An ICT-Based Hydrogen Sulfide Sensor with Good Water Solubility for Fluorescence Imaging in Living Cells

Full text of DOI:10.1007/s10895-015-1582-5

J Fluoresc
DOI 10.1007/s10895-015-1582-5
EDITORIAL
An ICT-Based Hydrogen Sulfide Sensor with Good Water
Solubility for Fluorescence Imaging in Living Cells
Qiuyan Xu1 & Longwei He1 & Haipeng Wei1 & Weiying Lin 1,2
Received: 19 March 2015 /Accepted: 4 May 2015
#
Springer Science+Business Media New York 2015
Introduction
Fluorescence imaging has been widely used as a powerful
tool for monitoring biomolecules within the context of living
systems with high spatial and temporal resolution [15–16].
Naphthalimide dye is the favorable building block for con-
structing fluorescent probes because of its excellent
photophysical properties, such as high extinction coefficients,
excellent quantum yields, and great photostability. In addition,
azido group is well-known for sensitively and selectively
responding to H2S, the electron withdrawing azide is very
easy reduced by H2S affording the electron donating amine
[17–26]. Thus, in this work, we employed the naphthalimide
chromophore as the signal reporter and azido group as the
responding site for H2S to construct a H2S probe based on
internal charge transfer (ICT) process (Scheme 1). In addition,
to enhance the water solubility, we introduced a hydrophilic
alcoholic group to the naphthalimide core. Prior to react with
H2S, we supposed that probe Nap-N3 has negligible fluores-
cence. However, interaction of naphthalimide azide with H2S
affords 4-amino-1,8-naphthalimide, which would elicit a sig-
nificant fluorescence enhancement via ICT process caused by
the electron donating amine and the electron withdrawing am-
ide. So, probe Nap-N3 might be employed as a turn-on fluo-
rescent probe for detecting H2S.
Hydrogen sulfide (H2S), traditionally considered to be a toxic
gas with the typical smell of rotten eggs and an antioxidant or
scavenger for reactive oxygen species (ROS) [1], has recently
emerged as a member of the endogenous gaseous transmitter
family of signaling molecules including nitric oxide (NO) and
carbon monoxide (CO) [2–3]. The endogenous H2S is pro-
duced by the enzymes (such as cystathionine β-synthase, cys-
t a t hi o n i n e γ - l ya s e, an d 3 - m e r ca pt op yr uva t e
sulphurtransferase) catalysis action of cysteine or cysteine de-
rivatives in mammalian tissues [4–6]. H2S plays an important
physiological role in many biological processes, for instance,
ischemia reperfusion injury, vasodilation, apoptosis, insulin
signaling, and oxygen sensing [7–11]. However, abnormal
levels of H2S is associated with various diseases, like
Alzheimer’s disease [12], Down’s syndrome [13], diabetes
[10], and liver cirrhosis [14]. Therefore, it’s very significant
for monitoring H2S using sensitive, selective, and water-
soluble fluorescent probe in the native biological
environment.
Electronic supplementary material The online version of this article
(doi:10.1007/s10895-015-1582-5) contains supplementary material,
which is available to authorized users.
Experimental
* Weiying Lin
weiyinglin2013@163.com
Materials and Instruments
1
State Key Laboratory of Chemo/Biosensing and Chemometrics,
Unless otherwise stated, all reagents were purchased from
commercial suppliers and used without further purification.
Solvents used were purified by standard methods prior to
use. Twice-distilled water was used throughout all experi-
ments; High resolution mass spectrometric (HRMS) analyses
were measured on a Finnigan MAT 95 XP spectrometer;
College of Chemistry and Chemical Engineering, Hunan University,
Changsha, Hunan 410082, China
2
Institute of Fluorescent Probes for Biological Imaging, School of
Chemistry and Chemical Engineering, School of Biological Science
and Technology, University of Jinan, Jinan, Shandong 250022,
People’s Republic of China

J Fluoresc
then extracted three times with dichloromethane. The organic
phase was collected, washed with brine, and dried with anhy-
drous MgSO4. The solvent was removed under reduced pres-
sure and the solid residue was purified by flash chromatogra-
phy column using methanol/dichloromethane (v/v 1:20) to
afford a yellow solid as compound Nap-N3 (10 mg, yield
70.9 %). 1H NMR (400 MHz, CDCl3) δ 8.64 (d, J = 7.2 Hz,
1H), 8.58 (d, J = 8.0 Hz, 1H), 8.45 (d, J = 8.4 Hz, 1H), 7.75 (t,
J = 7.8 Hz, 1H), 7.47 (d, J = 8.0 Hz, 1H), 4.45 (t, J = 5.2 Hz,
2H), 3.96 (t, J = 5.2 Hz, 2H).13C NMR (100 MHz, CDCl3) δ
164.87, 164.50, 143.84, 132.55, 132.06, 129.13, 126.91,
124.32, 122.29, 118.52, 114.72, 61.85, 42.80. MS (EI) m/z
282.1 [M]+. HRMS (EI) m/z calcd for C14H10N4O3 [M]+:
282.0747. Found 282.0758.
Scheme 1 A turn-on fluorescent H2S probe based on the proposed ICT
switching mechanism
NMR spectra were recorded on an INOVA-400 spectrometer,
using TMS as an internal standard; Electronic absorption
spectra were obtained on a LabTech UV Power spectrometer;
Photoluminescent spectra were recorded with a HITACHI
F4600 fluorescence spectrophotometer with a 1 cm standard
quartz cell; The fluorescence imaging of cells was performed
with OLYMPUS FV1000 (TY1318) confocal microscopy;
The pH measurements were carried out on a Mettler-Toledo
Delta 320 pH meter; TLC analysis was performed on silica gel
plates and column chromatography was conducted over silica
gel (mesh 200–300), both of which were obtained from the
Qingdao Ocean Chemicals.
Synthesis of Compound Nap-NH2
Sodium sulfide (20 mg, 0.25 mmol) was added to the
solution of compound Nap-N3 (14 mg, 0.05 mmol) in
3 mL DMF/H2O (v/v 9:1) mix-solvent. The mixture was
stirred for 1.5 h at room temperature in the dark. The
reaction mixture was poured into 100 mL of water and
then extracted two times with dichloromethane and ethyl
acetate (v/v 5:1). The organic phase was collected,
washed with brine, and dried with anhydrous MgSO4.
The solvent was removed under reduced pressure and
the solid residue was purified by flash chromatography
column using methanol/dichloromethane (v/v 1:10) to
afford a yellow solid as compound Nap-NH2 (9.5 mg,
Synthesis
The target compound Nap-N3 was readily synthesized in just
two steps as shown in Scheme 2.
1
yield 74.2 %). H NMR (400 MHz, DMSO) δ 8.69 (d,
Synthesis of Compound Nap-N3
J = 8.4 Hz, 1H), 8.42 (d, J = 7.2 Hz, 1H), 8.19 (d,
J = 8.4 Hz, 1H), 7.65 (t, J = 8.2 Hz, 1H), 7.56 (s, 2H),
6.89 (d, J = 8.4 Hz, 1H), 4.13 (t, J = 6.6 Hz, 2H), 3.59
(m, 2H).13C NMR (100 MHz, DMSO) δ 164.39,
163.49, 153.23, 134.35, 131.42, 130.18, 129.92,
124.35, 122.23, 119.79, 108.57, 107.91, 58.42,
41.78.MS (EI) m/z 256.1 [M]+. HRMS (EI) m/z calcd
for C14H12N2O3 [M]+: 256.0842. Found 256.0844.
The mixture of compounds 4-bromo-1,8-naphthalic anhy-
dride (82.8 mg, 0.3 mmol) and ethanolamine (90 mg,
1.5 mmol) in 5 mL of ethanol was heated at 80 °C under reflux
for 4 h. After cooling to room temperature, the reaction sol-
vent was removed under reduced pressure. The resulting res-
idue was a light yellow solid as the crude product of interme-
diate compound 2, which was directly carried on the next
reaction without further process. The mixture of compound
2 (16 mg, 0.05 mmol) and sodium azide (13 mg, 0.20 mmol)
in 3 mL of dry N,N-dimethylformide (DMF) was heated to
90 °C for 3 h in the dark. After cooling to room temperature,
the reaction mixture was poured into 100 mL of water and
Determination of the Fluorescence Quantum Yield
Fluorescence quantum yields for compounds Nap-N3 and
Nap-NH2 were determined by using fluorescein (Φf = 0.95
Scheme 2 Synthesis of H2S
sensor Nap-N3 and the reduced
product Nap-NH2

J Fluoresc
Fig. 1 a Fluorescence spectra (λex = 408 nm) of 10 μM Nap-N3 with 0–
3.5 eq. of Na2S in 25 mM phosphate buffer (pH 7.4, containing 2 %
ethanol). The inset shows the visual fluorescence color of Nap-N3
before (left) and after (right) addition of H2S (UV lamp, 365 nm). b
The linear relationship between the fluorescence intensity at 546 nm
and the concentration of H2S
in 0.1 M NaOH aqueous solution) as fluorescence standard
[27]. The quantum yields were calculated using the following
equation.
Results and Discussion
Fluorescence Responses to H2S
2
ΦFðXÞ ¼ ΦFðSÞðAS FX =AX FSÞðnX =nSÞ
With compound Nap-N3 in hand, we first evaluated the capa-
bility of Nap-N3 to detect H2S in aqueous buffer. The titration
of H2S to the probe Nap-N3 (10.0 μM) was performed in
25 mM PBS buffer (pH 7.4) with just 2 % ethanol. As de-
signed, upon excitation at 438 nm, the free sensor displayed
faint fluorescence at around 546 nm (Fig. 1a). However, ad-
dition of Na2S (a standard source for hydrogen sulfide) elicit-
ed a significant emission enhancement, suggesting that Nap-
N3 was reduced affording Nap-NH2 in the presence of H2S
and ICT process occurred. The fluorescence intensities at 546
exhibited 7.9-fold increase, it’s the ICT resonance structure of
compound Nap-NH2 that contributes mainly to the enhanced
fluorescence. The quantum yields of compounds Nap-N3 and
Nap-NH2 were determined 0.0056 and 0.1175, respectively,
using a reference fluorescein dye (with a quantum yield of
0.95 in 0.1 M NaOH aqueous solution). Importantly, the
Where ΦF is the fluorescence quantum yield, A is the ab-
sorbance at the excitation wavelength, F is the area in the
corrected emission spectrum, and n is the refractive index of
the solvent used. Subscripts S and X refer to the standard and to
the unknown, respectively.
HeLa Cells Culture
HeLa cells were cultured in DMEM (Dulbecco’s modified
Eagle’s medium) supplemented with 10 % FBS (fetal bovine
serum) in an atmosphere of 5 % CO2 and 95 % air at 37 °C.
Imaging of Exogenous H2S in Living Cells
HeLa cells were incubated with 5.0 μM Nap-N3 for 20 min in
an atmosphere of 5 % CO2 and 95 % air, and then treated with
20 μM Na2S for 10 min. Subsequently, the cells were imaged
using OLYMPUS FV1000 (TY1318) confocal microscope
with an excitation filter of 405 nm and emission channels of
520–570 nm (green channel).
Imaging of Endogenous H2S in Living Cells
PC-3 cells were incubated with 200 μM cysteine for 60 min in
an atmosphere of 5 % CO2 and 95 % air, and then treated with
5.0 μM Nap-N3 for 20 min. Subsequently, the cells were
imaged using OLYMPUS FV1000 (TY1318) confocal micro-
scope with an excitation filter of 405 nm and emission chan-
nels of 520–560 nm (green channel).
Fig. 2 Absorption spectra of probe Nap-N3 with 0–3.5 eq. of Na2S in
25 mM phosphate buffer (pH 7.4, containing 2 % ethanol)

J Fluoresc
Fig. 3 The comparison of the
partial 1H NMR spectra of a Nap-
N3 in CD3Cl and b Nap-NH2 in
d6-DMSO
chemodosimeter shows an excellent linear relationship be-
tween the fluorescent emission intensities at 546 nm and the
concentrations of H2S from 0.2 to 3.0 equivalent (Fig. 1b),
suggesting that the chemodosimeter is potentially useful for
quantitative determination of H2S. The ratio changes were
observed in absorption spectra, as shown in Fig. 2, upon ad-
dition of H2S, the absorption of naphthalimide azide at around
376 nm gradually faded and simultaneously a new red-shifted
(62 nm) absorption band at around 438 nm (characteristic
absorption of amino naphthalimide) was enhanced. There is
Fig. 4 Interfacial plots of the
orbitals for Nap-N3 (left) and
Nap-NH2 (right) in the excited
states

J Fluoresc
a clear isosbestic point (at 407 nm) was appeared, suggesting
that the conversion from Nap-N3 to Nap-NH2 is a concerted
process.
Mechanism Studies
To shed light on the H2S-triggered fluorescence turn-on re-
sponse, we decided to characterize the reduced product. Incu-
bation of Nap-N3 with Na2S afforded the product, which was
isolated and characterized by standard NMR and mass spec-
1
trometry (Figs. S1–3). The HNMR spectra comparison of
probe Nap-N3 and the product Nap-NH2, the peak of NH2
in Nap-NH2 was obviously observed, which suggested the
resulting product should be reduced compound Nap-NH2
(Fig. 3). We further employed time-dependent density func-
tion (TD-DFT) to calculate the molecular orbital plots of Nap-
N3 and Nap-NH2 by a suite of Gaussian 03 programs (6-
31G(d) basis sets). As shown in Fig. 4, for the probe Nap-
N3, both the highest occupied molecular orbital (HOMO) and
lowest unoccupied molecular orbital (LUMO) in the excited
states are distributed on the naphthalimide backbone and azi-
do group, however, the LUMO +1 is mainly located on the
electron-withdrawing azido group. The theoretical data sug-
gested the π electrons do not transfer from the azido unit to the
naphthalimide moiety and ICT process could not occur in
Nap-N3. By contrast, in the case of Nap-NH2, the LUMO
+1 is primarily resided on the naphthalimide backbone, but
not on the electron-donating amino group, which indicated the
π electrons transfer to the naphthalimide moiety and ICT pro-
cess occurs. These experimental facts and theoretical data
supported our proposed ICT switching mechanism.
Fig. 6 Time-dependent fluorescence intensity (at 546 nm) responses of
sensor Nap-N3 (10 μM) to 3 eq.Na2S in PBS buffer
Selectivity Studies
The detection limit was calculated to be 1.09 × 10−6
M
(S/N = 3), which locates in the range of physiological concen-
tration of H2S in vivo, indicating that the probe is sensitive to
H2S and might be suitable for detecting endogenous H2S in
biological samples. To examine the selectivity, the probe Nap-
N3 (10 μM) was treated with various biologically relevant
species in the aqueous buffer, such as the representative an-
ions, reactive oxygen species, reducing agents, small-
molecule thiols, and Na2S. As shown in Fig. 5, addition of
the representative interfering species including F−, Cl−, I−,
N3 , CO32−, SO42−, HPO42−, NO2 , NO3 , AcO−, SCN−, cit-
rate at 1 mM, and S2O32−, SO32−, ClO−, H2O2, NO, ascorbic
acid at 200 μM induced negligible changes. Notably, small-
molecule biothiols such as glutathione (GSH) and cysteine at
1 mM triggered only a small emission intensity increase.
−
−
−
Fig. 5 Fluorescence intensity (at 546 nm) of sensor Nap-N3 (10 μM) in
the presence of various analytes in aqueous solution (pH 7.4 PBS,
containing 2 % ethanol): 1, blank; 2, F−; 3, Cl−; 4, I−; 5, N3−; 6, CO3
;
2−
7, SO42−; 8, HPO42−; 9, NO2−; 10, NO3−; 11, Ac−; 12, SCN−; 13, citrate;
14, S2SO32−; 15, ClO−; 16, H2O2; 17, SO32−; 18, NO; 19, ascorbic acid;
20, Cys; 21, GSH; 22, Na2S
Fig. 7 The pH influence on the fluorescence intensity (at 546 nm) of
Nap-N3 (10 μM) in the absence (black square) or presence (red circle) of
Na2S (30 μM)

J Fluoresc
Fig. 8 Confocal fluorescence images of HeLa cells incubated with 5 μM Nap-N3 for 20 min (a, b) and then 20 μM Na2S for 20 min (c, d). Images (a, c)
were acquired using 405 nm excitation and green emission channels of 520–570 nm; (b, d) bright field images. Scale bar = 40 μm
However, just upon addition of 30 μM Na2S could elicit ob-
vious increase of emission intensity.
Fig. 8, the cells incubated with only Nap-N3 exhibited faint
fluorescence in the green channel. However, after co-
incubation with Na2S, strong green fluorescence was ob-
served in living cells, suggesting Nap-N3 was reduced by
H2S affording fluorescent compound Nap-NH2. Thus, these
results revealed that Nap-N3 is cell membrane permeable and
capable of monitoring H2S in living cells.
In addition to detection of extraneous H2S, we further de-
tected the biosynthesis H2S inside the cells. It’s well-known
that cystathionine β-synthase (CBS) and cystathionine γ-
lyase (CSE) could catalyze cysteine for producing H2S in
living cells [28–29]. HeLa cells were firstly incubated with
200 μM cysteine for 1 h, after rinsed three times by PBS
buffer, followed addition of 5 μM Nap-N3 and incubated an-
other 20 min. As exhibited in Fig. 9, in comparison with the
cells loaded with only the probe, the cells co-incubated with
cysteine and Nap-N3 elicited a marked increase of fluores-
cence intensity in the green channel. These results further
indicated that Nap-N3 is capable of detecting not only external
H2S in living cells, but also endogenous H2S biologically
produced by the cells.
Reaction-Time
In addition, the fluorescence intensity reached its maximum at
about 15 min (Fig. 6). The results suggested the probe Nap-N3
has a high selectivity for H2S over other biological species.
pH Effect
What’s more, probe Nap-N3 could respond well to H2S at
round physiological pH (Fig. 7). The results indicated Nap-
N3 may be suitable for studies of H2S in the living systems.
Fluorescence Imaging in Living Cells
Firstly, to examine the cell membrane permeability of
naphthalimide dye, we employed compound Nap-NH2 for
imaging living cells. The cells incubated with Nap-NH2 ex-
hibited strong fluorescence, while control cells showed none
fluorescence (Fig. S4), suggesting the naphthalimide dye is
cell membrane permeable. Then to examine the utility of the
sensor, we intended to image H2S in living cells. For proof-of-
concept, Nap-N3 (5 μM) was initially incubated with HeLa
cells for 20 min, after rinsed three times by PBS buffer, then
treated with 20 μM Na2S for another 20 min. As shown in
Conclusions
In summary, we have introduced a high water-soluble H2S
fluorescent probe based on naphthalimide chromophore. The
Fig. 9 Confocal fluorescence images of HeLa cells incubated with 5 μM
Nap-N3 only for 20 min (a, b) and 200 μM cystine for 1 h followed by
5 μM Nap-N3 for 20 min (c, d). Images (a, c) were acquired using 405 nm
excitation and green emission channels of 520–570 nm; (b, d) bright field
images. Scale bar = 40 μm

J Fluoresc
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Acknowledgments This work was financially supported by the
National Science Foundation of China (Nos. 21172063,
21472067) and the startup fund of University of Jinan.
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