DOI: 10.1002/cbic.201500249
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Rational Design of a Ratiometric Fluorescent Probe Based on Arene–Metal-Ion Contact for Endogenous Hydrogen Sulfide Detection in Living Cells Ryosuke Kawagoe, Ippei Takashima, Kazuteru Usui, Anna Kanegae, Yusuke Ozawa, and Akio Ojida*[a] We report the design and development of a fluorescent CdII ion complex that is capable of the ratiometric detection of H2S in living cells. This probe exploits the metal-ion-induced emission red shift resulting from direct contact between the aromatic ring of a fluorophore and a metal ion (i.e., arene–metalion or “AM” contact). The CdII complex displays a large emission blue shift upon interaction with H2S as the CdII-free ligand is released by the formation of cadmium sulfide. Screening of potential ligands and fluorophores led to the discovery of a py-
ronine-type probe, 6·CdII, that generated a sensitive and rapid ratio value change upon interaction with H2S, without interference from the glutathione that is abundant in the cell. The membrane-impermeable 6·CdII was successfully translocated into live cells by using an oligo-arginine peptide and pyrenebutylate as carriers. As such, 6·CdII was successfully applied to the ratiometric detection of both exogenous and endogenous H2S produced by the enzymes in living cells, thus demonstrating the utility of 6·CdII in biological fluorescence analysis.
Introduction Hydrogen sulfide (H2S) is the most recently identified gaseous signaling molecule in living organisms.[1] Although H2S has long been considered a toxic compound, there is now an accumulation of scientific evidence that this redox-active molecule plays important metabolic roles, for example in angiogenesis,[2] cardioprotection[3] and the induction of hippocampal longterm potentiation.[4] In mammals, H2S is enzymatically produced by cystathionine-b-synthase, cystathionine-g-lyase (CSE), and cysteine amino transferase in combination with 3-mercaptosulfurtransferase.[5] To better understand the biological functions of this compound, it is critical to develop a noninvasive real-time detection method for H2S in living cell systems, and thus a number of fluorescent probes for H2S have been developed in recent years.[6] These probes were designed based on three different sensing mechanisms: H2S-mediated azide reduction,[7] nucleophilic reaction of H2S,[8] and metal–sulfide complex precipitation.[9] Most such probes were designed to detect H2S levels by increases in single fluorescence emission. However, single-emission sensing is readily perturbed by various aspects of the cellular environment, such as intracellular pH, cell volume, and inhomogeneous probe localization, thus making it difficult to quantitatively detect H2S under in vivo conditions. Ratiometric fluorescence sensing, which is based on the simultaneous detection of two emission signals, can eliminate these unfavorable effects and allows quantitative [a] R. Kawagoe, I. Takashima, Dr. K. Usui, A. Kanegae, Y. Ozawa, Prof. Dr. A. Ojida Graduate School of Pharmaceutical Sciences, Kyushu University 3-1-1 Maidashi, Higashi-ku, Fukuoka 812-8582 (Japan) E-mail: [email protected] Supporting information for this article is available on the WWW under http://dx.doi.org/10.1002/cbic.201500249.
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data analysis with significant precision.[10] Several ratiometric probes for H2S have been reported in the last few years.[8c, 11] Unfortunately, despite having elaborate molecular designs, these probes still suffer from limited fluorescence, slow responses to the H2S flux, or an inability to detect endogenously produced H2S. Recently, we reported the unique fluorescence sensing of metal ions based on direct contact between the aromatic ring of a fluorophore and a metal ion (i.e., arene–metal-ion or “AM” contact).[12] Fluorescent probes with this sensing mechanism exhibited large emission red shifts upon coordination with various metal ions, including CdII, AgI, HgII, and PbII. By exploiting AM-contact sensing, we previously developed the xanthenetype CdII complex 1·CdII as a ratiometric fluorescence probe for H2S (Scheme 1).[12a] The 1·CdII system was able to sense H2S based on the significant emission blue shift (DF = 18 nm) induced by loss of the arene–metal-ion contact when the CdII ion is released from 1·CdII to form cadmium sulfide. Although 1·CdII could be applied to the ratiometric detection of exogenously supplied H2S in live cells, there was unexpectedly strong quenching of the fluorescence emission by glutathione (GSH). This quenching, the resulting weak fluorescence signal, and associated small change in signal ratio significantly diminished the usefulness of 1·CdII, and precluded its further application to the detection of endogenously produced H2S in living cells. In this paper, we report the design of a ratiometric fluorescent probe for H2S with improved sensing properties. Careful optimization of both the ligand and fluorophore structures provided a new CdII complex, 6·CdII, that functioned as a highly sensitive fluorescent probe for H2S, rapidly generating a large ratio change. Notably, the fluorescence of 6·CdII was not quenched by high concentrations of GSH (5 mm).
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Full Papers different ligand units (Scheme 2). The syntheses of these probes are described in the Supporting Information (Scheme S1). The probes’ fluorescence spectral changes upon the addition of H2S are summarized in Figure S2; Table 1 gives the fluorescence responses of the probes for H2S (30 mm) and GSH (5 mm) under aqueous MeOH conditions (50 mm HEPES, pH 7.4/MeOH 1:1). Probe 2·CdII, which has two 2,2’-dipicolylamine (Dpa) ligands, exhibited Scheme 1. Ratiometric fluorescence sensing of 1·CdII for H2S based on arene-metal ion a large emission blue shift (DF = ¢16 nm) upon the contact. addition of H2S. A significant emission shift (DF = ¢8 nm) was also observed in the case of 3·CdII, with Given these superior sensing properties, 6·CdII was successfully iminodiacetic acid (Ida) ligands, whereas the emission blue applied to the ratiometric detection of not only exogenously shift of 4·CdII, which has 2-picolylaminoacetic acid (Paa) lisupplied H2S, but also the endogenously produced H2S in gands, was small (DF = ¢2 nm). The large emission blue shifts living cells, thus highlighting its usefulness in biological fluorescence analysis.
Result and Discussion Screening of the ligand unit Figure 1 presents the fluorescence and absorption spectral changes of 1·CdII (10 mm, 20 mm HEPES, pH 7.4) upon the addition of high concentrations of GSH (0–5 mm). The significant quenching of the fluorescence concomitant with the decrease in absorbance around 500 nm suggests that the conjugated structure of the xanthene ring of 1·CdII was disrupted by the presence of GSH. Analysis of a mixture of 1·CdII and GSH by ESI-MS identified a new product with a m/z value of 1054, corresponding to the GSH adduct of 1·CdII (Figure S1 in the Supporting Information). Based on these spectral data, we concluded that the GSH-induced fluorescence quenching primarily results from nucleophilic attack of GSH on the xanthene ring of 1·CdII at the C9 carbon, which is known to be susceptible to such attack.[13]
Scheme 2. Fluorescent CdII complexes for ratiometric detection of H2S.
Table 1. Summary of the fluorescence response of the probes toward H2S and glutathione (GSH).
Figure 1. A) Absorption and B) fluorescence spectral changes of 1·CdII (10 mm) upon addition of glutathione (GSH, 0–5 mm). Measurement conditions: 20 mm HEPES, pH 7.4, lex = 500 nm.
Initially, we attempted to find suitable ligands for the probe that were capable of preventing the nucleophilic attack of GSH on the fluorophore. Since AM-contact sensing can work with various fluorophores,[12a] we prepared the readily available anthracene-type probes 2·CdII to 4·CdII, each of which possessed ChemBioChem 2015, 16, 1608 – 1615
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DF [nm][a] FGSH/Fo[b]
1·CdII
2·CdII
3·CdII
4·CdII
¢21 0.16
¢16 0.79
¢8 1.01
¢2 0.98
[a] Shift of the emission maximum wavelength upon addition of Na2S: 30 mm for 1·CdII, 100 mm for 2·CdII, 3·CdII, and 4·CdII. [b] Relative fluorescence intensity in the absence (Fo) and in presence of 5 mm GSH (FGSH), lex = 500 nm (1·CdII), lex = 360 nm (2·CdII, 3·CdII, 4·CdII).
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Full Papers seen for 2·CdII and 3·CdII in the presence of H2S were attributed to the intrinsically large emission red shifts of ligands 2 (DF = 15 nm) and 3 (DF = 10 nm) induced by CdII coordination, whereas the emission red shift of 4 upon CdII complexation is minimal (DF = 5 nm). These data suggest that the structure of the ligand unit, which assists the direct contact between the CdII and the fluorophore, can influence the ratiometric sensing property of the probe. In contrast to the xanthene-type probe 1·CdII, the anthracene-type CdII complexes showed very little decrease in fluorescence in the presence of 5 mm GSH (Table 1, Figure S2). This can be explained by the inertness of the anthracene ring toward the nucleophilic attack of GSH at the C9 position. We subsequently used isothermal titration calorimetry (ITC) to evaluate the coordination interactions of the anthracene-type probes with GSH. In the case of 2·CdII, which has Dpa ligands, endothermic binding was observed upon titration with GSH, and the binding constant (Ka) was found to be 5.48 Õ 104 m¢1 (n = 0.57) by curve-fitting analysis (Figure S3). We believe that this coordination interaction could enhance the nucleophilic attack of GSH on the xanthene ring of 1·CdII. In contrast, the ITC curves of 3·CdII and 4·CdII upon titration with GSH exhibited minimal change, thus suggesting that the binding affinities of these compounds are very weak (Ka < 103 m¢1). The weak binding affinities of 3·CdII and 4·CdII could result from the negative charges of the Ida and Paa ligands, which attenuate the electrostatic interaction between the anionic GSH and the coordinated CdII ions. Overall, these fluorescence and ITC data indicate that the Ida ligand in 3·CdII, which displays a large
emission blue shift upon binding with H2S and also reduces unwanted interactions with GSH, is the most suitable ligand for the ratiometric detection of H2S. Optimization of the fluorophore We next sought to optimize the fluorophore of the CdII complex to obtain a probe capable of reacting to H2S with a large emission blue shift, while remaining resistant to GSH-induced fluorescence quenching. The CdII complexes 5·CdII to 7·CdII were each designed with a different xanthene fluorophore (Scheme 2). Given the results of the ligand-optimization work, these probes had Ida as ligands. The synthetic route to the pyronine-type ligand 6 is given in Scheme 3. Briefly, the 4,7-dihydroxyxanthone 9 obtained from 8[12a] by deprotection of the pivaloyl groups was converted to ditriflate 10, which was treated with pyrrolidine to give the 4,7-diaminoxanthone 11.[14] After reduction with borane-dimethylsulfide complex, 12 was oxidized by 2,3-dichloro-5,6-dicyano-1,4-benzoquinone (DDQ) to afford pyronine 13. Finally, deprotection of tert-butyl esters and subsequent HPLC purification provided the pure ligand 6. The photophysical properties of the ligands and their CdII complexes are summarized in Table S1. In CdCl2 titration (Figure S4), each ligand 5 to 7 displayed a large emission red shift upon coordination with CdII ions, together with simultaneous absorption red shifts; these are indicative of contact interaction between the CdII ions and the fluorophores. Figures 2 A and S5 show the fluorescence responses of 5·CdII to 7·CdII toward H2S under aqueous conditions. All probes displayed
Scheme 3. Synthesis of ligand 6. a) 4 n NaOH, THF/MeOH (86 %); b) Tf2O, pyridine, CH2Cl2 (82 %); c) pyrrolidine, DMSO (20 %); d) BH3SMe2, THF (63 %); e) DDQ, EtOH/CH2Cl2 (78 %); f) TFA, CH2Cl2 ; g) HPLC (25 %).
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Figure 2. A) Change in fluorescence spectrum of 6·CdII (10 mm) upon addition of Na2S (0– 20 mm). Measurement conditions: 20 mm HEPES, pH 7.4, lex = 550 nm. B) Plot of the ratio value, R, of 6·CdII (10 mm) upon addition of Na2S. Measurement conditions: 20 mm HEPES, pH 7.4, lex = 550 nm. C) Change in the ratio value (DR) of the CdII complexes (10 mm) upon addition of Na2S (30 mm). 1·CdII : DR = F527 nm/F549 nm, 5·CdII : DR = F528 nm/ F546 nm, 7·CdII : DR = F593 nm/F613 nm. Measurement conditions: 20 mm HEPES, pH 7.4, lex = 500 nm (1·CdII and 5·CdII), lex = 550 nm (6·CdII and 7·CdII). D) Change in the relative fluorescence intensity (F/Fo) of 1·CdII (*), 5·CdII (&), 6·CdII (~), and 7·CdII (^) upon addition of GSH (0–5 mm). Measurement conditions: [probe] = 10 mm, 20 mm HEPES, pH 7.4.
large emission blue shifts upon the addition of sodium sulfide (Na2S) as a donor of H2S, leading to changes in the fluorescence ratio value R, as shown in Figure 2 B and C. Simultaneously, the absorption spectra of the probes were hypsochromically shifted (Figure S5); this suggests that the emission shifts were induced by the loss of AM-contact as a result of the release of CdII-free ligands. Most intriguingly, the probes showed different magnitudes of fluorescence quenching response to GSH. Figure 2 D summarizes the changes in the fluorescence intensities of the probes upon the addition of 0–5 mm GSH. The decrease in the fluorescence intensity of 5·CdII was less than that of the previously developed 1·CdII, thus suggesting that 5·CdII was less reactive toward GSH than 1·CdII was. This could possibly be ascribed to the lower binding affinity of the Ida·CdII complex with GSH compared to that of the Dpa·CdII complex, as was observed in the case of anthracene-type CdII complexes (Figure S3). Interestingly, the pyronine-type probe 7·CdII, with Nacetylpiperazine ligands, lost essentially all fluorescence upon the addition of 2 mm of GSH, whereas the pyronine-type probe 6·CdII, with pyrrolidine ligands, maintained its original fluorescence even in the presence of 5 mm of GSH. We speculate that the different reactivities of these complexes can be explained in terms of the LUMO energy levels of the fluorophores, as reported by Urano et al.[13d] Scheme 4 gives the calculated LUMO energy levels of fluorophores 5’ to 7’, all of which are analogues of the fluorophores of 5·CdII to 7·CdII. Density functional theory calculations suggest that 6’ has ChemBioChem 2015, 16, 1608 – 1615
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a higher LUMO energy level than 7’; this is indicative of the lower electrophilic reactivity of 6·CdII relative to 7·CdII,[15] and is consistent with the experimental results, which show a higher tolerance of 6·CdII for GSH. The LUMO energy level of the xanthene-based 5’ was calculated to be higher than that of 6’, which cannot explain the higher reactivity of 5·CdII with GSH compared with that of 6·CdII. However, when the hydrogen bonding of the carbonyl oxygen of 5’ with surrounding water molecules (according to Jorgensen’s model[16]) were taken into account, the LUMO energy level of 5’ was predicted to be lower than that of 6’, and this is consistent with the experimental observations of the higher resistance of 6·CdII toward GSH. Overall, the data in Figure 2 suggest that 6·CdII is the most suitable probe for H2S sensing among the three CdII complexes. The sensing properties of 6·CdII were further evaluated by fluorescence titration experiments. Figure 3 A plots the time-dependent fluorescence response of 6·CdII toward H2S in the presence and absence of GSH. In the absence of GSH, the fluorescence ratio value R (F599 nm/F619 nm) of 6·CdII gradually increased and reached a plateau (R = 2.0) 5 min after treatment with Na2S (a donor of gaseous H2S); no ratio change was induced by GSH (5 mm) even after 10 min. Interestingly, the response of 6·CdII toward H2S was accelerated by the presence of GSH; the R value rapidly (within ca. 1 min) increased to 2.0
Scheme 4. LUMO energy levels of the fluorophores from DFT calculations.
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Figure 3. A) Time trace plot of the ratio value, R, of 6·CdII (10 mm) upon addition 5 mm of GSH (~), and upon addition of 30 mm of Na2S in the presence (&) and absence (*) of 5 mm of GSH. Measurement conditions: 20 mm HEPES, pH 7.4, lex = 550 nm. The error bars represent standard deviations obtained from triplicate experiments. B) The ratio value of 6·CdII (10 mm) in the presence of various thiol species; 1) none, 2) Na2S (30 mm), 3) Na2S (30 mm) + GSH (5 mm), 4) GSH (5 mm), 5) l-cysteine (1 mm), 6) dl-homocysteine (1 mm), 7) lipoic acid (1 mm). Measurement conditions: 20 mm HEPES, pH 7.4, lex = 550 nm.
in the presence of 5 mm GSH. It should be noted that the fluorescence response of 6·CdII in the presence of GSH is apparently faster than those of most previously reported H2S probes, including 1·CdII.[7c, 8c, 9c] The rate acceleration induced by GSH might be due to the coordination of GSH to the CdII ion of 6·CdII, possibly inducing a change in the coordination geometry and making the complex more susceptible to nucleophilic attack by H2S. The sensing selectivity of 6·CdII toward biologically important thiol species was further examined. As shown in Figure 3 B, 6·CdII selectively responded to H2S with an increase of its R value, whereas significant changes in the R value were not induced by other physiologically abundant thiol species, such as cysteine (1 mm), homocysteine (1 mm), lipoic acid (1 mm) and GSH (5 mm). In addition, 6·CdII and ligand 6 did not show any significant pH-dependent ratio change over the physiological pH range (4.5 to 8.5; Figure S6), thus providing evidence for the accurate H2S sensing properties of 6·CdII under the different pH conditions in living cells. Live-cell imaging of hydrogen sulfide In subsequent work, the ratiometric fluorescence imaging of H2S in living cells was performed, taking advantage of the superior properties of 6·CdII, including selective, rapid, and accurate sensing. Unfortunately, 6·CdII was found to barely penetrate the cell membrane, likely because of its negatively charged, hydrophilic structure. We thus attempted to use a cell-penetrating peptide (CPP) to translocate the CdII complex to the cell interior. The carrier CR8 was designed with cysteine as a coordination site for 6·CdII, together with octa-arginine (R8), a well-known CPP sequence that has been used for delivering various molecular cargoes (Figure 4 A).[17] According to previous reports by Futaki et al.,[18] CR8 has been used in combination with pyrenebutyrate (PB) to deliver CdII complexes directly to the cytosol. When HeLa cells were sequentially treated with PB, 6·CdII and CR8 in HEPES-buffered saline solution, folChemBioChem 2015, 16, 1608 – 1615
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Figure 4. A) Structures of pyrenebutylate and the oligo-arginine peptides. B)–E) Fluorescence staining of HeLa cells with 6·CdII (30 mm) in the presence or absence of the oligo-arginine peptide (30 mm) and PB (50 mm): B) CR8 + PB at 37 8C, C) CR8 at 37 8C, D) CR8 + PB at 4 8C, and E) R8 + PB at 37 8C. Detailed conditions are given in the Experimental Section.
lowed by 20 min of incubation at 37 8C, bright fluorescence from 6·CdII was observed throughout the inside of the cells (Figure 4 B). Treatment of the cells with CR8 in the absence of PB gave weak, localized fluorescent spots inside the cells (Figure 4 C); this suggested that, in this case, 6·CdII was taken up in the endocytic vesicles. Treatment of the cells with CR8 and PB at 4 8C, at which temperature endocytosis does not occur, gave a diffuse fluorescence within the cell interiors similar to that observed at 37 8C (Figure 4 D). These results suggest that the main cell uptake pathway of 6·CdII is direct membrane translocation with the help of CPP and PB, as proposed by Futaki et al.[18] Interestingly, no fluorescence spots were observed when using R8 (without cysteine) as the CPP (Figure 4 E). These data imply that the coordination of 6·CdII with CR8 at its cysteine residues greatly facilitates the intracellular delivery of 6·CdII. The sensing ability of 6·CdII for H2S was subsequently examined in HeLa cells. A Trypan Blue viability test indicated that 6·CdII did not exhibit significant cell toxicity after introduction into cells by CR8 and PB (Figure S7). When cells stained with 6·CdII were treated with Na2S (150 mm final concentration), the R value (F580–600 nm/F610–630 nm) gradually increased from 0.87 to
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Figure 5. Ratiometric fluorescence imaging of H2S in HeLa cells with 6·CdII. Ratio images of the cells A) at before addition of Na2S, B) 10 min after addition of Na2S (150 mm), C) after 10 min without addition of Na2S. D) Time trace plot of the ratio value, R, of 6·CdII in HeLa cells in the presence (*) and absence (&) of Na2S (150 mm). n = 3; * P < 0.01 vs. absence of Na2S.
1.83 (DR = 0.96) over a 10 min interval (Figure 5 D).[19] This ratio value change was much larger than in the case of the previously developed probe 1·CdII, which displayed a ratio value change of DR = 0.1 under the same imaging conditions.[12a] In a control experiment without Na2S, the ratio value scarcely changed (Figure 5 D). These results indicate that 6·CdII effectively detects exogenous H2S inside cells, generating a large ratio change without interference from the intracellular environment. Ratiometric imaging of endogenously produced H2S in living cells We subsequently utilized 6·CdII to measure endogenously produced H2S in living cells. Szabo et al. recently reported that exposure of HUVEC cells to vascular endothelial growth factor (VEGF) stimulates the production of H2S by CSE.[20] The 6·CdII probe was successfully introduced into HUVEC cells by using CR8 and PB as carriers (Figure 6 A). When the cells were stimulated with VEGF, a gradual increase in R was observed inside the cells, up to a factor of 1.35 after 10 min (Figure 6 B and D). This value was greater than that observed for cells without stimulation (R10 min/R0 min = 1.25, Figure 6 C and D). Pre-treatment of the cells with propargyl glycine (PAG), an inhibitor of CSE, suppressed the R value increase (R10 min/R0 min) to a value of 1.19, thus suggesting that the basal production level of H2S decreased as a result of the inactivation of CSE by PAG. Furthermore, PAG effectively suppressed the VEGF-stimulated increase in the R value. We also confirmed that 6·CdII did not change its ChemBioChem 2015, 16, 1608 – 1615
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Figure 6. Ratiometric detection of endogenously produced H2S in HUVEC cells. Ratio images of the cells A) before addition of VEGF, B) 10 min after addition of VEGF (40 ng mL¢1 final conc.), and C) after 10 min without the addition of VEGF. D) Comparison of the ratio value changes in HUVEC cells in the presence or absence of the additives: 1) none, 2) VEGF (40 ng mL¢1), 3) PAG (100 mm), 4) VEGF (40 ng mL¢1) + PAG (100 mm). n = 4, * P < 0.05 vs. none and # P < 0.01 vs. VEGF + PAG.
ratio value in response to reactive oxygen and nitrogen species (ROS and RNS) such as H2O2, HOCl, COOH, NO and ONOO¢ , all of which are produced during VEGF stimulation of HUVEC cells (Figure S8).[7b, 21] These data suggest that 6·CdII allows measurement of the endogenously produced H2S in HUVEC cells based on its ratiometric fluorescence signal.
Conclusion We have developed a fluorescent CdII complex as a ratiometric probe for H2S based on arene–metal contact. The design strategy for this probe involved structural optimization of the coordination ligand and the fluorophore, wherein we were able to fully exploit the high compatibility of AM-contact sensing with various fluorophores and ligands. Evaluation of the sensing properties of several CdII complexes indicated that 6·CdII had the best resistance to high concentrations of GSH and allowed rapid and sensitive detection of H2S based on a significant ratio value change under in vivo conditions. The 6·CdII probe was also successfully transported into live cells by using the oligo-arginine peptide CR8 and PB as carriers, a technique originally developed for the intracellular delivery of macromolecules such as peptides and proteins.[17] We thus found that this technique can also be used to deliver the hydrophilic, polar metal ion complex into cells. The improved sensing ability of 6·CdII together with its efficient membrane translocation allowed us to detect endogenously produced H2S in live HUVEC cells. In the future, we intend to further improve the sensing
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Full Papers ability of our probe, with the particular aim of achieving a more rapid response to H2S. Eventually, this elaborate molecular design could provide a ratiometric probe that enables real-time detection of the rapid H2S flux in living cells.
Experimental Section Synthesis and characterization of the probes: Probes 1, 2, 3, and 5 were synthesized according to reported methods.[12a, 22] The syntheses and characterizations of probes 4 and 7 are described in the Supporting Information. Synthesis of 9: NaOH (4 n, 10 mL) was added dropwise to a cooled (0 8C) solution of 8[12a] (1.08 g, 1.19 mmol) in THF/MeOH (1:1, 15 mL). The reaction mixture was stirred at 0 8C for 90 min, then neutralized with HCl (1 n). After dilution with CHCl3, the organic layers were washed with brine. The solvent was removed in vacuo to give 9 (750 mg 86 %) as a yellow solid. 1H NMR (400 MHz, CD3OD): d = 1.41 (s, 36 H), 3.86 (s, 8 H), 4.66 (s, 4 H), 6.80 (s, 2 H), 7.0411 ppm (s, 2 H); MALDI-TOF-MS (CHCA): m/z for C39H54N2O12 : calcd: 743.38 [M+ +H] + ; found: 743.17. Synthesis of 10: A solution of trifluoromethanesulfonic anhydride (Tf2O; 852 mL, 4.85 mmol) in dry CH2Cl2 (20 mL) was added dropwise to a cooled (0 8C) solution of 9 (750 mg, 1.01 mmol) and pyridine (813 mL, 10.1 mmol) in dry CH2Cl2 (5 mL). The reaction mixture was stirred at RT for 90 min. After the reaction had been quenched with saturated NaHCO3, the mixture was extracted with CHCl3. The organic layers were washed with saturated NaHCO3 and dried over Na2SO4. After removal of the solvent in vacuo, the residue was purified by flash column chromatography on silica gel (CHCl3/ MeOH 30:1!15:1!10:1) to give 10 (840 mg, 82 %) as a brown solid. 1H NMR (400 MHz, CDCl3): d = 1.46 (s, 36 H), 3.48 (s, 8 H), 4.60 (s, 4 H), 7.29 (s, 2 H), 7.98 ppm (s, 2 H); ESI-TOF-MS: m/z for C41H52F6N2O16S2 : calcd: 1029.26 [M+ +Na] + ; found: 1029.23. Synthesis of 11: A solution of 10 (400 mg, 0.4 mmol) and pyrrolidine (368 mL, 4.49 mmol) in dry DMSO (5 mL) was stirred at 90 8C for 15 h.[14] After being cooled to RT, the reaction mixture was diluted with CHCl3 and washed with saturated NaHCO3, then dried over Na2SO4. After removal of the solvent in vacuo, the residue was purified by flash column chromatography on silica gel (CHCl3/ MeOH 60:1) to give 11 (68.5 mg, 20 %) as a white powder. 1H NMR (400 MHz, CDCl3): d = 1.46 (s, 36 H), 2.00–2.04 (br s, 8 H), 3.39–3.42 (br s, 8 H), 3.50 (s, 8 H), 4.60 (s, 4 H), 6.20 (s, 2 H), 7.16 ppm (s, 2 H); ESI-TOF-MS: m/z for C47H68N4O10 : calcd: 849.50 [M+ +H] + ; found: 849.50. Synthesis of 12: A solution of 11 (40.6 mg, 48 mmol) in dry THF (6 mL) was added dropwise to a solution of borane dimethylsulfide complex (485 mL, 0.96 mmol) in dry THF (0.48 mL). The reaction mixture was stirred at 40 8C for 2 h. Borane dimethylsulfide (240 mm, 480 mmol) in THF (2 m) was added dropwise, and the reaction mixture was stirred at 40 8C for 2 h. After the mixture had been cooled to RT, the reaction was quenched with water. The mixture was diluted with AcOEt and washed with saturated NaHCO3 and brine, then dried over Na2SO4. After removal of the solvent in vacuo, the residue was purified by flash column chromatography on silica gel (CHCl3/MeOH 60:1) to give 12 (25.2 mg, 63 %) as a purple oil. 1H NMR (400 MHz, CDCl3): d = 1.44 (s, 36 H), 1.98 (s, 8 H), 3.26 (s, 8 H), 3.47 (s, 8 H), 3.93 (s, 4 H), 3.97 (s, 2 H), 6.15 (s, 2 H), 6.36 ppm (s, 2 H); ESI-TOF-MS: m/z for C47H70N4O9 : calcd: 857.50 [M+ +Na] + ; found: 857.51. ChemBioChem 2015, 16, 1608 – 1615
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Synthesis of 13: A solution of DDQ (3.2 mg, 43.2 mmol) in dry EtOH (1 mL) was added to a solution of 12 (12.0 mg, 14.4 mmol) in dry EtOH (1 mL) and dry CH2Cl2(2 mL), and the reaction mixture was stirred at RT for 15 min. After removal of the solvent in vacuo, the residue was purified by flash column chromatography on silica gel (CHCl3/MeOH 20:1!10:1) to give 13 (9.3 mg, 78 %) as a purple oil. 1H NMR (400 MHz, CD3OD): d = 1.45 (s, 36 H), 2.15 (br s, 8 H), 3.48 (s, 8 H), 3.64 (s, 4 H), 4.34 (s, 4 H), 6.69 (s, 2 H), 7.10 (s, 2 H), 9.94 ppm (s, 1 H); ESI-TOF-MASS: m/z for C47H69N4O9 : calcd: 833.51 [M] + ; found: 833.50. Synthesis of 6: Trifluoroacetic acid (2 mL) was added dropwise to a cooled (0 8C) solution of 13 (9.3 mg, 11.1 mmol) in dry CH2Cl2 (2 mL). The mixture was stirred at RT for 29 h, then concentrated in vacuo. The residue was purified by reversed-phase HPLC (YMCPack, Triant-C18, 250 Õ 20 mm, acetonitrile/water (0.1 % TFA) 5:95! 50:50 (40 min)) to give 6 (1.7 mg, 25 %) as a purple solid. The purity of 6 was confirmed to be > 96 % by HPLC analysis. 1H NMR (400 MHz, CD3OD): d = 2.15 (br s, 8 H), 3.60 (s, 8 H), 3.64 (br s, 4 H), 4.40 (s, 4 H), 6.72 (s, 2 H), 7.09 (s, 2 H), 10.05 ppm (s, 1 H); ESI-TOFHRMS: m/z for C31H37N4O9 : calcd: 609.2555 [M] + , found: 609.2554. Fluorescence measurement: Fluorescence titration of the probe with CdCl2 was conducted with a solution of the probe (3 mL) in HEPES buffer (50 mm, pH 7.4)/MeOH (1:1). In the titration of the CdII complexes, 1 equiv of CdCl2 was added to an aqueous solution of the ligand before use. Fluorescence titration with Na2S was conducted with a solution of the CdII complex (3 mL) in HEPES buffer (20 mm, pH 7.4) at 37 8C. A freshly prepared aqueous stock solution of Na2S was added to the solution with a micropipette, and the fluorescence emission spectra were recorded after 5 min. In the case of titration with the thiol species (Figure 3 B), ROS, and RNS (Figure S8), the fluorescence spectra were recorded 30 min after addition of the analytes. Relative fluorescence quantum yields (F) of pyronine derivatives 6 and 7 were determined by in a solution of rhodamine B (F = 0.65) in EtOH as standard. Cell culture: HeLa cells were cultured in high-glucose Dulbecco’s Modified Eagle’s Medium (DMEM; Sigma) supplemented with 10 % fetal bovine serum (FBS; Gibco) and 1 % antibiotic-antimycotic solution (Gibco) at 37 8C under a humidified atmosphere of 5 % CO2 in air. HUVEC cells were cultured in Endothelial Cell Growth Medium (TaKaRa) supplemented with 2 % fetal calf medium, 0.4 % endothelial cell growth factor, epidermal growth factor (0.1 ng mL¢1), basic fibroblast growth factor (1 ng mL¢1), heparin (90 mg mL¢1), and hydrocortisone (1 mg mL¢1) at 37 8C under a humidified atmosphere of 5 % CO2 in air. A subculture was performed every 3–4 days from subconfluent (< 80 %) cultures by using a trypsin/EDTA solution. For the fluorescence bioimaging, cells (3.0 Õ 104 cells per well) were cultured for 2 days in a 35 mm glass-bottomed dish (Iwaki Scitech, Japan). Fluorescence imaging of exogenous H2S in HeLa cells: Fluorescence imaging was conducted with a confocal-laser scanning microscope equipped with a 60 Õ objective lens (A1Rsi, Nikon) or equipped with a 63 Õ objective lens (LSM 780, Zeiss). The following detection channels were chosen for the ratiometric imaging; Ch1 lex = 562 nm, lem = 580–600 nm, and Ch2 lex = 562 nm, lem = 610– 630 nm (A1Rsi), and lex = 561 nm, lem = 600–630 nm (LSM 780). In a glass-based dish, HeLa cells in modified HEPES buffer solution (HBS; 107 mm NaCl, 6 mm KCl, 11.5 mm glucose, 20 mm HEPES, pH 7.4) were first incubated with pyrenebutylate (50 mm) for 2 min then with 6·CdII (30 mm) for 5 min. The cells were further incubated with CR8 (30 mm) at 37 8C for 20 min. After being washed twice
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Full Papers with HBS, the cells were treated with Na2S (150 mm) and subjected to the fluorescence imaging. Fluorescence imaging of hydrogen sulfide in HUVEC cells: HUVEC cells were grown to 80 % confluence, washed with DMEM, and incubated for 8–12 h. They were then washed with modified HBS and incubated with a solution of 6·CdII (30 mm), CR8 (30 mm), and pyrene butylate (50 mm) for 20 min at 37 8C. After exchange of the medium to HEPES (20 mm; pH 7.4, containing NaCl (107 mm), KCl (6 mm), glucose (11.5 mm), CaCl2 (2.0 mm), and MgCl2 (1.2 mm)), the cells were treated with VEGF (40 ng mL¢1 in 0.1 % BSA/H2O, Invitrogen) and then subjected the fluorescence imaging. For the inhibition of CSE activity, HUVEC cells were pretreated with PAG (dl-propargylglycine, 100 mm, Sigma) in the modified HBS buffer for 10 min before being incubated with 6·CdII, and PAG (100 mm) was added again 10 min before VEGF addition.
Acknowledgements We appreciate technical support from the Research Support Center, Graduate School of Medical Sciences, Kyushu University. This work was performed under the Cooperative Research Program of the “Network Joint Research Center for Materials and Devices” organized by (MEXT, Japan). A.O. acknowledges the Toray Science Foundation for financial support. I.T. acknowledges the Japan Society for the Promotion of Science (JSPS) Research Fellowships for Young Scientists. Keywords: analytical methods · cadmium · fluorescent probes · hydrogen sulfide · imaging agents · noncovalent interactions [1] a) C. Szabû, Nat. Rev. Drug Discovery 2007, 6, 917 – 935; b) T. Ida, T. Sawa, H. Ihara, Y. Tsuchiya, Y. Watanabe, Y. Kumagai, M. Suematsu, H. Motohashi, S. Fujii, T. Matsunaga, M. Yamamoto, K. Ono, N. O. DevarieBaez, M. Xian, J. M. Fukuto, T. Akaike, Proc. Natl. Acad. Sci. USA 2014, 111, 7606 – 7611. [2] a) G. Yang, L. Wu, B. Jiang, W. Yang, J. Qi, K. Cao, Q. Meng, A. K. Mustafa, W. Mu, S. Zhang, S. H. Snyder, R. Wang, Science 2008, 322, 587 – 590; b) W. Zhao, J. Zhang, Y. Lu, R. Wang, EMBO J. 2001, 20, 6008 – 6016. [3] M. Nishida, T. Sawa, N. Kitajima, K. Ono, H. Inoue, H. Ihara, H. Motohashi, M. Yamamoto, M. Suematsu, H. Kurose, A. van der Vliet, B. A. Freeman, T. Shibata, K. Uchida, Y. Kumagai, T. Akaike, Nat. Chem. Biol. 2012, 8, 714 – 724. [4] a) A. Kazuho, K. Hideo, J. Neurosci. 1996, 16, 1066 – 1071; b) H. Kimura, Biochem. Biophys. Res. Commun. 2000, 267, 129 – 133. [5] P. Kamoun, Amino Acids 2004, 26, 243 – 254. [6] V. S. Lin, C. J. Chang, Curr. Opin. Chem. Biol. 2012, 16, 595 – 601. [7] a) A. R. Lippert, E. J. New, C. J. Chang, J. Am. Chem. Soc. 2011, 133, 10078 – 10080; b) V. S. Lin, A. R. Lippert, C. J. Chang, Proc. Natl. Acad. Sci. USA 2013, 110, 7131 – 7135; c) H. Peng, Y. Cheng, C. Dai, A. L. King, B. L. Predmore, D. J. Lefer, B. Wang, Angew. Chem. Int. Ed. 2011, 50, 9672 – 9675; Angew. Chem. 2011, 123, 9846 – 9849.
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[8] a) Y. Qian, L. Karpus, O. Kabil, S. Y. Zhang, H. L. Zhu, R. Banerjee, J. Zhao, C. He, Nat. Commun. 2011, 2, 495 – 501; b) C. Liu, J. Pan, S. Li, Y. Zhao, L. Y. Wu, C. E. Berkman, A. R. Whorton, M. Xian, Angew. Chem. Int. Ed. 2011, 50, 10327 – 10329; Angew. Chem. 2011, 123, 10511 – 10513; c) Y. Chen, C. Zhu, Z. Yang, J. Chen, Y. He, Y. Jiao, W. He, L. Qiu, J. Cen, Z. Guo, Angew. Chem. Int. Ed. 2013, 52, 1688 – 1691; Angew. Chem. 2013, 125, 1732 – 1735. [9] a) E. Galardon, A. Tomas, P. Roussel, I. Artaud, Dalton Trans. 2009, 9126 – 9130; b) M. G. Choi, S. Cha, H. Lee, H. L. Jeon, S.-K. Chang, Chem. Commun. 2009, 7390 – 7392; c) K. Sasakura, K. Hanaoka, N. Shibuya, Y. Mikami, Y. Kimura, T. Komatsu, T. Ueno, T. Terai, H. Kimura, T. Nagano, J. Am. Chem. Soc. 2011, 133, 18003 – 18005. [10] G. Grynkiewicz, M. Poenie, R. Y. Tsien, J. Biol. Chem. 1985, 260, 3440 – 3450. [11] a) F. Yu, P. Li, P. Song, B. Wang, J. Zhao, K. Han, Chem. Commun. 2012, 48, 2852 – 2854; b) S. K. Bae, C. H. Heo, D. J. Choi, D. Sen, E.-H. Joe, B. R. Cho, H. M. Kim, J. Am. Chem. Soc. 2013, 135, 9915 – 9923; c) X. Wang, J. Sun, W. Zhang, X. Ma, J. Lv, B. Tang, Chem. Sci. 2013, 4, 2551 – 2556. [12] a) I. Takashima, M. Kinoshita, R. Kawagoe, S. Nakagawa, M. Sugimoto, I. Hamachi, A. Ojida, Chem. Eur. J. 2014, 20, 2184 – 2192; b) I. Takashima, A. Kanegae, M. Sugimoto, A. Ojida, Inorg. Chem. 2014, 53, 7080 – 7082. [13] a) S. Kenmoku, Y. Urano, H. Kojima, T. Nagano, J. Am. Chem. Soc. 2007, 129, 7313 – 7318; b) A. Ojida, I. Takashima, T. Kohira, H. Nonaka, I. Hamachi, J. Am. Chem. Soc. 2008, 130, 12095 – 12101; c) H. N. Kim, M. H. Lee, H. J. Kim, J. S. Kim, J. Yoon, Chem. Soc. Rev. 2008, 37, 1465 – 1472; d) M. Sakabe, D. Asanuma, M. Kamiya, R. J. Iwatate, K. Hanaoka, T. Terai, T. Nagano, Y. Urano, J. Am. Chem. Soc. 2013, 135, 409 – 414. [14] L. Wu, K. Burgess, J. Org. Chem. 2008, 73, 8711 – 8718. [15] In the optimized geometry of 6’, the both pyrrolidine rings exist in the almost same plane of the xanthene ring (average twisted angle = 3.98), whereas the N-acetylpiperazine rings of 7’ have a twisted conformation (average twisted angle = 17.98). The planer conformation of 6’ might facilitate conjugation of the sp2 lone pairs of the pyrroridine nitrogens with the pyronine ring so that the LUMO energy level of 6’ is higher than that of 7’. [16] a) W. L. Jorgensen, J. F. Blake, D. C. Lim, D. L. Severance, J. Chem. Soc. Faraday Trans. 1994, 90, 1727 – 1732; b) D. L. Severance, W. L. Jorgensen, J. Am. Chem. Soc. 1992, 114, 10966 – 10968. [17] S. Futaki, Adv. Drug Delivery Rev. 2005, 57, 547 – 558. [18] a) F. Perret, M. Nishihara, T. Takeuchi, S. Futaki, A. N. Lazar, A. W. Coleman, N. Sakai, S. Matile, J. Am. Chem. Soc. 2005, 127, 1114 – 1115; b) T. Takeuchi, M. Kosuge, A. Tadokoro, Y. Sugiura, M. Nishi, M. Kawata, N. Sakai, S. Matile, S. Futaki, ACS Chem. Biol. 2006, 1, 299 – 303. [19] Treatment of HeLa cells with the CdII·pyrithione complex (10 mm) and Na2S (150 mm) for 1 h did not lead to a significant change in cell viability (Figure S7), thus suggesting that CdII released from 6·CdII as Na2S exerts little toxicity on the cells. [20] A. Papapetropoulos, A. Pyriochou, Z. Altaany, G. Yang, A. Marazioti, Z. Zhou, M. G. Jeschke, L. K. Branski, D. N. Herndon, R. Wang, C. Szabû, Proc. Natl. Acad. Sci. USA 2009, 106, 21972 – 21977. [21] A. Papapetropoulos, G. Garca-CardeÇa, J. A. Madri, W. C. Sessa, J. Clin. Invest. 1997, 100, 3131 – 3139. [22] M. Choi, M. Kim, K. D. Lee, K.-N. Han, I.-A. Yoon, H.-J. Chung, J. Yoon, Org. Lett. 2001, 3, 3455 – 3457.
Manuscript received: May 17, 2015 Accepted article published: May 28, 2015 Final article published: June 26, 2015
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Rational Design of a Ratiometric Fluorescent Probe Based on Arene–Metal-Ion Contact for Endogenous Hydrogen Sulfide Detection in Living Cells Ryosuke Kawagoe, Ippei Takashima, Kazuteru Usui, Anna Kanegae, Yusuke Ozawa, and Akio Ojida*[a] We report the design and development of a fluorescent CdII ion complex that is capable of the ratiometric detection of H2S in living cells. This probe exploits the metal-ion-induced emission red shift resulting from direct contact between the aromatic ring of a fluorophore and a metal ion (i.e., arene–metalion or “AM” contact). The CdII complex displays a large emission blue shift upon interaction with H2S as the CdII-free ligand is released by the formation of cadmium sulfide. Screening of potential ligands and fluorophores led to the discovery of a py-
ronine-type probe, 6·CdII, that generated a sensitive and rapid ratio value change upon interaction with H2S, without interference from the glutathione that is abundant in the cell. The membrane-impermeable 6·CdII was successfully translocated into live cells by using an oligo-arginine peptide and pyrenebutylate as carriers. As such, 6·CdII was successfully applied to the ratiometric detection of both exogenous and endogenous H2S produced by the enzymes in living cells, thus demonstrating the utility of 6·CdII in biological fluorescence analysis.
Introduction Hydrogen sulfide (H2S) is the most recently identified gaseous signaling molecule in living organisms.[1] Although H2S has long been considered a toxic compound, there is now an accumulation of scientific evidence that this redox-active molecule plays important metabolic roles, for example in angiogenesis,[2] cardioprotection[3] and the induction of hippocampal longterm potentiation.[4] In mammals, H2S is enzymatically produced by cystathionine-b-synthase, cystathionine-g-lyase (CSE), and cysteine amino transferase in combination with 3-mercaptosulfurtransferase.[5] To better understand the biological functions of this compound, it is critical to develop a noninvasive real-time detection method for H2S in living cell systems, and thus a number of fluorescent probes for H2S have been developed in recent years.[6] These probes were designed based on three different sensing mechanisms: H2S-mediated azide reduction,[7] nucleophilic reaction of H2S,[8] and metal–sulfide complex precipitation.[9] Most such probes were designed to detect H2S levels by increases in single fluorescence emission. However, single-emission sensing is readily perturbed by various aspects of the cellular environment, such as intracellular pH, cell volume, and inhomogeneous probe localization, thus making it difficult to quantitatively detect H2S under in vivo conditions. Ratiometric fluorescence sensing, which is based on the simultaneous detection of two emission signals, can eliminate these unfavorable effects and allows quantitative [a] R. Kawagoe, I. Takashima, Dr. K. Usui, A. Kanegae, Y. Ozawa, Prof. Dr. A. Ojida Graduate School of Pharmaceutical Sciences, Kyushu University 3-1-1 Maidashi, Higashi-ku, Fukuoka 812-8582 (Japan) E-mail: [email protected] Supporting information for this article is available on the WWW under http://dx.doi.org/10.1002/cbic.201500249.
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data analysis with significant precision.[10] Several ratiometric probes for H2S have been reported in the last few years.[8c, 11] Unfortunately, despite having elaborate molecular designs, these probes still suffer from limited fluorescence, slow responses to the H2S flux, or an inability to detect endogenously produced H2S. Recently, we reported the unique fluorescence sensing of metal ions based on direct contact between the aromatic ring of a fluorophore and a metal ion (i.e., arene–metal-ion or “AM” contact).[12] Fluorescent probes with this sensing mechanism exhibited large emission red shifts upon coordination with various metal ions, including CdII, AgI, HgII, and PbII. By exploiting AM-contact sensing, we previously developed the xanthenetype CdII complex 1·CdII as a ratiometric fluorescence probe for H2S (Scheme 1).[12a] The 1·CdII system was able to sense H2S based on the significant emission blue shift (DF = 18 nm) induced by loss of the arene–metal-ion contact when the CdII ion is released from 1·CdII to form cadmium sulfide. Although 1·CdII could be applied to the ratiometric detection of exogenously supplied H2S in live cells, there was unexpectedly strong quenching of the fluorescence emission by glutathione (GSH). This quenching, the resulting weak fluorescence signal, and associated small change in signal ratio significantly diminished the usefulness of 1·CdII, and precluded its further application to the detection of endogenously produced H2S in living cells. In this paper, we report the design of a ratiometric fluorescent probe for H2S with improved sensing properties. Careful optimization of both the ligand and fluorophore structures provided a new CdII complex, 6·CdII, that functioned as a highly sensitive fluorescent probe for H2S, rapidly generating a large ratio change. Notably, the fluorescence of 6·CdII was not quenched by high concentrations of GSH (5 mm).
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Full Papers different ligand units (Scheme 2). The syntheses of these probes are described in the Supporting Information (Scheme S1). The probes’ fluorescence spectral changes upon the addition of H2S are summarized in Figure S2; Table 1 gives the fluorescence responses of the probes for H2S (30 mm) and GSH (5 mm) under aqueous MeOH conditions (50 mm HEPES, pH 7.4/MeOH 1:1). Probe 2·CdII, which has two 2,2’-dipicolylamine (Dpa) ligands, exhibited Scheme 1. Ratiometric fluorescence sensing of 1·CdII for H2S based on arene-metal ion a large emission blue shift (DF = ¢16 nm) upon the contact. addition of H2S. A significant emission shift (DF = ¢8 nm) was also observed in the case of 3·CdII, with Given these superior sensing properties, 6·CdII was successfully iminodiacetic acid (Ida) ligands, whereas the emission blue applied to the ratiometric detection of not only exogenously shift of 4·CdII, which has 2-picolylaminoacetic acid (Paa) lisupplied H2S, but also the endogenously produced H2S in gands, was small (DF = ¢2 nm). The large emission blue shifts living cells, thus highlighting its usefulness in biological fluorescence analysis.
Result and Discussion Screening of the ligand unit Figure 1 presents the fluorescence and absorption spectral changes of 1·CdII (10 mm, 20 mm HEPES, pH 7.4) upon the addition of high concentrations of GSH (0–5 mm). The significant quenching of the fluorescence concomitant with the decrease in absorbance around 500 nm suggests that the conjugated structure of the xanthene ring of 1·CdII was disrupted by the presence of GSH. Analysis of a mixture of 1·CdII and GSH by ESI-MS identified a new product with a m/z value of 1054, corresponding to the GSH adduct of 1·CdII (Figure S1 in the Supporting Information). Based on these spectral data, we concluded that the GSH-induced fluorescence quenching primarily results from nucleophilic attack of GSH on the xanthene ring of 1·CdII at the C9 carbon, which is known to be susceptible to such attack.[13]
Scheme 2. Fluorescent CdII complexes for ratiometric detection of H2S.
Table 1. Summary of the fluorescence response of the probes toward H2S and glutathione (GSH).
Figure 1. A) Absorption and B) fluorescence spectral changes of 1·CdII (10 mm) upon addition of glutathione (GSH, 0–5 mm). Measurement conditions: 20 mm HEPES, pH 7.4, lex = 500 nm.
Initially, we attempted to find suitable ligands for the probe that were capable of preventing the nucleophilic attack of GSH on the fluorophore. Since AM-contact sensing can work with various fluorophores,[12a] we prepared the readily available anthracene-type probes 2·CdII to 4·CdII, each of which possessed ChemBioChem 2015, 16, 1608 – 1615
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DF [nm][a] FGSH/Fo[b]
1·CdII
2·CdII
3·CdII
4·CdII
¢21 0.16
¢16 0.79
¢8 1.01
¢2 0.98
[a] Shift of the emission maximum wavelength upon addition of Na2S: 30 mm for 1·CdII, 100 mm for 2·CdII, 3·CdII, and 4·CdII. [b] Relative fluorescence intensity in the absence (Fo) and in presence of 5 mm GSH (FGSH), lex = 500 nm (1·CdII), lex = 360 nm (2·CdII, 3·CdII, 4·CdII).
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Full Papers seen for 2·CdII and 3·CdII in the presence of H2S were attributed to the intrinsically large emission red shifts of ligands 2 (DF = 15 nm) and 3 (DF = 10 nm) induced by CdII coordination, whereas the emission red shift of 4 upon CdII complexation is minimal (DF = 5 nm). These data suggest that the structure of the ligand unit, which assists the direct contact between the CdII and the fluorophore, can influence the ratiometric sensing property of the probe. In contrast to the xanthene-type probe 1·CdII, the anthracene-type CdII complexes showed very little decrease in fluorescence in the presence of 5 mm GSH (Table 1, Figure S2). This can be explained by the inertness of the anthracene ring toward the nucleophilic attack of GSH at the C9 position. We subsequently used isothermal titration calorimetry (ITC) to evaluate the coordination interactions of the anthracene-type probes with GSH. In the case of 2·CdII, which has Dpa ligands, endothermic binding was observed upon titration with GSH, and the binding constant (Ka) was found to be 5.48 Õ 104 m¢1 (n = 0.57) by curve-fitting analysis (Figure S3). We believe that this coordination interaction could enhance the nucleophilic attack of GSH on the xanthene ring of 1·CdII. In contrast, the ITC curves of 3·CdII and 4·CdII upon titration with GSH exhibited minimal change, thus suggesting that the binding affinities of these compounds are very weak (Ka < 103 m¢1). The weak binding affinities of 3·CdII and 4·CdII could result from the negative charges of the Ida and Paa ligands, which attenuate the electrostatic interaction between the anionic GSH and the coordinated CdII ions. Overall, these fluorescence and ITC data indicate that the Ida ligand in 3·CdII, which displays a large
emission blue shift upon binding with H2S and also reduces unwanted interactions with GSH, is the most suitable ligand for the ratiometric detection of H2S. Optimization of the fluorophore We next sought to optimize the fluorophore of the CdII complex to obtain a probe capable of reacting to H2S with a large emission blue shift, while remaining resistant to GSH-induced fluorescence quenching. The CdII complexes 5·CdII to 7·CdII were each designed with a different xanthene fluorophore (Scheme 2). Given the results of the ligand-optimization work, these probes had Ida as ligands. The synthetic route to the pyronine-type ligand 6 is given in Scheme 3. Briefly, the 4,7-dihydroxyxanthone 9 obtained from 8[12a] by deprotection of the pivaloyl groups was converted to ditriflate 10, which was treated with pyrrolidine to give the 4,7-diaminoxanthone 11.[14] After reduction with borane-dimethylsulfide complex, 12 was oxidized by 2,3-dichloro-5,6-dicyano-1,4-benzoquinone (DDQ) to afford pyronine 13. Finally, deprotection of tert-butyl esters and subsequent HPLC purification provided the pure ligand 6. The photophysical properties of the ligands and their CdII complexes are summarized in Table S1. In CdCl2 titration (Figure S4), each ligand 5 to 7 displayed a large emission red shift upon coordination with CdII ions, together with simultaneous absorption red shifts; these are indicative of contact interaction between the CdII ions and the fluorophores. Figures 2 A and S5 show the fluorescence responses of 5·CdII to 7·CdII toward H2S under aqueous conditions. All probes displayed
Scheme 3. Synthesis of ligand 6. a) 4 n NaOH, THF/MeOH (86 %); b) Tf2O, pyridine, CH2Cl2 (82 %); c) pyrrolidine, DMSO (20 %); d) BH3SMe2, THF (63 %); e) DDQ, EtOH/CH2Cl2 (78 %); f) TFA, CH2Cl2 ; g) HPLC (25 %).
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Figure 2. A) Change in fluorescence spectrum of 6·CdII (10 mm) upon addition of Na2S (0– 20 mm). Measurement conditions: 20 mm HEPES, pH 7.4, lex = 550 nm. B) Plot of the ratio value, R, of 6·CdII (10 mm) upon addition of Na2S. Measurement conditions: 20 mm HEPES, pH 7.4, lex = 550 nm. C) Change in the ratio value (DR) of the CdII complexes (10 mm) upon addition of Na2S (30 mm). 1·CdII : DR = F527 nm/F549 nm, 5·CdII : DR = F528 nm/ F546 nm, 7·CdII : DR = F593 nm/F613 nm. Measurement conditions: 20 mm HEPES, pH 7.4, lex = 500 nm (1·CdII and 5·CdII), lex = 550 nm (6·CdII and 7·CdII). D) Change in the relative fluorescence intensity (F/Fo) of 1·CdII (*), 5·CdII (&), 6·CdII (~), and 7·CdII (^) upon addition of GSH (0–5 mm). Measurement conditions: [probe] = 10 mm, 20 mm HEPES, pH 7.4.
large emission blue shifts upon the addition of sodium sulfide (Na2S) as a donor of H2S, leading to changes in the fluorescence ratio value R, as shown in Figure 2 B and C. Simultaneously, the absorption spectra of the probes were hypsochromically shifted (Figure S5); this suggests that the emission shifts were induced by the loss of AM-contact as a result of the release of CdII-free ligands. Most intriguingly, the probes showed different magnitudes of fluorescence quenching response to GSH. Figure 2 D summarizes the changes in the fluorescence intensities of the probes upon the addition of 0–5 mm GSH. The decrease in the fluorescence intensity of 5·CdII was less than that of the previously developed 1·CdII, thus suggesting that 5·CdII was less reactive toward GSH than 1·CdII was. This could possibly be ascribed to the lower binding affinity of the Ida·CdII complex with GSH compared to that of the Dpa·CdII complex, as was observed in the case of anthracene-type CdII complexes (Figure S3). Interestingly, the pyronine-type probe 7·CdII, with Nacetylpiperazine ligands, lost essentially all fluorescence upon the addition of 2 mm of GSH, whereas the pyronine-type probe 6·CdII, with pyrrolidine ligands, maintained its original fluorescence even in the presence of 5 mm of GSH. We speculate that the different reactivities of these complexes can be explained in terms of the LUMO energy levels of the fluorophores, as reported by Urano et al.[13d] Scheme 4 gives the calculated LUMO energy levels of fluorophores 5’ to 7’, all of which are analogues of the fluorophores of 5·CdII to 7·CdII. Density functional theory calculations suggest that 6’ has ChemBioChem 2015, 16, 1608 – 1615
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a higher LUMO energy level than 7’; this is indicative of the lower electrophilic reactivity of 6·CdII relative to 7·CdII,[15] and is consistent with the experimental results, which show a higher tolerance of 6·CdII for GSH. The LUMO energy level of the xanthene-based 5’ was calculated to be higher than that of 6’, which cannot explain the higher reactivity of 5·CdII with GSH compared with that of 6·CdII. However, when the hydrogen bonding of the carbonyl oxygen of 5’ with surrounding water molecules (according to Jorgensen’s model[16]) were taken into account, the LUMO energy level of 5’ was predicted to be lower than that of 6’, and this is consistent with the experimental observations of the higher resistance of 6·CdII toward GSH. Overall, the data in Figure 2 suggest that 6·CdII is the most suitable probe for H2S sensing among the three CdII complexes. The sensing properties of 6·CdII were further evaluated by fluorescence titration experiments. Figure 3 A plots the time-dependent fluorescence response of 6·CdII toward H2S in the presence and absence of GSH. In the absence of GSH, the fluorescence ratio value R (F599 nm/F619 nm) of 6·CdII gradually increased and reached a plateau (R = 2.0) 5 min after treatment with Na2S (a donor of gaseous H2S); no ratio change was induced by GSH (5 mm) even after 10 min. Interestingly, the response of 6·CdII toward H2S was accelerated by the presence of GSH; the R value rapidly (within ca. 1 min) increased to 2.0
Scheme 4. LUMO energy levels of the fluorophores from DFT calculations.
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Figure 3. A) Time trace plot of the ratio value, R, of 6·CdII (10 mm) upon addition 5 mm of GSH (~), and upon addition of 30 mm of Na2S in the presence (&) and absence (*) of 5 mm of GSH. Measurement conditions: 20 mm HEPES, pH 7.4, lex = 550 nm. The error bars represent standard deviations obtained from triplicate experiments. B) The ratio value of 6·CdII (10 mm) in the presence of various thiol species; 1) none, 2) Na2S (30 mm), 3) Na2S (30 mm) + GSH (5 mm), 4) GSH (5 mm), 5) l-cysteine (1 mm), 6) dl-homocysteine (1 mm), 7) lipoic acid (1 mm). Measurement conditions: 20 mm HEPES, pH 7.4, lex = 550 nm.
in the presence of 5 mm GSH. It should be noted that the fluorescence response of 6·CdII in the presence of GSH is apparently faster than those of most previously reported H2S probes, including 1·CdII.[7c, 8c, 9c] The rate acceleration induced by GSH might be due to the coordination of GSH to the CdII ion of 6·CdII, possibly inducing a change in the coordination geometry and making the complex more susceptible to nucleophilic attack by H2S. The sensing selectivity of 6·CdII toward biologically important thiol species was further examined. As shown in Figure 3 B, 6·CdII selectively responded to H2S with an increase of its R value, whereas significant changes in the R value were not induced by other physiologically abundant thiol species, such as cysteine (1 mm), homocysteine (1 mm), lipoic acid (1 mm) and GSH (5 mm). In addition, 6·CdII and ligand 6 did not show any significant pH-dependent ratio change over the physiological pH range (4.5 to 8.5; Figure S6), thus providing evidence for the accurate H2S sensing properties of 6·CdII under the different pH conditions in living cells. Live-cell imaging of hydrogen sulfide In subsequent work, the ratiometric fluorescence imaging of H2S in living cells was performed, taking advantage of the superior properties of 6·CdII, including selective, rapid, and accurate sensing. Unfortunately, 6·CdII was found to barely penetrate the cell membrane, likely because of its negatively charged, hydrophilic structure. We thus attempted to use a cell-penetrating peptide (CPP) to translocate the CdII complex to the cell interior. The carrier CR8 was designed with cysteine as a coordination site for 6·CdII, together with octa-arginine (R8), a well-known CPP sequence that has been used for delivering various molecular cargoes (Figure 4 A).[17] According to previous reports by Futaki et al.,[18] CR8 has been used in combination with pyrenebutyrate (PB) to deliver CdII complexes directly to the cytosol. When HeLa cells were sequentially treated with PB, 6·CdII and CR8 in HEPES-buffered saline solution, folChemBioChem 2015, 16, 1608 – 1615
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Figure 4. A) Structures of pyrenebutylate and the oligo-arginine peptides. B)–E) Fluorescence staining of HeLa cells with 6·CdII (30 mm) in the presence or absence of the oligo-arginine peptide (30 mm) and PB (50 mm): B) CR8 + PB at 37 8C, C) CR8 at 37 8C, D) CR8 + PB at 4 8C, and E) R8 + PB at 37 8C. Detailed conditions are given in the Experimental Section.
lowed by 20 min of incubation at 37 8C, bright fluorescence from 6·CdII was observed throughout the inside of the cells (Figure 4 B). Treatment of the cells with CR8 in the absence of PB gave weak, localized fluorescent spots inside the cells (Figure 4 C); this suggested that, in this case, 6·CdII was taken up in the endocytic vesicles. Treatment of the cells with CR8 and PB at 4 8C, at which temperature endocytosis does not occur, gave a diffuse fluorescence within the cell interiors similar to that observed at 37 8C (Figure 4 D). These results suggest that the main cell uptake pathway of 6·CdII is direct membrane translocation with the help of CPP and PB, as proposed by Futaki et al.[18] Interestingly, no fluorescence spots were observed when using R8 (without cysteine) as the CPP (Figure 4 E). These data imply that the coordination of 6·CdII with CR8 at its cysteine residues greatly facilitates the intracellular delivery of 6·CdII. The sensing ability of 6·CdII for H2S was subsequently examined in HeLa cells. A Trypan Blue viability test indicated that 6·CdII did not exhibit significant cell toxicity after introduction into cells by CR8 and PB (Figure S7). When cells stained with 6·CdII were treated with Na2S (150 mm final concentration), the R value (F580–600 nm/F610–630 nm) gradually increased from 0.87 to
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Figure 5. Ratiometric fluorescence imaging of H2S in HeLa cells with 6·CdII. Ratio images of the cells A) at before addition of Na2S, B) 10 min after addition of Na2S (150 mm), C) after 10 min without addition of Na2S. D) Time trace plot of the ratio value, R, of 6·CdII in HeLa cells in the presence (*) and absence (&) of Na2S (150 mm). n = 3; * P < 0.01 vs. absence of Na2S.
1.83 (DR = 0.96) over a 10 min interval (Figure 5 D).[19] This ratio value change was much larger than in the case of the previously developed probe 1·CdII, which displayed a ratio value change of DR = 0.1 under the same imaging conditions.[12a] In a control experiment without Na2S, the ratio value scarcely changed (Figure 5 D). These results indicate that 6·CdII effectively detects exogenous H2S inside cells, generating a large ratio change without interference from the intracellular environment. Ratiometric imaging of endogenously produced H2S in living cells We subsequently utilized 6·CdII to measure endogenously produced H2S in living cells. Szabo et al. recently reported that exposure of HUVEC cells to vascular endothelial growth factor (VEGF) stimulates the production of H2S by CSE.[20] The 6·CdII probe was successfully introduced into HUVEC cells by using CR8 and PB as carriers (Figure 6 A). When the cells were stimulated with VEGF, a gradual increase in R was observed inside the cells, up to a factor of 1.35 after 10 min (Figure 6 B and D). This value was greater than that observed for cells without stimulation (R10 min/R0 min = 1.25, Figure 6 C and D). Pre-treatment of the cells with propargyl glycine (PAG), an inhibitor of CSE, suppressed the R value increase (R10 min/R0 min) to a value of 1.19, thus suggesting that the basal production level of H2S decreased as a result of the inactivation of CSE by PAG. Furthermore, PAG effectively suppressed the VEGF-stimulated increase in the R value. We also confirmed that 6·CdII did not change its ChemBioChem 2015, 16, 1608 – 1615
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Figure 6. Ratiometric detection of endogenously produced H2S in HUVEC cells. Ratio images of the cells A) before addition of VEGF, B) 10 min after addition of VEGF (40 ng mL¢1 final conc.), and C) after 10 min without the addition of VEGF. D) Comparison of the ratio value changes in HUVEC cells in the presence or absence of the additives: 1) none, 2) VEGF (40 ng mL¢1), 3) PAG (100 mm), 4) VEGF (40 ng mL¢1) + PAG (100 mm). n = 4, * P < 0.05 vs. none and # P < 0.01 vs. VEGF + PAG.
ratio value in response to reactive oxygen and nitrogen species (ROS and RNS) such as H2O2, HOCl, COOH, NO and ONOO¢ , all of which are produced during VEGF stimulation of HUVEC cells (Figure S8).[7b, 21] These data suggest that 6·CdII allows measurement of the endogenously produced H2S in HUVEC cells based on its ratiometric fluorescence signal.
Conclusion We have developed a fluorescent CdII complex as a ratiometric probe for H2S based on arene–metal contact. The design strategy for this probe involved structural optimization of the coordination ligand and the fluorophore, wherein we were able to fully exploit the high compatibility of AM-contact sensing with various fluorophores and ligands. Evaluation of the sensing properties of several CdII complexes indicated that 6·CdII had the best resistance to high concentrations of GSH and allowed rapid and sensitive detection of H2S based on a significant ratio value change under in vivo conditions. The 6·CdII probe was also successfully transported into live cells by using the oligo-arginine peptide CR8 and PB as carriers, a technique originally developed for the intracellular delivery of macromolecules such as peptides and proteins.[17] We thus found that this technique can also be used to deliver the hydrophilic, polar metal ion complex into cells. The improved sensing ability of 6·CdII together with its efficient membrane translocation allowed us to detect endogenously produced H2S in live HUVEC cells. In the future, we intend to further improve the sensing
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Full Papers ability of our probe, with the particular aim of achieving a more rapid response to H2S. Eventually, this elaborate molecular design could provide a ratiometric probe that enables real-time detection of the rapid H2S flux in living cells.
Experimental Section Synthesis and characterization of the probes: Probes 1, 2, 3, and 5 were synthesized according to reported methods.[12a, 22] The syntheses and characterizations of probes 4 and 7 are described in the Supporting Information. Synthesis of 9: NaOH (4 n, 10 mL) was added dropwise to a cooled (0 8C) solution of 8[12a] (1.08 g, 1.19 mmol) in THF/MeOH (1:1, 15 mL). The reaction mixture was stirred at 0 8C for 90 min, then neutralized with HCl (1 n). After dilution with CHCl3, the organic layers were washed with brine. The solvent was removed in vacuo to give 9 (750 mg 86 %) as a yellow solid. 1H NMR (400 MHz, CD3OD): d = 1.41 (s, 36 H), 3.86 (s, 8 H), 4.66 (s, 4 H), 6.80 (s, 2 H), 7.0411 ppm (s, 2 H); MALDI-TOF-MS (CHCA): m/z for C39H54N2O12 : calcd: 743.38 [M+ +H] + ; found: 743.17. Synthesis of 10: A solution of trifluoromethanesulfonic anhydride (Tf2O; 852 mL, 4.85 mmol) in dry CH2Cl2 (20 mL) was added dropwise to a cooled (0 8C) solution of 9 (750 mg, 1.01 mmol) and pyridine (813 mL, 10.1 mmol) in dry CH2Cl2 (5 mL). The reaction mixture was stirred at RT for 90 min. After the reaction had been quenched with saturated NaHCO3, the mixture was extracted with CHCl3. The organic layers were washed with saturated NaHCO3 and dried over Na2SO4. After removal of the solvent in vacuo, the residue was purified by flash column chromatography on silica gel (CHCl3/ MeOH 30:1!15:1!10:1) to give 10 (840 mg, 82 %) as a brown solid. 1H NMR (400 MHz, CDCl3): d = 1.46 (s, 36 H), 3.48 (s, 8 H), 4.60 (s, 4 H), 7.29 (s, 2 H), 7.98 ppm (s, 2 H); ESI-TOF-MS: m/z for C41H52F6N2O16S2 : calcd: 1029.26 [M+ +Na] + ; found: 1029.23. Synthesis of 11: A solution of 10 (400 mg, 0.4 mmol) and pyrrolidine (368 mL, 4.49 mmol) in dry DMSO (5 mL) was stirred at 90 8C for 15 h.[14] After being cooled to RT, the reaction mixture was diluted with CHCl3 and washed with saturated NaHCO3, then dried over Na2SO4. After removal of the solvent in vacuo, the residue was purified by flash column chromatography on silica gel (CHCl3/ MeOH 60:1) to give 11 (68.5 mg, 20 %) as a white powder. 1H NMR (400 MHz, CDCl3): d = 1.46 (s, 36 H), 2.00–2.04 (br s, 8 H), 3.39–3.42 (br s, 8 H), 3.50 (s, 8 H), 4.60 (s, 4 H), 6.20 (s, 2 H), 7.16 ppm (s, 2 H); ESI-TOF-MS: m/z for C47H68N4O10 : calcd: 849.50 [M+ +H] + ; found: 849.50. Synthesis of 12: A solution of 11 (40.6 mg, 48 mmol) in dry THF (6 mL) was added dropwise to a solution of borane dimethylsulfide complex (485 mL, 0.96 mmol) in dry THF (0.48 mL). The reaction mixture was stirred at 40 8C for 2 h. Borane dimethylsulfide (240 mm, 480 mmol) in THF (2 m) was added dropwise, and the reaction mixture was stirred at 40 8C for 2 h. After the mixture had been cooled to RT, the reaction was quenched with water. The mixture was diluted with AcOEt and washed with saturated NaHCO3 and brine, then dried over Na2SO4. After removal of the solvent in vacuo, the residue was purified by flash column chromatography on silica gel (CHCl3/MeOH 60:1) to give 12 (25.2 mg, 63 %) as a purple oil. 1H NMR (400 MHz, CDCl3): d = 1.44 (s, 36 H), 1.98 (s, 8 H), 3.26 (s, 8 H), 3.47 (s, 8 H), 3.93 (s, 4 H), 3.97 (s, 2 H), 6.15 (s, 2 H), 6.36 ppm (s, 2 H); ESI-TOF-MS: m/z for C47H70N4O9 : calcd: 857.50 [M+ +Na] + ; found: 857.51. ChemBioChem 2015, 16, 1608 – 1615
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Synthesis of 13: A solution of DDQ (3.2 mg, 43.2 mmol) in dry EtOH (1 mL) was added to a solution of 12 (12.0 mg, 14.4 mmol) in dry EtOH (1 mL) and dry CH2Cl2(2 mL), and the reaction mixture was stirred at RT for 15 min. After removal of the solvent in vacuo, the residue was purified by flash column chromatography on silica gel (CHCl3/MeOH 20:1!10:1) to give 13 (9.3 mg, 78 %) as a purple oil. 1H NMR (400 MHz, CD3OD): d = 1.45 (s, 36 H), 2.15 (br s, 8 H), 3.48 (s, 8 H), 3.64 (s, 4 H), 4.34 (s, 4 H), 6.69 (s, 2 H), 7.10 (s, 2 H), 9.94 ppm (s, 1 H); ESI-TOF-MASS: m/z for C47H69N4O9 : calcd: 833.51 [M] + ; found: 833.50. Synthesis of 6: Trifluoroacetic acid (2 mL) was added dropwise to a cooled (0 8C) solution of 13 (9.3 mg, 11.1 mmol) in dry CH2Cl2 (2 mL). The mixture was stirred at RT for 29 h, then concentrated in vacuo. The residue was purified by reversed-phase HPLC (YMCPack, Triant-C18, 250 Õ 20 mm, acetonitrile/water (0.1 % TFA) 5:95! 50:50 (40 min)) to give 6 (1.7 mg, 25 %) as a purple solid. The purity of 6 was confirmed to be > 96 % by HPLC analysis. 1H NMR (400 MHz, CD3OD): d = 2.15 (br s, 8 H), 3.60 (s, 8 H), 3.64 (br s, 4 H), 4.40 (s, 4 H), 6.72 (s, 2 H), 7.09 (s, 2 H), 10.05 ppm (s, 1 H); ESI-TOFHRMS: m/z for C31H37N4O9 : calcd: 609.2555 [M] + , found: 609.2554. Fluorescence measurement: Fluorescence titration of the probe with CdCl2 was conducted with a solution of the probe (3 mL) in HEPES buffer (50 mm, pH 7.4)/MeOH (1:1). In the titration of the CdII complexes, 1 equiv of CdCl2 was added to an aqueous solution of the ligand before use. Fluorescence titration with Na2S was conducted with a solution of the CdII complex (3 mL) in HEPES buffer (20 mm, pH 7.4) at 37 8C. A freshly prepared aqueous stock solution of Na2S was added to the solution with a micropipette, and the fluorescence emission spectra were recorded after 5 min. In the case of titration with the thiol species (Figure 3 B), ROS, and RNS (Figure S8), the fluorescence spectra were recorded 30 min after addition of the analytes. Relative fluorescence quantum yields (F) of pyronine derivatives 6 and 7 were determined by in a solution of rhodamine B (F = 0.65) in EtOH as standard. Cell culture: HeLa cells were cultured in high-glucose Dulbecco’s Modified Eagle’s Medium (DMEM; Sigma) supplemented with 10 % fetal bovine serum (FBS; Gibco) and 1 % antibiotic-antimycotic solution (Gibco) at 37 8C under a humidified atmosphere of 5 % CO2 in air. HUVEC cells were cultured in Endothelial Cell Growth Medium (TaKaRa) supplemented with 2 % fetal calf medium, 0.4 % endothelial cell growth factor, epidermal growth factor (0.1 ng mL¢1), basic fibroblast growth factor (1 ng mL¢1), heparin (90 mg mL¢1), and hydrocortisone (1 mg mL¢1) at 37 8C under a humidified atmosphere of 5 % CO2 in air. A subculture was performed every 3–4 days from subconfluent (< 80 %) cultures by using a trypsin/EDTA solution. For the fluorescence bioimaging, cells (3.0 Õ 104 cells per well) were cultured for 2 days in a 35 mm glass-bottomed dish (Iwaki Scitech, Japan). Fluorescence imaging of exogenous H2S in HeLa cells: Fluorescence imaging was conducted with a confocal-laser scanning microscope equipped with a 60 Õ objective lens (A1Rsi, Nikon) or equipped with a 63 Õ objective lens (LSM 780, Zeiss). The following detection channels were chosen for the ratiometric imaging; Ch1 lex = 562 nm, lem = 580–600 nm, and Ch2 lex = 562 nm, lem = 610– 630 nm (A1Rsi), and lex = 561 nm, lem = 600–630 nm (LSM 780). In a glass-based dish, HeLa cells in modified HEPES buffer solution (HBS; 107 mm NaCl, 6 mm KCl, 11.5 mm glucose, 20 mm HEPES, pH 7.4) were first incubated with pyrenebutylate (50 mm) for 2 min then with 6·CdII (30 mm) for 5 min. The cells were further incubated with CR8 (30 mm) at 37 8C for 20 min. After being washed twice
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Full Papers with HBS, the cells were treated with Na2S (150 mm) and subjected to the fluorescence imaging. Fluorescence imaging of hydrogen sulfide in HUVEC cells: HUVEC cells were grown to 80 % confluence, washed with DMEM, and incubated for 8–12 h. They were then washed with modified HBS and incubated with a solution of 6·CdII (30 mm), CR8 (30 mm), and pyrene butylate (50 mm) for 20 min at 37 8C. After exchange of the medium to HEPES (20 mm; pH 7.4, containing NaCl (107 mm), KCl (6 mm), glucose (11.5 mm), CaCl2 (2.0 mm), and MgCl2 (1.2 mm)), the cells were treated with VEGF (40 ng mL¢1 in 0.1 % BSA/H2O, Invitrogen) and then subjected the fluorescence imaging. For the inhibition of CSE activity, HUVEC cells were pretreated with PAG (dl-propargylglycine, 100 mm, Sigma) in the modified HBS buffer for 10 min before being incubated with 6·CdII, and PAG (100 mm) was added again 10 min before VEGF addition.
Acknowledgements We appreciate technical support from the Research Support Center, Graduate School of Medical Sciences, Kyushu University. This work was performed under the Cooperative Research Program of the “Network Joint Research Center for Materials and Devices” organized by (MEXT, Japan). A.O. acknowledges the Toray Science Foundation for financial support. I.T. acknowledges the Japan Society for the Promotion of Science (JSPS) Research Fellowships for Young Scientists. Keywords: analytical methods · cadmium · fluorescent probes · hydrogen sulfide · imaging agents · noncovalent interactions [1] a) C. Szabû, Nat. Rev. Drug Discovery 2007, 6, 917 – 935; b) T. Ida, T. Sawa, H. Ihara, Y. Tsuchiya, Y. Watanabe, Y. Kumagai, M. Suematsu, H. Motohashi, S. Fujii, T. Matsunaga, M. Yamamoto, K. Ono, N. O. DevarieBaez, M. Xian, J. M. Fukuto, T. Akaike, Proc. Natl. Acad. Sci. USA 2014, 111, 7606 – 7611. [2] a) G. Yang, L. Wu, B. Jiang, W. Yang, J. Qi, K. Cao, Q. Meng, A. K. Mustafa, W. Mu, S. Zhang, S. H. Snyder, R. Wang, Science 2008, 322, 587 – 590; b) W. Zhao, J. Zhang, Y. Lu, R. Wang, EMBO J. 2001, 20, 6008 – 6016. [3] M. Nishida, T. Sawa, N. Kitajima, K. Ono, H. Inoue, H. Ihara, H. Motohashi, M. Yamamoto, M. Suematsu, H. Kurose, A. van der Vliet, B. A. Freeman, T. Shibata, K. Uchida, Y. Kumagai, T. Akaike, Nat. Chem. Biol. 2012, 8, 714 – 724. [4] a) A. Kazuho, K. Hideo, J. Neurosci. 1996, 16, 1066 – 1071; b) H. Kimura, Biochem. Biophys. Res. Commun. 2000, 267, 129 – 133. [5] P. Kamoun, Amino Acids 2004, 26, 243 – 254. [6] V. S. Lin, C. J. Chang, Curr. Opin. Chem. Biol. 2012, 16, 595 – 601. [7] a) A. R. Lippert, E. J. New, C. J. Chang, J. Am. Chem. Soc. 2011, 133, 10078 – 10080; b) V. S. Lin, A. R. Lippert, C. J. Chang, Proc. Natl. Acad. Sci. USA 2013, 110, 7131 – 7135; c) H. Peng, Y. Cheng, C. Dai, A. L. King, B. L. Predmore, D. J. Lefer, B. Wang, Angew. Chem. Int. Ed. 2011, 50, 9672 – 9675; Angew. Chem. 2011, 123, 9846 – 9849.
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Manuscript received: May 17, 2015 Accepted article published: May 28, 2015 Final article published: June 26, 2015
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