Development of a novel H₂S and GSH detection cocktail for fluorescence imaging

Abstract

Hydrogen sulfide (H₂S) and reduced glutathione (GSH), which basically represent the two terminals of the reactivity order of small-molecular biothiols, are also the most intriguing signal molecules from this family, and their simultaneous detection is challenging but of great importance to clarify their physiopathology. Herein, we reported a borondipyrromethene (BODIPY) derivative (probe ZS2) equipped with a o-formylchalcone moiety for the simultaneous detection of H₂S and GSH in bio-samples. Due to favorable steric and electronic effects, ZS2 can selectively react with H₂S and GSH, accompanied by dramatic fluorescence intensity enhancement. When ZS2 is used in combination with our previously reported H₂S-specific probe ZS1, the simultaneous and discriminative detection of the highest reactive H₂S and the lowest reactive GSH can be realized. As a proof of concept, this detection cocktail has been applied for the quantification of endogenous H₂S and GSH in fresh rat plasma, and for their imaging in live cells. Moreover, ZS2 also features a bioorthogonal module which is readily transformed to organelle-specific functionalities. Facilitated by this module, a mitochondria-targeting version, ZS3, was straightforwardly prepared which realized the simultaneous visualization of H₂S and GSH localized in the mitochondria.

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Introduction

Free mercapto groups are widely present in living organisms in the forms of small molecular hydrogen sulfide (H₂S), cysteine (Cys), homocysteine (Hcy), glutathione (GSH), or even macro-sized Cys-bearing proteins. They generally show remarkable reductive activity which is vital for the maintenance of favorable cytosolic reduction states, and significant nucleophilic properties which is crucial for the proper functioning of some enzymes. Among the several low-molecular-weight thiols, H₂S and GSH have aroused the most research attention due to their significant yet mysterious roles in biology. The once notorious H₂S is now experiencing a revival as a gasotransmitter signalling neuromodulation in the brain and smooth muscle relaxation in the vascular system,¹ whereas the ubiquitous glutathione plays versatile roles in detoxification, redox regulation, as well as cellular signaling.² Endogenous concentrations of H₂S and GSH are finely modulated by a series of biosynthetic and metabolic processes while abnormal levels are closely related to the progress of various diseases. For example, H₂S deficiency contributes to the pathogenesis of hypertension and diabetes mellitus,³ whereas GSH imbalance is involved in the progress of aging, cancers, stroke, Alzheimer's, Parkinson's disease, vitamin deficiencies and auto-immune disorders.⁴ Given the sophisticated physiological and pathological roles H₂S and GSH play, it is important to have a robust assay available to track and qualify their existence in live organisms, which is not only important to facilitate their biological function study, but may be helpful to the prediction of H₂S and GSH related diseases.

Traditional H₂S or GSH assays, including the maleimide assay⁵ and the enzymatic colorimetric assay,⁶ are often carried out in conjugation with separation. They can therefore no longer satisfy current research needs for in situ detection. Fluorescence imaging employing chemical sensors, on the other hand, have gained a wide popularity due to their high specificity and the compatibility with live systems.⁷ More importantly, by virtue of being “switchable”, fluorescent sensors can result in extremely high target-to-background ratios.⁸ This, together with the biological significance of various biothiols, explains considerable contemporary efforts devoted to the development of biothiol fluorescent sensors.⁹ Indeed great research progress has been made in this field as typically represented by the development of probes that can selectively respond to a certain biothiol species,¹⁰ or even can discriminate different biothiols.¹¹ Noteworthy, most of these probes are devised based on the nucleophilic property of the mercapto group. Since the nucleophilic reactivity of various small-molecular biothiols generally follows the order of H₂S > Cys > Hcy ≫ GSH,¹² it therefore remains a great challenge to design GSH specific probes and more formidable to develop a probe that can simultaneously distinguish the most reactive H₂S and the most inert GSH from other biothiol species in the complex live biological context.

Apart from the specificity issue, there constitutes another challenge to design organelle-targeting probes which are important tools to study the sub-cellular localization and function of biomolecules. Current organelle-specific probes are generally developed by fusing organelle-anchoring groups, e.g. triphenylphosphonium (TPP) salts targeting mitochondria or lipophilic amines targeting lysosome,^13,14 to well-defined probes. Though technically sufficient, one limitation of this method is the laborious chemistry. Since the anchoring groups sometimes have to be incorporated into probe skeleton at the beginning of synthesis, new anchoring demands can only be satisfied by the synthesis of new probes from the beginning. To circumvent this issue, a bio-orthogonal module may be incorporated into the probe to facilitate its diversification with a variety of organelle-targeting function groups.

Here in this manuscript, we report our recent discovery of the first fluorescent probe that can selectively respond to H₂S and GSH. This probe, termed ZS2, when used in combination with our previously reported H₂S-specific probe ZS1,¹⁵ composes a detection cocktail that realizes the simultaneous sensing of H₂S and GSH. In addition, ZS2 also features a versatile handle for the fast building of organelle-targeting derivatives. As a proof of concept, its mitochondria-targeting version, ZS3 was synthesized and evaluated.

Results and discussion

Probe design and synthesis

In our previous work, a H₂S-selective fluorescent probe ZS1 was developed based on the aldehyde-acrylate sulfide-trapping chemistry developed in He's lab.¹⁶ Though being highly specific for H₂S, ZS1 suffers from the major limitation of its slow detecting kinetics. To improve the detection rate of ZS1, the acrylate group was switched to a more electron-deficient α,β-unsaturated phenyl ketone. Moreover, to facilitate further equipment of organelle-targeting groups, in the phenyl ring was preset an alkynyl group which may be readily transformed to any organelle-anchoring moiety through the facile azide–alkyne 1,3-dipolar cycloaddition (click chemistry).¹⁷ With all these considerations, ZS2 was designed as illustrated in Fig. 1. We envisioned that the electron-rich borondipyrromethene (BODIPY) scaffold and the electron-poor chalcone scaffold might form an electron donor–acceptor complex and hence quench the emission of the BODIPY fluorophore via efficient internal charge transfer (ICT).¹⁸ While the electron-density augment of the phenyl ring triggered by nucleophilic addition would block the charge transfer process and therefore restore the strong fluorogenic property of the probe. Synthesis of ZS2 and its derivatization to ZS3 was detailed in the ESI.†

Fig. 1 Structures of ZS1–3 and the design philosophy.

Selectivity of ZS2 among various small molecular biothiol species

To check if the more electron-deficient o-formylchalcone functions to accelerate the sensing kinetics of ZS2, its time-dependent fluorescent response towards H₂S was evaluated in phosphate buffered saline (PBS). As hypothesized, ZS2 alone (5 μM) was almost non-emissive in PBS (10 mM, pH 7.4, 20% acetonitrile). While treating ZS2 with sodium bisulfide (NaHS) as a sulfide source (400 μM) initiated a robust increase in fluorescent intensity and a dramatic emission band centred at 562 nm emerged (Fig. 2). Inspiringly, the detection reaction went fast enough to complete in about 20 minutes, which is a great improvement as compared to our previous acrylate-bearing ZS1 who needed almost 50 minutes to arrive at its maximum response.¹⁵ This result justifies our design philosophy to improve probe sensitivity simply by taking advantage of electronic effects.

Fig. 2 ZS2 (5 μM) responded to H₂S (400 μM) with fluorescent intensity increasing in a time-dependent way (λ_ex 530 nm).

Having gained confidence in the sensing kinetics of ZS2, we were next interested in if the o-formylchalcone moiety could also guarantee the specificity of ZS2 towards H₂S as the acrylate group did in ZS1. For this purpose, the responses of both ZS1 and ZS2 to various biothiols including H₂S, Cys, Hcy and GSH at concentrations corresponding to or higher than their respective physiological levels were tested and compared.¹⁹ As shown in Fig. 3, ZS2 responded to H₂S with a much stronger signal than ZS1 under otherwise the same conditions, in agreement with its greatly improved sensitivity. While strikingly, unlike ZS1 who exclusively preferred H₂S, ZS2 also responded to GSH with a significant fluorescent enhancement. This result, though disfavored the specificity of ZS2, was on the other hand arousing our great curiosity. As we all know, greater reactivity can often compromise substrate selectivity. From this point of view, it is not surprising that ZS2 can respond to other thiol species besides H₂S. However, given the nucleophilicity order of H₂S > Cys > Hcy ≫ GSH, it is surprising that ZS2 responds to the most reactive H₂S and the least reactive GSH while leaving the moderate reactive Cys and Hcy intact. Apart from the scientific point of view, this result is also of practical significance because, as major biological anti-oxidants, GSH and H₂S play important roles in an assortment of physiological and pathological processes, while their simultaneous detection assay is still unavailable. Since ZS1 is specific for H₂S while ZS2 for both H₂S and GSH, a detection cocktail may be developed. To this end, the detection mechanism of ZS2 was studied as the first step.

Fig. 3 Responses of ZS1 and ZS2 to various biothiols. Data shown were the fluorescence spectra (a) and intensity at 562 nm (b) of the probes (5 μM) in PBS (10 mM, pH 7.4, 20% acetonitrile) (λ_ex 530 nm) after incubating with biothiols (200 μM for Cys, Hcy, H₂S and 1.0 mM for GSH) for 1 hour.

Sensing mechanism of ZS2 towards H₂S and GSH

To shed light on the trick of ZS2 that makes it responsible to both H₂S and GSH, HPLC-HRMS analysis was carried out to reveal the possible product structures. When the detection mixtures prepared as shown in the caption of Fig. 3 were analyzed, a main signal of m/z 577.1550 was observed in the ZS2–H₂S system (Fig. S1†) and m/z 828.2679 in the ZS2–GSH system (Fig. S2†), suggesting the production of 1 : 1 adduct between ZS2 and H₂S or GSH. While when the ZS2–Cys system was analyzed, no signal other than that of ZS2 alone (m/z 543.1671) could be observed (Fig. S3†). These results were in agreement with the fact that ZS2 was fluorescently responsive to H₂S and GSH while silent to Cys (Fig. 3). Since the aldehyde-acrylate sulfide-trapping chemistry has been well established (Fig. 1), it was straightforward to speculate that ZS2 responded to H₂S by the tandem nucleophilic addition and thereafter the formation of the electron-rich 1,3-dihydrobenzo[c]thiophene moiety. While for the sterically bulky GSH, it remained unclear how the 1 : 1 adduct was formed. To get the detailed mechanism, ¹H NMR spectra of the thiol-recognition structure, compound 7, were investigated in the absence and presence of H₂S or GSH (Fig. S4†). As expected, treating 7 (9.4 μM, DMSO-d₆/D₂O (3 : 1)) with NaHS (1 equiv.) resulted in the disappearance of the signals at 10.11 (s, 1H), 8.51 (d, J = 15.6 Hz, 1H), 7.71 (d, J = 15.6 Hz, 1H) assigned to the aldehyde and the trans-olefin, indicating the total transformation of 7 to the dihydrobenzo[c]thiophene product 7a. While the addition of GSH led to the proportional decrease of these two pairs of signals, implying that a tandem nucleophilic addition was taking place and a macrocyclic product 7b formed (Fig. 4). These results, on the one hand unveiled the detection mechanism of ZS2 towards H₂S and GSH shown below, also offered a possible explanation of its special selectivity mode (Fig. S5†). We speculated that the thiol group of GSH first added to the aldehyde of ZS2 resulting in a hemithioacetal intermediate, which brought the amine group of GSH in close proximity to the reactive α,β-unsaturated carbonyl of ZS2, leading to the Michael addition between the amine and the active olefin. While for Cys or Hcy, the unfavorable steric hindrance prevented the second nucleophilic addition from taking place, and the system remained dark in equilibrium of ZS2 and the hemithioacetal (Fig. S5†). ZS1 may also react with GSH to form the hemithioacetal, yet its acrylate is not reactive enough for the amino nucleophilic attack (Fig. S5†). It is therefore postulated that both favorable spatial distance between the amine and the Michael receptor, and favorable reactivity of the latter, are necessary for the probe to respond to GSH.

Fig. 4 Tandem nucleophilic addition between the o-formylchalcone moiety and H₂S or GSH as indicated by HRMS and NMR analysis.

Detection kinetics of ZS2 towards H₂S and GSH

To establish a feasible protocol for the simultaneous detection of H₂S and GSH employing ZS2, we next set out to study its reaction kinetics in detail. For this purpose, the fluorescent responses of ZS2 towards various concentrations of H₂S or GSH were recorded as time lapsed. As shown in Fig. S6–S9,† the emission intensity at 562 nm of ZS2 (5 μM in PBS) treated with varying concentrations of NaHS (50–400 μM) increased gradually and reached the maxima in about 20 minutes. Under pseudo-first-order conditions (5 μM ZS2 and 400 μM NaHS), the observed rate constant for NaHS detection is k_obs = 7.13 × 10⁻⁴ S⁻¹ (Fig. 5a). While the plot of k_obs versus [NaHS] indicated that the process was overall 2nd order with k₂ = 1.61 M⁻¹ S⁻¹ (Fig. 5b). Similar results were obtained for GSH detection process (Fig. S10–S15†), except that it proceeded in a relatively sluggish way with k_obs = 2.69 × 10⁻⁴ S⁻¹ and k₂ = 0.29 M⁻¹ S⁻¹ (Fig. 5c and d), which are two and five times slower than those of the ZS2–H₂S reaction. These results are in agreement with the afore-proposed sequential-two-step-nucleophilic–addition mechanism that the rate-limiting thiol-aldehyde addition goes first followed by the fast intramolecular cyclization.

Fig. 5 Reaction kinetics profiling of ZS2 towards NaHS (a and b) and GSH (c and d). ZS2 (5 μM) was treated with different concentrations of NaHS (100, 200, 400 μM) or GSH (100, 200, 400, 800, 1000 μM) and the fluorescent spectra were recorded at varying time intervals (λ_ex 530 nm). Kinetic plot of fluorescence emission intensity at 562 nm gave k_obs and k₂ as shown.

Concentration-dependent responses of ZS2 towards H₂S and GSH

To evaluate the sensitivity of ZS2, its fluorescent spectra after incubating with various concentrations of NaHS or GSH were recorded (Fig. S16 and S17†). Much to our delight, ZS2 responded to different concentrations of NaHS or GSH with fluorescent intensity increasing in a linearly correlated way (Fig. 6), which is a highly desirable quality since linear correlation usually facilitates quantification. For H₂S detection, the linear range is 0–350 μM while 0–1400 μM for GSH analysis, both of which fall into the ranges of reported native concentrations of biological H₂S/GSH,¹⁹ indicating the great potency of ZS2 to quantify endogenous H₂S/GSH. Moreover, the in vitro detection limits were calculated to be 0.80 μM for H₂S and 8.0 μM for GSH (S/N = 3), which are sufficiently sensitive to qualify endogenous H₂S or GSH.

Fig. 6 Probe ZS2 responded to H₂S or GSH in a concentration-dependent manner. Plot of the emission intense at 562 nm (λ_ex 530 nm) of ZS2 (5 μM) in PBS (10 mM, pH 7.4, 20% acetonitrile) vs. the concentrations of H₂S (a) or GSH (b) gave linear correlation.

Selectivity of ZS2 among various bio-relevant species

We also assessed the selectivity of ZS2 across a panel of biologically relevant analytes including various amino acids, cations and anions (PBS, 10 mM, pH 7.4). As shown in Fig. 7, none species other than H₂S and GSH could trigger obvious intensity enhancement. Moreover, ZS2 could also respond to H₂S or GSH in the presence of other analytes with fluorescence readouts similar to those without the presence of other species, indicating that ZS2 is exclusively selective for H₂S and GSH and may be applicable in complex biological samples.

Fig. 7 Fluorescence intensity of ZS2 (5 μM) at 562 nm in the presence of various bio-relevant analytes: blank (1), H₂O₂ (2), ClO⁻ (3), Mg²⁺ (4), Fe²⁺ (5), Fe³⁺ (6), Zn²⁺ (7), Cu²⁺ (8), Cys (9), Ala (10), Gly (11), Hcy (12), EtSH (13), SO₃²⁻ (14), NaHS (15), GSH (16). Concentrations: 1.0 mM for GSH and 200 μM for others. Data were obtained after one hour of incubation (λ_ex 530 nm).

Fluorescence responses of ZS2 to co-existing GSH and H₂S

Based on the selectivity experiments, we are confident about the preference of ZS2 for H₂S and GSH. However, it remained unknown how this probe would respond to co-existing GSH and H₂S. To address this question, we recorded the fluorescence spectra of ZS2 after reacting with various concentrations of GSH in the co-presence of H₂S. When the intensities at emission maxima (562 nm) were plotted, it was observed that the final signal readouts were the sum of those obtained when H₂S or GSH was present alone, with the relative error less than 5% (Fig. S18†). This result demonstrates that H₂S doesn't interfere the accurate quantification of GSH by probe ZS2 in PBS, and vice versa. Also, this result consolidates the feasibility of cocktail detection of biological H₂S and GSH employing ZS2 in combination with the H₂S-specific ZS1.

Quantifying endogenous sulfides in fresh rat plasma

With the possibility of ZS2 in combination with ZS1 to simultaneously detect H₂S and GSH well characterized in PBS buffer, we next tested the feasibility of this cocktail to quantify endogenous H₂S and GSH in fresh male Sprague-Dawley (SD) rat plasma. For this purpose, fresh blood from the same rat was divided into two parts, with one aliquot for the quantification of H₂S and the other for the quantification of GSH after precipitation of endogenous H₂S by ZnCl₂. The spike method was employed for the measurement of endogenous H₂S or GSH, with details fully described in the ESI (Fig. S19–S21†). It turned out the rats tested had a mean endogenous concentration of 76.7 ± 13.3 μM for H₂S and 853.3 ± 80.4 μM for GSH, close to other reports on H₂S and GSH concentrations in rat blood plasma.²⁰

Live cell imaging of H₂S and GSH employing ZS2

Based on the in vitro characterization studies of ZS2, the feasibility of ZS2 to sense H₂S and GSH was well established. We therefore moved forward to the simultaneous imaging of H₂S and GSH in live cells. For this purpose, we first chose MEF cells endogenously expressing both H₂S and GSH as the model cells. As shown in Fig. 8, no detectable fluorescence signal was observed for the cells in the absence of ZS2, indicating that no undesired autofluorescence would interfere the results. While significant intracellular fluorescence was detected in the cells loaded with ZS2 (5 μM), evidencing the good cell permeability of ZS2 and the presence of abundant intracellular GSH/H₂S. Interestingly, when the cells preloaded with ZS2 were treated with exogenous GSH or H₂S, the intensity of the intracellular fluorescence was positively correlated with the concentrations of GSH or H₂S in the culture medium (Fig. 8). Even more, the increase of intracellular fluorescent intensity of cells treated with both GSH and H₂S was almost equivalent to the total increase when the cells were incubated with GSH or H₂S alone (Fig. 8d), suggesting the great potency of ZS2 to quantify intracellular GSH/H₂S. Similar results were obtained in other cell lines such as EA.hy926 (Fig. S22†). Moreover, the ability of probe ZS3 to target mitochondria was also tested by staining cells with both ZS3 and MitoTracker Green. As anticipated, ZS3 fluorescence showed strong colocalization with MitoTracker Green (Fig. S23†), suggesting the potency of ZS3 to track GSH and H₂S located in the mitochondrial components of cells, and the feasibility of the fast-transformation of fluorescent probes to organelle-targeting ones by virtue of a preset alkynyl group and last-step “click chemistry”.

Fig. 8 Simultaneous detection of GSH and H₂S in live MEF cells by ZS2. Data shown were the representative images of MEF cells without or with the preloading of ZS2 (5 μM), followed by the incubation with various concentrations of exogenous GSH (a), H₂S (b), or both GSH and H₂S (c). (d) Quantified fluorescence intensities of cells as represented in (a)–(c).

Conclusions

To conclude, by switching the acrylate in our previously reported H₂S-specific probe ZS1 to the more electron-poor 3-oxo-3-phenylprop-1-enyl, we have designed ZS2 which, due to the more electron-deficient Michael receptor, can not only sense H₂S with dramatically accelerated kinetics, but is reactive enough for the nucleophilic addition with the free amino in GSH in tandem with the aldehyde-thiol semi-mercaptalation. ZS2 represents the first fluorescent probe that realizes the simultaneous detection of the highest reactive H₂S and the lowest reactive GSH among various biothiols. When used in combination with ZS1, a detection cocktail is developed for the simultaneous and discriminative detection of H₂S and GSH. Feasibility of this cocktail for quantifying endogenous H₂S and GSH in fresh rat plasma and for imaging live cells has been well characterized. Moreover, ZS2 also features a versatile handle that is readily transformed to organelle-targeting groups, and its mitochondria version was prepared and tested as a proof of concept. Given the important yet mysterious physiopathology of H₂S and GSH, we think that ZS2 is a helpful tool for biology study and holds promise to illuminate the intracellular crosstalk between H₂S and GSH.

Acknowledgements

This research is supported by the National Science Fund for Distinguished Young Scholars (81125023), the National Natural Science Foundation of China (91213303), the Program of Shanghai Subject Chief Scientist (13XD1404300) and Zhejiang Provincial Natural Science Foundation of China (LY15H300003).

Notes and references

1. (a) B. D. Paul and S. H. Snyder, Nat. Rev. Mol. Cell Biol., 2012, 13, 499; (b) L. Li, P. Rose and P. K. Moore, Annu. Rev. Pharmacol. Toxicol., 2011, 51, 169; (c) H. Kimura, N. Shibuya and Y. Kimura, Antioxid. Redox Signaling, 2012, 17, 45; (d) W. L. Chen, Y. Y. Niu, W. Z. Jiang, H. L. Tang, C. Zhang, Q. M. Xia and X. Q. Tang, Rev. Neurosci., 2015, 26, 129; (e) X. Zhang and J. S. Bian, ACS Chem. Neurosci., 2014, 5, 876; (f) K. M. Holwerda, S. A. Karumanchi and A. T. Lely, Curr. Opin. Nephrol. Hypertens., 2015, 24, 170; (g) N. Skovgaard, A. Gouliaev, M. Aalling and U. Simonsen, Curr. Pharm. Biotechnol., 2011, 12, 1385; (h) Y. H. Liu, M. Lu, L. F. Hu, P. T. Wong, G. D. Webb and J. S. Bian, Antioxid. Redox Signaling, 2012, 17, 141; (i) K. Kashfi and K. R. Olson, Biochem. Pharmacol., 2013, 85, 689.
2. (a) K. Aquilano, S. Baldelli and M. R. Ciriolo, Front. Pharmacol., 2014, 5, 196; (b) P. G. Board and D. Menon, Biochim. Biophys. Acta, 2013, 1830, 3267; (c) M. Marí, A. Morales, A. Colell, C. García-Ruiz, N. Kaplowitz and J. C. Fernández-Checa, Biochim. Biophys. Acta, 2013, 1830, 3317; (d) M. L. Circu and T. Y. Aw, Biochim. Biophys. Acta, 2012, 1823, 1767; (e) R. Franco and J. A. Cidlowski, Antioxid. Redox Signaling, 2012, 17, 1694; (f) S. Singh, A. R. Khan and A. K. Gupta, J. Exp. Ther. Oncol., 2012, 9, 303.
3. (a) Y. Kaneko, Y. Kimura, H. Kimura and I. Niki, Diabetes, 2006, 55, 1391; (b) G. Yang, L. Wu, B. Jiang, W. Yang, J. Qi, K. Cao, Q. Meng, A. K. Mustafa, W. Mu, S. Zhang, S. H. Snyder and R. Wang, Science, 2008, 322, 587; (c) S. Mani, A. Untereiner, L. Wu and R. Wang, Antioxid. Redox Signaling, 2014, 20, 805.
4. (a) S. C. Lu, Mol. Aspects Med., 2009, 30, 42; (b) D. M. Townsend, K. D. Tew and H. Tapiero, Biomed. Pharmacother., 2003, 57, 145.
5. (a) R. J. Ellis, Nature, 1966, 211, 1266; (b) R. A. Winters, J. Zukowski, N. Ercal, R. H. Matthews and D. R. Spitz, Anal. Biochem., 1995, 227, 14.
6. (a) O. W. Griffith, Anal. Biochem., 1980, 106, 207; (b) U. Hannestad, S. Margheri and B. Sörbo, Anal. Biochem., 1989, 178, 394.
7. (a) K. M. Dean and A. E. Palmer, Nat. Chem. Biol., 2014, 10, 512; (b) A. Nadler and C. Schultz, Angew. Chem., Int. Ed., 2013, 52, 2408; (c) E. A. Lemke and C. Schultz, Nat. Chem. Biol., 2011, 7, 480; (d) H. Kobayashi, M. Ogawa, R. Alford, P. L. Choyke and Y. Urano, Chem. Rev., 2010, 110, 2620; (e) L. D. Lavis and R. T. Raines, ACS Chem. Biol., 2008, 3, 142; (f) D. L. Taylor and Y. L. Wang, Nature, 1980, 284, 405.
8. (a) D. W. Domaille, E. L. Que and C. J. Chang, Nat. Chem. Biol., 2008, 4, 168; (b) T. Ueno and T. Nagano, Nat. Methods, 2011, 8, 642; (c) M. Schäferling, Angew. Chem., Int. Ed., 2012, 51, 3532; (d) Z. Guo, S. Park, J. Yoon and I. Shin, Chem. Soc. Rev., 2014, 43, 16; (e) G. Patterson, M. Davidson, S. Manley and J. Lippincott-Schwartz, Annu. Rev. Phys. Chem., 2010, 61, 345.
9. (a) H. S. Jung, X. Chen, J. S. Kim and J. Yoon, Chem. Soc. Rev., 2013, 42, 6019; (b) K. Wang, H. Peng and B. Wang, J. Cell. Biochem., 2014, 115, 1007; (c) H. Peng, W. Chen, Y. Cheng, L. Hakuna, R. Strongin and B. Wang, Sensors, 2012, 12, 15907; (d) X. Chen, Y. Zhou, X. Peng and J. Yoon, Chem. Soc. Rev., 2010, 39, 2120.
10. (a) V. S. Lin, W. Chen, M. Xian and C. J. Chang, Chem. Soc. Rev., 2015, 44, 4596; (b) F. Yu, X. Han and L. Chen, Chem. Commun., 2014, 50, 12234; (c) J. Yin, Y. Kwon, D. Kim, D. Lee, G. Kim, Y. Hu, J. H. Ryu and J. Yoon, J. Am. Chem. Soc., 2014, 136, 5351; (d) L. Y. Niu, Y. S. Guan, Y. Z. Chen, L. Z. Wu, C. H. Tung and Q. Z. Yang, J. Am. Chem. Soc., 2012, 134, 18928; (e) X. Yang, Y. Guo and R. M. Strongin, Angew. Chem., Int. Ed., 2011, 50, 10690; (f) W. Wang, J. O. Escobedo, C. M. Lawrence and R. M. Strongin, J. Am. Chem. Soc., 2004, 126, 3400.
11. (a) Q. Miao, Q. Li, Q. Yuan, L. Li, Z. Hai, S. Liu and G. Liang, Anal. Chem., 2015, 87, 3460; (b) M. Y. Jia, L. Y. Niu, Y. Zhang, Q. Z. Yang, C. H. Tung, Y. F. Guan and L. Feng, ACS Appl. Mater. Interfaces, 2015, 7, 5907; (c) F. Wang, Z. Guo, X. Li, X. Li and C. Zhao, Chem.–Eur. J., 2014, 20, 11471; (d) J. Liu, Y. Q. Sun, H. Zhang, Y. Huo, Y. Shi and W. Guo, Chem. Sci., 2014, 5, 3183; (e) J. Liu, Y. Q. Sun, Y. Huo, H. Zhang, L. Wang, P. Zhang, D. Song, Y. Shi and W. Guo, J. Am. Chem. Soc., 2014, 136, 574; (f) X. F. Yang, Q. Huang, Y. Zhong, Z. Li, H. Li, M. Lowry, J. O. Escobedo and R. M. Strongin, Chem. Sci., 2014, 5, 2177.
12. W. X. Ren, J. Han, T. Pradhan, J. Y. Lim, J. H. Lee, J. Lee, J. H. Kim and J. S. Kim, Biomaterials, 2014, 35, 4157.
13. (a) A. T. Hoye, J. E. Davoren, P. Wipf, M. P. Fink and V. E. Kagan, Acc. Chem. Res., 2008, 41, 87; (b) A. Martin, A. Byrne, C. S. Burke, R. J. Forster and T. E. Keyes, J. Am. Chem. Soc., 2014, 136, 15300.
14. S. D. Wiedner, L. N. Anderson, N. C. Sadler, W. B. Chrisler, V. K. Kodali, R. D. Smith and A. T. Wright, Angew. Chem., Int. Ed., 2014, 53, 2919.
15. X. Li, S. Zhang, J. Cao, N. Xie, T. Liu, B. Yang, Q. He and Y. Hu, Chem. Commun., 2013, 49, 8656.
16. (a) Y. Qian, J. Karpus, O. Kabil, S. Y. Zhang, H. L. Zhu, R. Banerjee, J. Zhao and C. He, Nat. Commun., 2011, 2, 495; (b) Y. Qian, L. Zhang, S. Ding, X. Deng, C. He, X. E. Zheng, H. L. Zhu and J. Zhao, Chem. Sci., 2012, 3, 2920.
17. (a) H. C. Kolb, M. G. Finn and K. B. Sharpless, Angew. Chem., Int. Ed., 2001, 40, 2004; (b) V. V. Rostovtsev, L. G. Green, V. V. Fokin and K. B. Sharpless, Angew. Chem., Int. Ed., 2002, 41, 2596.
18. J. R. Lakowicz, Springer Science Business Media, 3rd edn, 2006, ch. 6, pp. 205–235.
19. (a) N. Tateishi, T. Higashi, A. Naruse, K. Nakashima and H. Shiozaki, J. Nutr., 1977, 107, 51; (b) A. Meister and M. E. Anderson, Annu. Rev. Biochem., 1983, 52, 711; (c) J. Furne, A. Saeed and M. D. Levitt, Am. J. Physiol.: Regul., Integr. Comp. Physiol., 2008, 295, 1479; (d) P. M. Ueland, H. Refsum, S. P. Stabler, M. R. Malinow, A. Andersson and R. H. Allen, Clin. Chem., 1993, 39, 1764.
20. (a) M. Whiteman and P. K. Moore, J. Cell. Mol. Med., 2009, 13, 488; (b) T. K. Chung, M. A. Funk and D. H. Baker, J. Nutr., 1990, 120, 158.

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Footnotes

† Electronic supplementary information (ESI) available: Synthesis and characterization of ZS2, ZS3; fluorometric analysis methods; cell imaging methods; ESI figures; NMR spectra of ZS2, ZS3 and intermediates. See DOI: 10.1039/c6ra08998d

‡ These authors contributed equally to this work.

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