The donor–acceptor complexes of quantum dots and ionic perylene diimides for ratiometric detection of double-stranded DNA

Abstract

This communication presents a new FRET donor–acceptor system based on hybrid inorganic–organic complexes formed by quantum dots (QDs) and perylene diimides (PDIs). The ratio of the emission of QDs and the FRET emission of PDIs changes as the ratio of the donor and the acceptor changes. This donor–acceptor system can be used for ratiometric detection of double-stranded DNA.

---

Dyes with excellent thermo- and photostability that emit in the visible light region with high quantum yields are useful as fluorescent probes in biological and biomedical research, life sciences, and clinical diagnostics.¹ Perylene diimides (PDIs) are one of these candidates that have been widely used in organic light-emitting diodes,² organic field-effect transistors,³ organic photovoltaic solar cells,⁴ and liquid crystals.⁵ However, their applications as fluorescent probes have only been minimally explored because most PDIs tend to aggregate in aqueous solution, forming non-fluorescent π–π stacks.⁶ To efficiently reduce the aggregation, the introduction of substituents on the perylene-core or large dendrons at the side chains is generally required.⁷ However these modifications require substantial synthesis and purification. Another approach to prevent aggregation of PDIs in aqueous solution is to form strong host–guest complexes with macrocyclic hosts such as cyclodextrin (CD), cucurbit[8]uril (CB[8]), and ExBox through non-covalent interactions.⁸ Supramolecular assembly and disassembly of host–guest complexes have been employed to develop fluorescent “on–off” or “off–on” probes for sensing small molecules (e.g. organophosphates, surfactants, and sulphides) and macromolecules.⁹

Herein, we report a new supramolecular approach to prevent aggregation of PDIs in aqueous solution by forming a strong donor–accepter complex of PDIs with negatively charged QDs through non-covalent electrostatic attractions and hydrophobic interactions (Scheme 1c).¹⁰ A previous study has shown very efficient QD photoluminescence (PL) quenching and energy transfer in QD–PDI nanoassemblies,¹¹ thus we anticipate QD : PDI complexes are capable of undergoing efficient fluorescence resonance energy transfer (FRET), where negatively charged luminescent QDs serve as FRET donors and positively charged fluorescent PDIs act as FRET acceptors. Upon excitation of QD : PDI complexes at 405 nm, energy can be transferred from QD to PDIs bound to the surface of the QD though non-covalent electrostatic interactions. FRET efficiency of QD : PDI complexes can be tuned by controlling the molar ratio of QDs and PDIs. In addition, the ratio of two emission intensities (the emission from the donor QD and the FRET emission from the accepter PDIs) changes as the molar ratio of QDs and PDIs changes. Previous studies have reported that PDIs bind strongly with DNA through non-covalent interactions (e.g. π–π interactions and electrostatic attractions) (Scheme 1b).¹² We hypothesized that dsDNA may compete effectively with QDs for binding to PDIs, and thus may allow QD : PDI complexes to serve as ratiometric probes for dsDNA (Scheme 1c). Since a ratiometric assay is based on the ratio of fluorescence emission intensities at two different wavelengths instead of the absolute fluorescence emission intensity at a single wavelength, errors due to inaccuracy in concentrations of fluorescent probes and fluctuation in the intensity of an excitation light source are minimized.¹³

Scheme 1 Synthesis, aggregation, deaggregation, and energy transfer diagram. (a) Synthesis of PDI-2; (b) deaggregation of PDI-2 by intercalating with double-stranded DNA; (c) deaggregation of PDI-2 by interacting with QDs and competition binding of dsDNA and QDs with PDI-2.

To apply this new deaggregation and FRET strategy, a water-soluble dicationic dye PDI-2 was designed and synthesized in two simple steps from perylene-3,4,9,10-tetracarboxylic dianhydride (PTCDA) outlined in Scheme 1a. PDI-2 is very soluble in polar organic solvents (e.g. methanol and DMSO), water and aqueous buffers (e.g. Tris–HCl and PBS buffer). The UV-vis spectrum of PDI-2 (5.0 × 10⁻⁶ M) in methanol showed characteristic features of three absorption bands with a highest peak at ∼535 nm indicating the presence of PDI-2 non-aggregated monomers in methanol. In contrast, the UV-vis spectra of PDI-2 (5.0 × 10⁻⁶ M) in water and Tris–HCl buffer showed a dominate peak at ∼500 nm and a small shoulder around 540 nm, suggesting aggregation of PDI-2 molecules in aqueous solution (Fig. 1a).¹⁴ It is known that the aggregation of PDIs in aqueous solution causes significant fluorescence quenching due to the formation of non-fluorescent aggregates via an excimer self-quenching mechanism.¹⁵ As expected, fluorescence intensity of PDI-2 (5.0 × 10⁻⁶ M) in water and Tris–HCl buffer is only about 12% and 5% respectively of that in methanol (Fig. S6†). The deaggregation of PDI-2 by double-stranded DNA was investigated using UV-vis spectroscopy. As shown in Fig. S7a,† the addition of dsDNA S1 : S2 to the buffer solution of PDI-2 caused an increase in the absorption band of monomers and a red spectral-shift to ∼550 nm, suggesting deaggregation of PDI-2 by DNA double helix intercalation, surface binding, or both.^13h In addition, the overall decrease in absorptivity suggested the stacking interactions between the bases and PDI-2 molecules (Fig. S6a†).¹⁶ The presence of dsDNA S1 : S2 as little as 20 nM caused a significant drop in the fluorescence emission of PDI-2 (80 nM) probably due to the static fluorescence quenching by DNA bases of S1 : S2 (Fig. S8†). The results shown above suggest that PDI-2 itself may be suitable for DNA assessment. However, the ratiometric assay presented in this manuscript is superior because it could minimize the errors due to inaccuracy in concentrations and fluctuation in the intensity of an excitation light source.

Fig. 1 UV-vis spectra of PDI-2 (5 × 10⁻⁶ M) (a) in methanol, water, and Tris–HCl buffer (pH 7.4); and (b) in the presence of QD-455 with a ratio from 0 to 1.0 equivalent in Tris–HCl buffer (pH 7.4).

To demonstrate deaggregation of PDI-2 by forming complexes with negatively charged QDs, we tested two commercially available QDs (QD-455 and QD-490) with same amphiphilic polymer surface coating, but different emission peaks centered at ∼455 nm and ∼490 nm, respectively. These polymer encapsulated CdSe/ZnS core/shell QDs with carboxylic acid groups are very stable in most aqueous buffer solutions in the pH range of 5–10 and can even survive autoclaving (121 °C for 30 min) and lyophilisation processes.¹⁷ In water or a buffer (pH 7.4), we anticipated that positively charged PDI-2 molecules would readily bind to the negatively charged surfaces of QDs. As shown in Fig. 1b, the signature band at ∼535 nm of PDI-2 monomers increased as the molar ratio of QD-455/PDI-2 increased from 0 to 1.0. The absorption spectrum of PDI-2 (5 × 10⁻⁶ M) with one equivalent of QD-455 in the range of 450 to 550 nm was similar to the spectrum of PDI-2 (5 × 10⁻⁶ M) in methanol, suggesting complete deaggregation of PDI-2 upon addition of one equivalent of QD-455 QDs. The fluorescence intensity of PDI-2 (5 × 10⁻⁶ M) enhanced about 13-fold in the presence of 1.0 equivalent of QD-455, confirming deaggregation of PDI-2 (Fig. 1b). In a similar experiment of QD-490 QDs, the absorption spectrum of PDI-2 showed characteristics of monomers in the presence of 1.0 equivalent of QD-490, indicating deaggregation of PDI-2 aggregates upon addition of QD-490 quantum dots (Fig. S7b†).

The primary requirement for FRET is that the emission energy of the donor (S₁–S₀) must match with the excitation energy of the accepter (S₀–S₁). In other words, the absorption spectrum of the acceptor must overlap with the emission spectrum of the donor, thus the FRET transfer rate is dependent on their overlap integral.¹⁸ As shown in Fig. 2a, PDI-2 absorbs visible light in a broad range from 420 to 600 nm and emits longer wavelength light in the range of 520 nm to 650 nm with a maximum peak at ∼545 nm. QD-455 quantum dots absorb violet-blue light and emit in the range of 420 nm to 520 nm. Therefore, the emission of QD-455 overlaps with the absorption of PDI-2 and it makes QD-455 and PDI-2 a suitable donor–acceptor pair for FRET energy transfer. In comparison, QD-490 quantum dots emit light in the range of 450 nm to 550 nm that overlaps ∼50% more with the absorption of PDI-2 compared to QD-455 QDs (Fig. S9a†). Thus, we anticipated that FRET would be observed in both QD-455 : PDI-2 and QD-490 : PDI-2 systems, but with a different FRET energy transfer rate. To demonstrate FRET between QDs and PDI-2, the fluorescence emission of QDs was investigated in the absence and presence of PDI-2. As shown in Fig. 2b, the emission intensity of QD-455 (2.0 × 10⁻⁸ M) at ∼455 nm decreased gradually as the molar ratio of PDI-2/QD increased and it dropped ∼45% and ∼70% as the molar ratio of PDI-2 increased to 10 and 30 (or 10 and 30 dye molecules per QD), respectively. Interestingly, FRET emission intensity at ∼545 nm increased initially as the molar ratio of PDI-2/QDs increased and the maximum intensity was achieved with a molar ratio of PDI-2/QDs around 4 to 6, then it decreased as the molar ratio of PDI-2/QDs continuously increased presumably due to enhanced self-quenching of PDI-2 at a high local concentration surrounding the surfaces of QDs. The FRET efficiency from QD-455 to PDI-2 was calculated based on the equation E = (F_D − F_DA)/F_D (where F_D and F_DA represent fluorescence intensities of the donor in the absence and presence of the acceptors, respectively). As shown in Fig. 2c, QD-455 PL emission intensity decreased as the molar ratio of PDI-2/QD-455 increased. As expected, the FRET efficiency increased as the molar ratio of PDI-2/QD-455 increased and an approximately 50% FRET efficiency was achieved when the molar ratio of PDI-2/QD-455 reached 15. In comparison, the study with QD-490 showed a higher FRET energy transfer rate and the FRET efficiency reached 50% when the molar ratio of PDI-2/QD-490 was 10 presumably due to the greater overlap integral between the emission of QD-490 and the absorption of PDI-2. As shown in Fig. 2d, the ratio of the PL emission (at 455 nm) of QD-455 and the FRET emission (at 545 nm) of PDI-2 decreased rapidly from ∼42 to ∼6 as the molar ratio of PDI-2/QD-455 increased from 0 to 4. As the molar ratio of PDI-2/QD-455 increased further, the FRET emission of PDI-2 decreased probably due to self-quenching of PDI-2 even though PL emission of QD-455 continuously decreased. Interestingly, the ratio of PL₄₅₅/FL₅₄₅ increased gradually from 6 to 10 as the molar ratio of PDI-2/QD-455 increased from 10 to 30 probably because the self-quenching efficiency is slightly greater than the FRET quenching efficiency. In comparison, the study of QD-490 with PDI-2 showed a constant ratio (∼6) of PL_{490 nm}/FL_{545 nm} as the molar ratio of PDI-2/QD-490 continuously increased from 10 to 30, suggesting that self-quenching efficiency is similar to the FRET quenching efficiency. This is consistent with the results shown in Fig. 2c where a higher FRET energy transfer rate was observed in the study of QD-490 QDs. In a control experiment, a perylene-based dye PDI-3 with two negative charges was designed and synthesized. Compared to PDI-2, PDI-3 showed similar optical properties in Tris–HCl buffer (Fig. S10†). As expected, PL emission of QD-455 (20 nM) at 455 nm remained almost the same in the presence of 1.0 to 5.0 equivalents of PDI-3. However a very small emission peak was observed around 545 nm in the presence of 5.0 equivalents of PDI-3 presumably due to either an emission of PDI-3 when excited directly at 405 nm or a weak FRET emission of PDI-3 that binds to the surface of QDs (Fig. S11†). These observations strongly suggested that there was very little binding of negatively charged PDI-3 dyes to the surface of QDs, probably due to charge repulsion.

Fig. 2 (a) FRET diagram. UV-vis molar absorptivity (dashed lines) of QD-455 (1 × 10⁻⁶ M) and PDI-2 (5 × 10⁻⁶ M). The absorptivity of PDI-2 was multiplied by 7. Emission spectra (solid lines, normalized) of QD-455 (2 × 10⁻⁸ M) and PDI-2 (1.0 × 10⁻⁶ M) in Tris–HCl buffer (pH 7.4) are shown; (b) emission spectra of QD-455 (2.0 × 10⁻⁸ M) in the absence and presence of PDI-2 with a molar ratio from 0 to 30. Inset: FRET fluorescence emission in the range from 500 to 750 nm; (c) plots show QD PL intensity and FRET efficiency vs. [PDI-2]/[QD]; (d) plots show the ratio PL_QD/FL_PDI-2 vs. [PDI-2]/[QD].

Further studies were performed to explore the ratiometric assay for the detection of double-stranded DNA using QD-455 : PDI-2 complexes. The PL emission of QD-455 (20 nM) in the presence of dsDNA was investigated. As shown in Fig. S12,† the presence of excess dsDNA S1 : S2 caused a ∼9% drop in PL emission intensity of QD-455 probably due to weak static quenching by DNA bases that bind to the surface of the QDs. Interesting to note, one could anticipate that the binding between DNA and QDs is very weak due to the competition of electrostatic repulsion and hydrophobic interactions.¹⁹ Since the minimum PL_455nm/FL_545nm ratio can be obtained when the molar ratio of PDI-2/QD-455 is about 4, solutions of complexes with four equivalents of PDI-2 and one equivalent of QD-455 were used in further studies with DNA. As shown in Fig. 3a, no significant changes were observed in either PL emission intensity (at ∼455 nm) or FRET emission intensity (at ∼545 nm) for QD-455 : PDI-2 (20 nM : 80 nM) complexes in Tris–HCl buffer (pH 7.4) in the presence of the ssDNA S1 (100 nM). In contrast, the presence of the dsDNA S1 : S2 (100 nM) caused a significant drop in the FRET emission intensity and an increase in PL emission intensity, suggesting competitive binding of dsDNA with PDI-2. These results suggested that dsDNA competes favourably with QDs for PDI-2 presumably due to the combination of electrostatic and π–π interactions. The changes in the ratio of PL_455nm/FL_545nm of QD-455 : PDI-2 (20 nM : 80 nM) complexes were further investigated with mixed base monomers (G, C, A, T), ssDNA sequences (S1, S3, S5, and S7) and dsDNA sequences (S1 : S2 (81% GC), S3 : S4 (50% GC), S5 : S6 (11% GC), and S7 : S8 (0% GC)) with different percentages of GC pairs. As shown in Fig. 3b, the presence of ssDNA sequences (S1, S3, S5 and S7) did not cause a significant change to the initial ratio (∼6.0) of PL_455nm/FL_545nm. In contrast, the ratio of PL_455nm/FL_545nm increased to 12.1, 9.7, 7.5, and 6.7 in the presence 100 nM of dsDNA S1 : S2, S3 : S4, S5 : S6, and S7 : S8, respectively. These results revealed that the binding ability of dsDNA to PDI-2 increases as the GC content increases. We may conclude that PDI-2 more favourably binds with GC base pairs through π–π interactions in addition to electrostatic interactions. The titration curve and detection limit of dsDNA using QD-455 : PDI-2 (20 nM : 80 nM) complexes were further investigated. As shown in Fig. 3c, a considerable change in the ratio of PL_455nm/FL_545nm can be observed in the presence of dsDNA S1 : S2 as low as 20 nM. A linear titration curve can be obtained in the range of 0 to 300 nM. In comparison, the study of S3 : S4 showed a similar titration curve with a lower binding effect and it confirmed that the binding ability is related to the GC content. In contrast, the ratio of PL_455nm/FL_545nm remained almost the same in the presence of ssDNA S1 with concentrations from 0 to 500 nM. The FRET emission of PDI-2 decreased as the concentration of dsDNA increased from 0 to 500 nM. The Stem–Volmer quenching constant was calculated to be 6.4 × 10⁶ M⁻¹, 4.3 × 10⁶ M⁻¹, and 4 × 10⁵ M⁻¹ for the presence of S1 : S2, S3 : S4, and S1, respectively (Fig. S13†).¹⁸ We also anticipated that a lower detection limit could be achieved by lowering the concentration of the complexes since the amount of dsDNA required to compete with QDs is dependent on the concentration of QDs in the complexes. A ratiometric assay of dsDNA S1 : S2 using QD-455 : PDI-2 (5 nM : 20 nM) complexes was performed. As shown in Fig. 3d, a considerable change in the ratio of PL_{455 nm}/FL_{545 nm} can be observed in the presence of as little as 2.0 nM of S1 : S2, which is a 10-fold decrease compared to the detection limit achieved using QD-455 : PDI-2 (20 nM : 80 nM) complexes. A linear titration curve can be obtained in the range of 0 to 50 nM.

Fig. 3 (a) Fluorescence spectra of QD-450 : PDI-2 (20 nM : 80 nM) in the absence and presence of S1 and S1 : S2; (b) the ratio of PL₄₅₅/FL₅₄₅ changes in the presence of the mixture of base monomers (0.1 mM of G, C, A, T), 100 nM of ssDNA S1, S3, S5, and S7, and 100 nM of dsDNA S1 : S2, S3 : S4, S5 : S6, and S7 : S8; (c) the titration of QD-450 : PDI-2 (20 nM : 80 nM) with S1, S1–S2, and S3–S4; (d) the titration of QD-450 : PDI-2 (5 nM : 20 nM) with S1–S2. The sequences of DNA oligonucleotides: S1 (5′-CCA CCC CAC CCC ACC CCA CCC CAC CCC-3′); complement S2 (5′-GGG GTG GGG TGG GGT GGG GTG GGG TGG-3′); S3 (5′-CAC ACA CAC ACA CAC ACA CAC ACA CAC-3′); complement S4 (5′-GTG TGT GTG TGT GTG TGT GTG TGT GTG-3′); S5 (5′-AAA AAA CAA AAA ACA AAA AAC AAA AAA-3′); and complement S6 (5′-TTT TTT GTT TTT TGT TTT TTG TTT TTT-3′); S7 (5′-AAA AAA AAA AAA AAA AAA AAA AA AAA-3′); complement S8 (5′-TTT TTT TTT TTT TTT TTT TTT TTT TTT-3′).

In conclusion, we have developed a new supramolecular approach to prevent the water-soluble perylene diimide PDI-2 from aggregation in aqueous solution by forming donor–accepter FRET complexes with negative charged quantum dots (QD-455 and QD490) through strong electrostatic interactions. More importantly, these FRET complexes are capable of ratiometric detection of double-stranded DNA (dsNDA) with a detection limit as low as 2 nM based on the competitive binding of dsDNA with PDI-2 over QDs. Interestingly, our studies also revealed that dsDNA with higher GC content binds more favourably with PDI-2, probably due to stronger π–π interactions between PDI-2 and GC base pairs in addition to electrostatic interactions. These newly designed QD-PDIs complexes could serve as sensitive fluorescence ratiometric probes for dsDNA.

Acknowledgements

We acknowledge the financial support from Bio-Med Scientific (Grant No. 1310043-01) and National Science Foundation for partial support.

References

1. (a) B. Valeur and M. N. Berberan-Santos, Molecular Fluorescence: Principles and Applications, John Wiley & Sons, 2012; (b) L. Yuan, W. Lin, K. Zheng and S. Zhu, Acc. Chem. Res., 2013, 46, 1462–1473; (c) H. Kobayashi, M. Ogawa, R. Alford, P. L. Choyke and Y. Urano, Chem. Rev., 2010, 110, 2620–2640.
2. (a) S.-C. Lo and P. L. Burn, Chem. Rev., 2007, 107, 1097–1116; (b) T. Qin, J. Ding, L. Wang, M. Baumgarten, G. Zhou and K. Mullen, J. Am. Chem. Soc., 2009, 131, 14329–14336.
3. (a) A. L. Briseno, S. C. B. Mannsfeld, C. Reese, J. M. Hancock, Y. Xiong, S. A. Jenekhe, Z. Bao and Y. Xia, Nano Lett., 2007, 7, 2847–2853; (b) X. Zhan, J. Zhang, S. Tang, Y. Lin, M. Zhao, J. Yang, H.-L. Zhang, Q. Peng, G. Yu and Z. Li, Chem. Commun., 2015, 51, 7156–7159.
4. (a) C. Li and H. Wonneberger, Adv. Mater., 2012, 24, 613–636; (b) D. N. Congreve, J. Lee, N. J. Thompson, E. Hontz, S. R. Yost, P. D. Reusswig, M. E. Bahlke, S. Reineke, T. Van Voorhis and M. A. Baldo, Science, 2013, 340, 334–337.
5. (a) S.-W. Tam-Chang and L. Huang, Chem. Commun., 2008, 1957–1967; (b) F. Wurthner, C. R. Saha-Moller, B. Fimmel, S. Ogi, P. Leowanawat and D. Schmidt, Chem. Rev., 2016, 116, 962–1052.
6. (a) F. Wurthner, Chem. Commun., 2004, 1564–1579; (b) F. Biedermann, E. Elmalen, I. Ghos, W. M. Nau and O. A. Sherman, Angew. Chem., Int. Ed., 2012, 51, 7739–7743.
7. (a) K. Peneva, G. Mihov, F. Nolde, S. Rocha, J.-I. Hotta, K. Braeckmans, J. Hofkens, H. Uji-I, A. Herrmann and K. Mullen, Angew. Chem., 2008, 120, 3420–3423; (b) Z. Xu, B. He, J. Shen, W. Yang and M. Yin, Chem. Commun., 2013, 49, 3646–3648; (c) T. Heek, C. Fasting, C. Rest, X. Zhang, F. Wurthner and R. Haag, Chem. Commun., 2010, 46, 1884–1886; (d) B. Gao, H. Li, H. Liu, L. Zhang, Q. Bai and X. Ba, Chem. Commun., 2011, 47(3), 894–3896.
8. (a) M. Zhu, G. H. Aryal, N. Zhang, H. Zhang, X. Su, R. Schmehl, X. Liu, J. Hu, J. Wei and J. Jayawickramarajah, Langmuir, 2015, 31, 578–586; (b) S. T. J. Ryan, J. D. Barrio, I. Ghos, F. Biedermann, A. I. Lazar, Y. Lan, R. J. Coulston, W. M. Nau and O. A. Sherman, J. Am. Chem. Soc., 2014, 136, 9053–9060; (c) F. Biedermann, E. Elmalen, I. Ghos, W. M. Nau and O. A. Sherman, Angew. Chem., Int. Ed., 2012, 51, 7739–7743; (d) R. N. Dsuza, U. Pischel and W. M. Nau, Chem. Rev., 2011, 111, 7941–7980.
9. (a) X. Feng, Y. An, Z. Yao, C. Li and G. Shi, ACS Appl. Mater. Interfaces, 2012, 4, 614–618; (b) C. Ren, J. Zhang, M. Chen and Z. Yang, Chem. Soc. Rev., 2014, 43, 7257–7266; (c) R. Huang, X. Wang, D. Wang, F. Liu, B. Mei, A. Tang, J. Jiang and G. Liang, Anal. Chem., 2013, 85, 6203–6207; (d) Y. Yuan, J. Zhang, Q. Cao, L. An and G. Liang, Anal. Chem., 2015, 87, 6180–6185.
10. J. Valanciunaite, A. S. Klymchenko, A. Skripka, L. Richert, S. Steponkiene, G. Streckyte, Y. Mely and R. Rotomskis, RSC Adv., 2014, 4, 52270–52278.
11. D. Kowerko, J. Schuster, N. Amecke, M. Abdel-Mottaleb, R. Dobrawa, F. Wurthner and C. von Borczyskowski, Phys. Chem. Chem. Phys., 2010, 12, 4112–4123.
12. (a) S. Krauß, M. Lysetska and F. Würthner, Lett. Org. Chem., 2005, 2, 349–353; (b) J. T. Kern and S. M. Kerwin, Bioorg. Med. Chem. Lett., 2002, 12, 3395–3398; (c) L. Rossetti, M. Franceschin, S. Schirripa, A. Bianco, G. Ortaggi and M. Savino, Bioorg. Med. Chem. Lett., 2005, 15, 413–420; (d) R. Samudrala, X. Zhang, R. M. Wadkins and D. L. Mattern, Bioorg. Med. Chem., 2007, 15, 186–193; (e) D. Baumstark and H. A. Wagenknecht, Angew. Chem., Int. Ed., 2008, 47, 2612–2614.
13. (a) S. Ohkuma and B. Poole, Proc. Natl. Acad. Sci. U. S. A., 1978, 5, 3327–3331; (b) J. A. Thomas, R. N. Buchsbaum, A. Zimniak and E. Racker, Biochemistry, 1979, 18, 2210–2218; (c) D. Srikun, E. W. Miller, D. W. Domaille and C. J. Chang, J. Am. Chem. Soc., 2008, 130, 4596–4597; (d) E. M. Nolan and S. J. Lippard, J. Mater. Chem., 2005, 15, 2778–2783; (e) E. M. Nolan and S. J. Lippard, J. Am. Chem. Soc., 2007, 129, 5910–5918; (f) Y. Zhang, X. Guo, W. Si, L. Jia and X. Qian, Org. Lett., 2008, 10, 473–476; (g) V. Montana, D. L. Farkas and L. M. Loew, Biochemistry, 1989, 28, 4536–4539; (h) L. Huang and S.-W. Tam-Chang, Chem. Commun., 2011, 47, 2291–2293.
14. F. Wurthner, Chem. Commun., 2004, 1564–1579.
15. C. P. N. Dsouza, U. Pischel and W. M. Nau, Chem. Rev., 2011, 111, 7941–7980.
16. R. R. Alexander and J. M. Griffiths, Basic Biochemical Methods, J.M. Wiley and Sons, 2nd edn, 1993.
17. (a) http://www.oceannanotech.com; (b) L. Yang, H. Mao, Y. A. Wang, Z. Cao, X. Peng, X. Wang, H. Duan, C. Ni, Q. Yuan, G. Adams, M. Q. Smith, W. C. Wood, X. Gao and S. Nie, Small, 2009, 5, 235–243.
18. J. R. Lakowicz, Principles of Fluorescence Spectroscopy, Springer, 3rd edn, 2006.
19. (a) A. H. Loo, Z. Sofer, D. Bousa, P. Ulbrich, A. Bonanni and M. Pumera, ACS Appl. Mater. Interfaces, 2016, 8, 1951–1957; (b) D. Siegberg and D. P. Herten, Aust. J. Chem., 2011, 64, 512–516.

---

Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c6ra16019k

---