating a terminal N-hydroxysuccinimidyl ester as a reactive group for labelling primary amine groups from biomolecules as (poly)amines, peptides or proteins. Their potential as chemosensors for primary amines, using Nα-Boc protected amino acid l-lysine as a model, was assessed through UV- Visible, fluorescence and 1H NMR titrations.

1. Introduction

Biomolecules containing primary amines, such as (poly)amines, peptides and proteins, are essential for nearly every cellular function, and fluorescence-based imaging techniques provide powerful tools for investigating their activity in real time within living cells [1,2]. Such capabilities are crucial for advancing our understanding of cellular processes.

Over the past several decades, numerous fluorescent chemosensors have been developed, becoming indispensable tools for bioimaging and disease investigation [3]. Small-molecule fluorescent compounds are essential as labelling agents and activatable sensors because they are versatile, cost-effective, easy to handle and can achieve high signal-to-noise ratios through advanced chemical design [4]. Current fluorescence-based imaging technology heavily relies on small molecule-based dyes owing to their compact size, facile chemical modification, excellent reproducibility and biocompatibility [5]. Among the developed organic small molecule-based fluorescent dyes for biomolecule labelling are rhodamine [6], coumarin [7], fluorescein [8], BODIPY [9] and cyanine [10].

The abundance of reactive biogenic primary amines, i.e. the N-terminus and lysine residues, on protein surfaces allows for easy labelling of any protein with amine-reactive chemical groups [11]. The same principle can be applied to any biomolecule containing primary amine functional groups. It is well established that N-hydroxysuccinimidyl (NHS) esters can react with primary amino groups under physiological conditions. This reaction is highly efficient and specific, making it a preferred method for the labelling of biomolecules containing primary amines. By conjugating fluorescent dyes to NHS esters, it is possible to directly attach these dyes to biomolecules containing primary amines, allowing their visualization under fluorescence microscopy [12-14].

Based on our previous experience in the synthesis of push-pull heterocyclic dyes based on pyrrole and bithiophene [15-20], we decided to prepare several bithiophene-based push-pull fluorophores modified with an NHS ester as a reactive group (figure 1) to form an amide bond with the primary amino groups of biomolecules. Using Boc-lysine as a model system, we demonstrate the effectiveness and use of this labelling technique. To the best of our knowledge, this is the first example of the use of bithiophene-based push-pull fluorophores for the labelling of molecules via NHS esters.

2. Material and methods

2.1. General methods

The reagents and solvents used in the syntheses were purchased from Sigma-Aldrich and were used without further purification. Proton and 13C nuclear magnetic resonance (1H NMR and 13C NMR) spectra were recorded on Bruker Avance 300, 400 or 500 MHz equipment and referenced to the solvent peak at 25°C. The solvents are indicated in parentheses before the chemical shift values (δ relative to tetramethyl silane (TMS) and given in ppm). High-resolution mass spectra (HRMS) were recorded in the positive ion mode on an AB SCIEX Triple TOFTM 5600 liquid chromatography/mass spectrometry spectrometer. Fluorescence measures were taken with a Shimadzu RF-6000 spectrofluorophotometer using a 1 cm path length cuvette. UV-Visible (UV-Vis) absorption spectra were registered with a Shimadzu UV-2600 spectrophotometer, using a 1 cm path length cuvette.

2.2. Syntheses

2.2.1. Synthesis of bithienyl-pyrrole 5

Precursor 5 was synthesized through a Suzuki coupling reaction following a reported procedure [21].

2-Bromo-2,2'-bithiophene (3) (1 equiv.) was coupled with 1-methyl-1H-pyrrole-2-boronic acid pinacol ester 4 (1.2 equiv.) in a mixture of dimethoxyethane (DME, 10 ml), aqueous 2 M Na2CO3 (1 ml) and Pd(PPh3)4 (5 mmol%) at 80°C under nitrogen atmosphere. The reaction was monitored by thin-layer chromatography (TLC), which determined the reaction time (16 h). After cooling, the mixture was extracted with AcOEt (20 ml) and H2O (3 × 20 ml), and the phases were separated. The organic phase obtained was dried over anhydrous MgSO4, filtered and the solvent was evaporated under reduced pressure to give a crude mixture. The crude product was purified through a silica gel chromatography column using mixtures of dichloromethane and light petroleum of increasing polarity to afford product 5 as a dark green oil (131 mg, 66% yield). 1H NMR (400 MHz, CDCl3) δ 7.21 (dd, J = 5.1, 1.2, 1H), 7.17 (dd, J = 3.6, 1.2, 1H), 7.12 (d, J = 3.7, 1H), 7.04-6.99 (m, 2H), 6.92 (d, J = 3.7, 1H), 6.71 (t, J = 2.3, 1H), 6.35 (dd, J = 3.7, 1.4, 1H), 3.76 (s, 3H) ppm.

2.2.2. Synthesis of 7

Compound 7 was obtained through an azo coupling as previously reported [16].

Hydrochloric acid 37% (333 µl) was added to a round-bottom flask with a suspension of 4-aminobenzoic acid (0.067 g, 1.0 equiv.) in water (833 µl) and stirred at 0-5°C until the mixture was homogenous. NaNO2 (0.041 g, 1.2 equiv.) dissolved in water (666 µl) was slowly added to the well-stirred previous solution. The reaction mixture was stirred for 30 min at 0-5°C. A solution with 5 (0.120 g, 1 equiv.) was prepared in MeOH (3.333 ml) and pyridine (167 µl). The previously prepared diazonium salt solution was added dropwise to the solution of 5 at 0-5°C. The resulting mixture was stirred for 3 h at 0-5°C and then concentrated under reduced pressure. The precipitate was filtered, washed with cold water and dried to afford the pure heterocyclic azo derivative 7 as a grey solid (164 mg, 83% yield). 1H NMR (400 MHz, deuterated dimethyl sulfoxide (DMSO-d6)) δ 8.06 (d, J = 8.5, 2H), 7.86 (d, J = 8.6, 2H), 7.59 (dd, J = 5.1, 1.2, 1H), 7.57 (d, J = 3.9, 1H), 7.45 (d, J = 3.9, 1H), 7.43 (dd, J = 3.6, 1.2, 1H), 7.14 (dd, J = 5.1, 3.6, 1H), 6.95 (d, J = 4.5, 1H), 6.84 (d, J = 4.5, 1H), 4.13 (s, 3H) ppm.

2.2.3 General procedure for the synthesis of aldehydes 12a-b

Aldehydes 12a-b were prepared by Suzuki coupling as previously described [21].

5'-Bromo-2,2'-bithiophene-5-carboxaldehyde 11 (0.73 mmol) was coupled with boronic acids 9 and 10 (0.88 mmol) in a mixture of DME (10 ml), aqueous 2 M Na2CO3 (1 ml) and Pd(PPh3)4 (5 mol%) at 80°C under nitrogen atmosphere. The reactions were monitored by TLC, which determined the different reaction times (17-21 h). After cooling, the mixture was filtered and washed with water and diethyl ether to afford aldehydes 12a-b.

12a as an orange solid (213 mg, 93% yield) 1H NMR (400 MHz, CDCl3) δ 9.84 (s, 1H), 7.66 (d, J = 4.0, 1H), 7.53-7.47 (m, 2H), 7.30 (d, J = 3.9, 1H), 7.21 (d, J = 4.0, 1H), 7.11 (d, J = 3.9, 1H), 6.76-6.70 (m, 2H), 3.01 (s, 6H) ppm. HRMS: m/z calcd for C17H16NOS2 [M+H]+: 314.0673; found: 314.0675.

12b as an orange solid (172 mg, 54% yield) 1H NMR (400 MHz, acetone-d6) δ 9.92 (s, 1H), 7.93 (d, J = 4.0, 1H), 7.66-7.58 (m, 2H), 7.52 (d, J = 3.9, 1H), 7.47 (d, J = 4.0, 1H), 7.42 (d, J = 3.9, 1H), 7.40-7.30 (m, 4H), 7.16-7.07 (m, 6H), 7.09-7.01 (m, 2H) ppm. HRMS: m/z calcd for C27H19NOS2 [M]+: 437.0908; found: 437.0891.

2.2.4 General procedure for the synthesis of derivatives 14a-b

Cyanoacetic derivatives 14a-b were synthesized by a Knoevenagel condensation with cyanoacetic acid as previously described [16].

To a mixture of aldehyde 12 (1 equiv.) and cyanoacetic acid (13) (1.2 equiv.) in acetonitrile (ACN, 10 ml) was added one drop of piperidine. The reaction mixture was heated under reflux for 3-4 h and then cooled to 0°C. The precipitates obtained were filtered and washed with ethyl ether to give the pure products 14a-b.

14a as a red solid (120 mg, 63% yield) 1H NMR (400 MHz, DMSO-d6) δ 8.10 (s, 1H), 7.69 (d, J = 3.9, 1H), 7.52 (d, J = 8.9, 2H), 7.44 (d, J = 3.9, 1H), 7.41 (d, J = 3.9, 1H), 7.32 (d, J = 3.9, 1H), 6.75 (d, J = 8.9, 2H), 2.95 (s, 6H) ppm.

14b as a red solid (132 mg, 96% yield) 1H NMR (400 MHz, DMSO-d6) δ 8.03 (s, 1H), 7.68-7.58 (m, 3H), 7.50-7.40 (m, 3H), 7.34 (t, J = 7.9, 4H), 7.15-7.03 (m, 6H), 7.00-6.94 (m, 2H) ppm.

2.2.5 Synthesis of dyes 1 and 2a-b

In a typical procedure, a mixture of 7 or 14a-b (1 equiv.), NHS (5 equiv.) and N-(3-dimethylaminopropyl)-N'-ethylcarbodiimide hydrochloride (EDC HCl, 5 equiv.) in dichloromethane (DCM, 25 ml) was stirred at room temperature for 24 h. The mixture was extracted with H2O (3 × 10 ml), and the phases were separated. The organic phase obtained was dried over anhydrous MgSO4, filtered and the solvent was evaporated under reduced pressure. The pure dye was obtained by recrystalization from DCM/hexane.

Dye 1 as a dark red solid (125 mg, 69% yield) 1H NMR (500 MHz, DMSO-d6) δ 8.21 (d, J = 8.7, 2H), 7.99 (d, J = 8.7, 2H), 7.62-7.56 (m, 2H), 7.48-7.41 (m, 2H), 7.15 (dd, J = 5.1, 3.6 Hz, 1H), 6.94 (d, J = 4.5 Hz, 1H), 6.85 (d, J = 4.5 Hz, 1H), 4.16 (s, 3H), 2.91 (s, 4H) ppm. 13C NMR (125 MHz, DMSO-d6) δ 170.85, 131.95, 131.40, 129.08, 128.83, 126.75, 125.63, 125.28, 122.77, 113.98, 32.08, 26.03 ppm. HRMS: m/z calculated for C24H18N4O4S2 [M+H]+: 491.0848; found 491.0831.

Dye 2a as a burgundy solid (52 mg, 34% yield). 1H NMR (500 MHz, DMSO-d6) δ 8.79 (s, 1H), 8.18 (d, J = 4.3 Hz, 1H), 7.70 - 7.42 (m, 5H), 6.75 (d, J = 8.3 Hz, 2H), 2.97 (s, 6H), 2.88 (s, 4H) ppm. 13C NMR (125 MHz, DMSO-d6) δ 173.24, 170.50, 159.93, 151.16, 151.06, 149.31, 146.04, 133.33, 131.41, 130.63, 127.20, 125.36, 123.40, 120.64, 115.61, 112.74, 88.44, 25.99, 25.69 ppm. HRMS: m/z calculated for C24H19N3O4S2 [M+H]+: 478.0897; found 478.0864.

Dye 2b as a dark red solid (134 mg, 86% yield). 1H NMR (500 MHz, DMSO-d6) δ 8.82 (s, 1H), 8.19 (d, J = 4.3 Hz, 1H), 7.74 (d, J = 4.0 Hz, 1H), 7.69 (d, J = 4.2 Hz, 1H), 7.63 (dd, J = 9.9, 3.2 Hz, 2H), 7.52 (d, J = 4.0 Hz, 1H), 7.34 (ddd, J = 11.4, 7.4, 2.9 Hz, 4H), 7.13-7.04 (m, 6H), 6.95 (dd, J = 8.9, 2.4 Hz, 2H), 2.88 (s, 3H) ppm. 13C NMR (125 MHz, DMSO-d6) δ 173.23, 170.46, 159.80, 152.17, 151.24, 150.44, 147.03, 133.82, 133.09, 130.19, 127.29, 126.45, 125.84, 125.25, 124.36, 122.61, 89.23, 25.72 ppm. HRMS: m/z calculated for C34H23N3O4S2 [M+H]+: 602.1208; found 602.1173.

2.3. UV-Visible and fluorescence titrations with Nα-Boc protected amino acid L-lysine

In a 3 ml quartz cell (1.0 cm of path length), to 2800 µl of a 10 µM solution of the corresponding dye (1, 2a or 2b) in DMSO/H2O 99 : 1, increasing additions of 10 µl of Nα-Boc protected amino acid l-lysine (Boc-Lys-OH; 3 mM in aq NaHCO3 10 mM buffer, pH 8.4) were added. For each aliquot added, the solution was stirred for 5 s and then the corresponding UV-Vis absorption and fluorescence emission spectra (at the corresponding excitation wavelength) of the solution were recorded.

2.4. 1H NMR titrations

A known amount of the corresponding dye was dissolved in DMSO-d6 (ca 400 µl), in an NMR tube in such a way that the final concentration of the dye was 10 mM. Then, increasing amounts of Boc-Lys-OH in MeOD-d4 (10 µl aliquots, 0-4 equiv.) were added to the tubes, and finally, more DMSO-d6 was added to obtain a total volume of 500 µl. The solutions were shaken for 5 min at room temperature, and the corresponding 1H NMR spectra were recorded.

3. Results and discussion

3.1. Synthesis of dye 1

The first prepared dye based on bithiophene (compound 1) was the NHS ester derivative of the azo compound 7 (scheme 1). Compound 7 contains an electron-rich π-bridge/donating moiety on one side of the azo group and an electron-withdrawing benzoic acid group on the other side, which will be used later to bind NHS. On the other hand, the bithienylpyrrole precursor 5 was synthesized in 66% yield through a Suzuki cross-coupling reaction between the commercially available 5-bromo-2,2'-bithiophene 3 and 1-methyl-1H-pyrrol-2-yl-2-boronic acid 4 in DME under an inert atmosphere [21]. Diazotation of 4-aminobenzoic acid with NaNO2 in HCl at 0-5°C afforded the corresponding diazonium salt 6, which was further reacted with bithienylpyrrole 5 in methanol and pyridine at 0-5°C to give the heterocyclic azo dye 7, which was isolated by precipitation as a grey solid in 83% yield. The terminal carboxyl group of the azo derivative was transformed into the NHS ester by using NHS with 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC) as a coupling agent. The final new product was obtained with a 69% yield after recrystallization.

3.2. Synthesis of dyes 2a-b

Another family of bithiophene-containing push-pull dyes was synthesized (scheme 2). They consist of two push-pull molecular systems, containing an N,N-dialkylaniline as the donor moiety, connected through a bithiophene π-bridge to a cyanoacetic acid derivative as the electron acceptor moiety. The carboxyl group of the cyanoacetic acid was later transformed into an NHS ester as an anchoring group. Formyl compounds 12a-b incorporating a dialkylaniline group connected to the 2,2-bithiophene moiety were synthesized through a Suzuki cross-coupling reaction of commercially available aryl-boronic acids 9 and 10 with 5'-bromo-5-formyl-2,2'-bithiophene 11 in DME and aqueous 2 M Na2CO3, using Pd(PPh3)4 as a palladium catalyst, under an inert atmosphere. In this case, products 12a and 12b were obtained with respective yields of 93 and 54%, which may be understandable owing to the different electron-donating character of the dialkyl or diaryl substituent. These formyl precursors were converted into the correspondent cyanoacetic acid derivatives 14a-b by Knoevenagel condensation with 2-cyanoacetic acid in refluxing ACN in the presence of piperidine as a catalyst, in standard conditions as previously described [16] and with good yields (63 and 96%, respectively). The carboxyl group was then transformed into the NHS ester using NHS with EDC as a coupling agent. The final products were obtained with 34% (2a) and 86% (2b) yield, respectively.

The new dyes 1 and 2a-b were fully characterized using 1H and 13C NMR and HRMS.

3.3. Preliminary sensing experiments with Nα-Boc protected amino acid L-lysine

To evaluate the labelling of biomolecules containing primary amine groups through dyes 1 and 2a-b, a preliminary study was undertaken using Boc-Lys-OH as a model. The selection of this N-terminal protected amino acid with a side chain containing a primary amine group as a target molecule allowed for evaluating the capabilities of the synthesized dyes as chemical sensors.

For this purpose, UV-Vis and fluorescence titration studies with dyes 1 and 2a-b in the presence of increasing amounts of Boc-Lys-OH were performed. In a standard procedure, DMSO solutions of dyes 1 and 2a-b (10 µM) were treated with increasing amounts of Boc-Lys-OH in NaHCO3 buffer pH = 8.4 (10 mM), and the corresponding UV-Vis and fluorescence spectra were recorded (figures 2 and 3; electronic supplementary material, figure S11).

Dye 1 (10 µM in DMSO) presents an absorption maximum at 519 nm and an emission maximum at 623 nm when excited at the absorption maximum. The addition of Boc-Lys-OH leads to hypochromic and hypsochromic shifts with a new maximum at 484 nm in the UV-Vis spectra (figure 2a). An enhancement of fluorescence emission was observed at 611 nm in the presence of an excess of Boc-Lys-OH (figure 2b). Dye 2 a (10 µM in DMSO, NaHCO3 buffer, pH = 8.4) exhibits an absorption maximum at 387 nm. Upon adding increasing amounts of Boc-Lys-OH, the absorbance decreases in the peak at 387 nm, and a new maximum peak emerges at 464 nm. Fluorescence titration studies (λexc = 387 nm) were also carried out (figure 3b). In the absence of Boc-Lys-OH, compound 2a showed an emission maximum at 481 nm, which was quenched by the addition of increasing amounts of the amino acid.

Dye 2b (10 µM in DMSO) shows a maximum absorption at 439 nm and an emission maximum at 621 nm when excited at the absorption maximum (see the electronic supplementary material, figure S11). Of the synthesized dyes, dye 2b showed the smallest variations in its UV-Vis and fluorescence spectra in the presence of Boc-Lys-OH, indicating a less pronounced response compared to the other dyes studied.

To test the dyes in a more realistic biological environment, fluorescence titration experiments of dye 2a with BOC-Lys-OH were conducted in a 1 : 1 mixture of DMSO and Roswell Park Memorial Institute (RPMI) 1640 medium (a cell culture medium with l-glutamine and sodium bicarbonate). As observed in the electronic supplementary material, figure S14, the fluorescence emission maximum of 2a, which appears at 511 nm in the absence of Boc-Lys-OH, experiences a gradual quenching upon increasing additions of the amino acid, similar to that observed in DMSO.

Finally, 1H NMR investigations were also conducted with dyes 1 and 2a-b, using increasing amounts of Boc-Lys-OH (figures 4, 5; electronic supplementary material, figure S12). Dye 2b presented the same trend as for dye 2a. As anticipated, the formation of the amide bond resulted in an upfield chemical shift of the hydrogens close to the carbonyl moiety, especially of the aromatic protons on the benzene ring of dye 1 and the olefinic hydrogen of compounds 2a-b. Furthermore, it was noted that the protons associated with the NHS in ester form at 2.9 ppm disappeared, revealing a new singlet at 2.6 ppm characteristic of the free NHS.

4. Conclusions

Three push-pull heterocyclic dyes based on bithiophene, incorporating a terminal NHS ester as an anchoring group for irreversibly binding to an amino group of a biomolecule through the formation of an amide bond, were successfully synthesized and characterized. Their potential as chemosensors for biomolecules containing primary amines, using Boc-Lys-OH as a model amino acid incorporating a primary amino group in its side chain, was assessed through UV-Vis, fluorescence and 1H NMR titrations. UV-Vis and fluorescence titrations of the dyes in the presence of increasing amounts of Boc-Lys-OH showed significant changes in their UV-Vis and fluorescence spectra, particularly for dyes 1 and 2a. In addition, the potential of dye 2a as a chemosensor for primary amines was also assessed in a more competitive medium, such as RPMI 1640 cell culture medium, which showed a quenching of the fluorescence emission in the presence of increasing amounts of Boc-Lys-OH. The 1H NMR titrations provided valuable insights, allowing us to visualize the formation of the amide bond between the synthesized dyes and Boc-Lys-OH. The distinctive changes in the 1H NMR spectra confirmed the successful reaction. Future studies will focus on optimizing these dyes towards their interaction with primary amine-containing proteins and studying their labelling properties using confocal microscopy.

Ethics. This work did not require ethical approval from a human subject or animal welfare committee.

Data accessibility. The data supporting this article have been uploaded as part of the eletronic supplementary material [22].

Declaration of AI use. We have not used AI-assisted technologies in creating this article.

Authors' contributions. M.B.: investigation, methodology, writing-original draft; P.A.: formal analysis, validation; J.A.S.: validation, writing-original draft; S.G.: formal analysis, validation; M.P.: methodology, supervision; S.P.G.C.: methodology, supervision, writing-review and editing; M.M.M.R.: conceptualization, funding acquisition, methodology, supervision, writing-review and editing; P.G.: conceptualization, funding acquisition, supervision, writing-original draft, writing-review and editing.

All authors gave final approval for publication and agreed to be held accountable for the work performed therein.

Conflict of interest declaration. We declare we have no competing interests.

Funding. The authors gratefully acknowledge grant PID2021-126304OB-C42 funded by the Spanish MCIN/AEI/ 10.13039/501100011033 and by "ERDF A way of making Europe, EU", grant PDC2022-133576-C22 funded by the Spanish MCIN/AEI/10.13039/501100011033 and by the European Union "NextGenerationEU"/PRTR" and grant CIAICO/2022/146 funded by Generalitat Valenciana (Conselleria de Educación, Universidades y Empleo). Thanks are also due to Fundação para a Ciência e Tecnologia (FCT) and FEDER (European Fund for Regional Development)-COMPETE-QRENEU for financial support through the Chemistry Research Centre of the University of Minho (Ref. CQ/UM UID/QUI/00686/2020). The NMR spectrometer Bruker Avance III 400 is part of the National NMR Network (PTNMR) and is partially supported by Infrastructure Project No 022161 (co-financed by FEDER through COMPETE 2020, POCI and PORL and FCT through PID-DAC).

Sidebar

Cite this article: Barros M, Arroyo P, Sáez JA, Gil S, Parra M, Costa SPG, Raposo MMM, Gaviña P. 2025 Push-pull fluorophores based on NHS esters of bithiophene for labelling of biomolecules containing primary amines. R. Soc. Open Sci. 12: 241816.

https://doi.org/10.1098/rsos.241816

Received: 21 October 2024

Accepted: 2 January 2025

Subject Category:

Chemistry

Subject Areas:

organic chemistry, analytical chemistry

Authors for correspondence:

M. Manuela M. Raposo e-mail: [email protected]

Pablo Gaviña e-mail: [email protected]

This article has been edited by the Royal Society of Chemistry, including the commissioning, peer review process and editorial aspects up to the point of acceptance.

Electronic supplementary material is available online at https://doi.org/10.6084/ m9.figshare.c.7647014.

References

References

1. Kielland N, Vendrell M, Lavilla R, Chang YT. 2012 Imaging histamine in live basophils and macrophages with a fluorescent mesoionic acid fluoride. Chem. Commun. 48, 7401-7403. (doi:10.1039/c2cc32292g)

2. Sintes M, De Moliner F, Caballero-Lima D, Denning DW, Read ND, Kielland N, Vendrell M, Lavilla R. 2016 Electrophilic, activation-free fluorogenic reagent for labeling bioactive amines. Bioconjugate Chem. 27, 1430-1434. (doi:10.1021/acs.bioconjchem.6b00245)

3. Wu D, Sedgwick AC, Gunnlaugsson T, Akkaya EU, Yoon J, James TD. 2017 Fluorescent chemosensors: the past, present and future. Chem. Soc. Rev. 46, 7105-7123. (doi:10.1039/C7CS00240H)

4. Terai T, Nagano T. 2013 Small-molecule fluorophores and fluorescent probes for bioimaging. Pflugers Arch. 465, 347-359. (doi:10.1007/ s00424-013-1234-z)

5. Yin J, Huang L, Wu L, Li J, James TD, Lin W. 2021 Small molecule based fluorescent chemosensors for imaging the microenvironment within specific cellular regions. Chem. Soc. Rev. 50, 12098-12150. (doi:10.1039/d1cs00645b)

6. Si D, Li Q, Bao Y, Zhang J, Wang L. 2023 Fluorogenic and cell-permeable rhodamine dyes for high-contrast live-cell protein labeling in bioimagingandbiosensing.Angew.Chem.Int.Ed.62,e202307641.(doi:10.1002/anie.202307641)

7. Fan Y, Wu Y, Hou J, Wang P, Peng X, Ge G. 2023 Coumarin-based near-infrared fluorogenic probes: Recent advances, challenges and future perspectives. Coord. Chem. Rev. 480, 215020. (doi:10.1016/j.ccr.2023.215020)

8. Watanabe S, Mizukami S, Hori Y, Kikuchi K. 2010 Multicolor protein labeling in living cells using mutant β-lactamase-tag technology. Bioconjug. Chem. 21, 2320-2326. (doi:10.1021/bc100333k)

9. Murale DP, Hong SC, Yun J, Yoon CN, Lee JS. 2015 Rational design of a photo-crosslinking BODIPY for in situ protein labeling. Chem. Commun. 51, 6643-6646. (doi:10.1039/C5CC01068C)

10. Yapici I et al. 2015 'Turn-on' protein fluorescence: in situ formation of cyanine dyes. J. Am. Chem. Soc. 137, 1073-1080. (doi:10.1021/ja506376j)

11. Pham GH et al. 2018 Tuning a protein-labeling reaction to achieve highly site selective lysine conjugation. Chembiochem 19, 799-804. (doi:10. 1002/cbic.201700611)

12. Dempsey DR, Jiang H, Kalin JH, Chen Z, Cole PA. 2018 Site-specific protein labeling with N-hydroxysuccinimide-esters and the analysis of ubiquitinligasemechanisms.J.Am.Chem.Soc.140,9374-9378.(doi:10.1021/jacs.8b05098)

13. Jiang H, Cole PA. 2021 N-terminal protein labeling with N-hydroxysuccinimide esters and microscale thermophoresis measurements of proteinprotein interactions using labeled protein. Curr. Protoc. 1, e14. (doi:10.1002/cpz1.14)

14. Jiang H, D'Agostino GD, Cole PA, Dempsey DR. 2020 Methods in enzymology, pp. 333-353, vol. 639. Elsevier.

15. Fernandes SS, Herbivo C, Aires-de-Sousa J, Comel A, Belsley M, Raposo MMM. 2018 Theoretical and experimental studies of aryl-bithiophene based push-pull π-conjugated heterocyclic systems bearing cyanoacetic or rhodanine-3-acetic acid acceptors for SHG nonlinear optical applications. Dyes Pigm. 149, 566-573. (doi:10.1016/j.dyepig.2017.10.001)

16. Garcia-Amorós J, Castro MCR, Nonell S, Vílchez S, Esquena J, Raposo MMM, Velasco D. 2019 Adaptable photochromic switches with selfaggregatingheterocyclicazodyes.J.Phys.Chem.C123,23140-23144.(doi:10.1021/acs.jpcc.9b07527)

17. Fernandes SSM, Castro MCR, Ivanou D, Mendes A, Raposo MMM. 2022 Push-pull heterocyclic dyes based on pyrrole and thiophene: synthesis and evaluation of their optical, redox and photovoltaic properties. Coatings 12, 34. (doi:10.3390/coatings12010034)

18. Fernandes SS, Belsley M, Pereira AI, Ivanou D, Mendes A, Justino LLG, Burrows HD, Raposo MMM. 2018 Push-pull N,N-diphenylhydrazones bearing bithiophene or thienothiophene spacers as nonlinear optical second harmonic generators and as photosensitizers for nanocrystalline TiO2dye-sensitizedsolarcells.ACSOmega3,12893-12904.(doi:10.1021/acsomega.8b01045)

19. Fuse S, Sugiyama S, Maitani MM, Wada Y, Ogomi Y, Hayase S, Katoh R, Kaiho T, Takahashi T. 2014 Elucidating the structure-property relationships of donor-π-acceptor dyes for dye-sensitized solar cells (DSSCs) through rapid library synthesis by a one-pot procedure. Chemistry 20, 10685-10694. (doi:10.1002/chem.201402093)

20. Garcia-Amorós J, Castro MCR, Coelho P, Raposo MMM, Velasco D. 2013 New heterocyclic systems to afford microsecond green-light isomerisable azo dyes and their use as fast molecular photochromic switches. Chem. Commun. 49, 11427-11429. (doi:10.1039/c3cc46736h)

21. Herbivo C, Comel A, Kirsch G, Raposo MMM. 2009 Synthesis of 5-aryl-5'-formyl-2,2'-bithiophenes as new precursors for nonlinear optical (NLO) materials. Tetrahedron 65, 2079-2086. (doi:10.1016/j.tet.2008.12.078)

22. Barros M, Arroyo P, Sáez JA, Gil S, Parra M, Costa SPG et al. 2025 Supplementary material from: Push-pull fluorophores based on NHS esters of bithiophene for labelling of biomolecules containing primary amines. FigShare (doi:10.6084/m9.figshare.c.7647014)