In situ visualization and detection of protein sulfenylation responses in living cells through a dimedone-based fluorescent probe

Article abstract of DOI:10.1039/c3ob41434e

Sulfenylation is one of the reversible post-translational modifications, playing significant roles in cellular redox homeostasis and signaling systems. Herein, small fluorescent probe (CPD and CPDDM) based live-cell labelling technology for the visualization of protein sulfenylation responses in living cells has been developed. This approach enables the detection of protein sulfenylation without the need for cell lysis, fixation or purification, and permits the noninvasive study of protein sulfenylation in live cells through the direct fluorescent readout. This technology also can realize dynamic tracking of protein sulfenylation in situ with minimal perturbation to sulfenylated proteins and less interference with cellular function. Information on the global distribution and dynamic changes of endogenous protein sulfenylation has been obtained.

Full text of DOI:10.1039/c3ob41434e

Organic &
Biomolecular Chemistry
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In situ visualization and detection of protein
sulfenylation responses in living cells through a
Cite this: Org. Biomol. Chem., 2013, 11,
566
7
dimedone-based fluorescent probe†
a
a,b
a
a
a
Qin Yin,‡ Chusen Huang,‡ Chao Zhang, Weiping Zhu,* Yufang Xu,*
a
a
Xuhong Qian and Yi Yang*
Sulfenylation is one of the reversible post-translational modifications, playing significant roles in cellular
redox homeostasis and signaling systems. Herein, small fluorescent probe (CPD and CPDDM) based live-
cell labelling technology for the visualization of protein sulfenylation responses in living cells has been
developed. This approach enables the detection of protein sulfenylation without the need for cell lysis,
fixation or purification, and permits the noninvasive study of protein sulfenylation in live cells through
the direct fluorescent readout. This technology also can realize dynamic tracking of protein sulfenylation
in situ with minimal perturbation to sulfenylated proteins and less interference with cellular function.
Information on the global distribution and dynamic changes of endogenous protein sulfenylation has
been obtained.
Received 12th July 2013,
Accepted 17th September 2013
DOI: 10.1039/c3ob41434e
www.rsc.org/obc
reduced to thiol quickly, or undergoes reaction with a nearby
thiol to form a disulfide, enzymatically or nonenzymatically.
Introduction
8
The homeostasis of the cellular redox environment is one of In the presence of strong oxidants, sulfenic acid would be
the most important foundations of living systems. Among the further converted to sulfinic (Cys–SO H) and/or sulfonic (Cys–
2
9
10
important factors associated with redox homeostasis and SO H) acid derivatives, which are barely reversible. There-
3
signaling, protein thiol is especially attractive for its direct fore, sulfenic acid, usually the first product of thiol oxidation,
involvement in many biological processes through post-trans- acts as an active intermediate in the complex thiol redox
1
4,8,11
lational modification. Sulfenylation, i.e. reversible oxidation network.
of protein thiols into sulfenic acid (R-SOH), has received tre- Due to the significant role and high reactivity of protein
2
mendous attention in recent years. It was initially found to sulfenic acid, a variety of approaches have been developed for
regulate the activity of an important metabolic enzyme, glycer- detecting sulfenic acid in proteins. By monitoring the differ-
3
aldehyde 3-phosphate dehydrogenase (GAPDH), in 1969. It ence in UV-vis absorption spectra between the adduct of
has been proven now that sulfenylation is one of the most 7-chloro-4-nitrobenzo-2-oxa-1,3-diazol (NBD-Cl) with sulfenic
direct and reversible post-translational modifications in the acid and thiol, the sulfenylation of purified protein was able to
4
12
cell antioxidant defense system, and has significant roles as be detected in vitro. The C-terminal cysteine rich domain of
3
the intermediate during enzyme catalytic cycles, the regulator the yeast Yap1 transcription factor was engineered to trap the
of protein translocation, the modulator of gene expression
and the global mechanism for cell signaling. Protein sulfenic and versatile chemical approaches. A method with sodium
acid is a highly reactive intermediate because it can be arsenite was used to reveal the proteomic sulfenylation
5
6
2
sulfenic acid. More effort was directed towards the specific
7
13
1
4
response of cells towards hydrogen peroxide. The dimedone
5,5-dimethyl-1,3-cyclohexanedione) and its derivatives were
the popular ligands for their specific interaction with sulfenic
(
a
State Key Laboratory of Bioreactor Engineering, Shanghai Key Laboratory of
Chemical Biology, School of Pharmacy, East China University of Science and
Technology, 130 Meilong Road, Shanghai 200237, PR China.
E-mail: wpzhu@ecust.edu.cn, yfxu@ecust.edu.cn, yiyang@ecsut.edu.cn;
Fax: (+86) 21-64252603
1
5
8,16
acid. Some mass spectrometry
and immunochemical
2
,17
approaches
have been introduced to monitor the formation
of sulfenic acids in proteins or even the in situ imaging of
protein sulfenic acid distribution in fixed cells based on dime-
done. The small-molecule probes have also been synthesized
b
Department of Chemistry, Life and Environmental Science College, Shanghai
Normal University, 100 Guilin Road, Shanghai 200234, PR China
Electronic supplementary information (ESI)
0.1039/c3ob41434e
†
1
‡
available. See DOI:
1
8
2
through the incorporation of biotin, fluorophore, isotope-
2
,19
2,20
These authors contributed equally to this work.
coded affinity tags
or series of azide-based tags
into
7
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Scheme 1 Structures of CPD and CPDDM.
Fig. 1 Plausible mechanism for the specific detection of sulfenic acid moieties
in proteins with CPD.
dimedone-like derivatives, as well as the 1,3-cyclopentane-
2
1
22
dione and β-ketoesters, which can selectively label sulfenic
acid in protein and make proteomic analysis of protein sulfe-
nylation possible in biological media. Despite this progress,
as sulfenic acid is a highly reactive intermediate, some time-
consuming procedures with cell lysis, fixation or purification
might make the data interpretation complicated and difficult.
Furthermore, to investigate the dynamic and compartmentali-
zation of cellular protein in different thiol oxidation states,
these kinds of thiol should be specifically labeled and imaged,
As the sulfenic acid is an unstable transient intermediate
during the oxidation processes in living cells, it should be
detected rapidly and non-invasively with a small molecular
fluorescent probe with low steric bulkiness and fast labeling
2
6
kinetics, which can track and visualize the protein sulfenyla-
tion directly. The sulfenic acid reacts with dimedone and its
2
,15
derivatives to form a stable thioether derivative,
whereas
free thiol has no reactivity towards dimedone under the physio-
logical conditions (Fig. 1). Thus it is possible to selectively
discriminate sulfenic acid from other forms of thiols in pro-
2
3
ideally in living cells. We previously developed in situ chemi-
cal staining methods for S-nitrosothiols and disulfides in fixed
cells, imaged their intracellular distribution and observed the
central role of mitochondria produced oxygen species in the
2
teins. In this research, the fluorescent probe CPD was
designed and synthesized through the conjugation of dime-
done analogue with a 7-aminocoumarin fluorophore, which
had a suitable excitation wavelength, stable fluorescence
2
4
oxidative modification of thiols. Hence a noninvasive and
versatile approach with in situ labeling and imaging in living
cells is still an urgent need for investigating the dynamic and
compartmentalization of cellular protein sulfenylation.
In this study, we reported the visualization of protein sulfe-
nylation in living cells using a dimedone-based fluorescent
probe (CPD, Scheme 1). This small fluorescent probe based live-
cell-labeling technology ensured more rapid detection of
protein sulfenylation through direct fluorescent readout, avoid-
ing the time consuming nature of cell lysis, fixation or purifi-
cation. Additionally, CPD with low concentration (5 μM) used in
labeling, minimized perturbation to sulfenylated proteins and
less interference with cellular ROS. Therefore, this noninvasive
labeling protocol enabled us to visualize cellular sulfenylated
proteins in the native state aimed at studying their global distri-
bution and dynamic changes.
2
7
signal and moderate quantum yield (Φ ) under the physio-
F
logical conditions (Table 1). In order to tune the biocompat-
ibility of CPD, the piperazine and urethane group (Scheme 1)
was introduced as the linker. A control compound (CPDDM,
Scheme 1) with dimethylated dimedone to block its reactivity
towards sulfenic acids was synthesized. The partition coeffi-
cient (P) assay showed that the log P value of CPD was 1.49 ±
0.16, and that of CPDDM was 1.95 ± 0.11 (Table 1), indicating
that they were cell-permeable probes and soluble in PBS buffer
(
pH 7.4, 1% DMSO as the co-solvent).
We then investigated the spectroscopic characteristics of
CPD and CPDDM. The UV absorption and fluorescence inten-
sity of CPD displayed negligible changes at pH 7.0–10 (ESI
Fig. 1a and b†). The UV absorption and fluorescence spectra of
CPDDM were almost identical to those of CPD in PBS buffer
(
Fig. 2a). Because protein sulfenylation is related to the redox
environment of living cells, we next investigated the effect of
redox agents on the fluorescence intensity of CPD. As shown
Results and discussion
To visualize protein sulfenylation in living cells, the probe in Fig. 2b, little fluorescence changes were observed in the
2
5
must be selective, stable, water-soluble and cell permeable.
presence of different concentrations of dithiothreitol (DTT)
a
Table 1 Spectral properties of CPD and CPDDM in PBS
ε (M− cm−1
1
)
Φ
Log P
Solvent
PBS
Compounds
λ
ex, max (nm)
λ
em, max (nm)
Stokes shift
F
CPD
CPDDM
415 ± 2
416 ± 2
485 ± 3
483 ± 3
70
67
24 800
23 000
0.11
0.10
1.49 ± 0.16
1.95 ± 0.11
a
F
N-Butyl-4-butylamino-1,8-naphthalimide was taken as the standard for Φ .
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Fig. 2 The spectroscopic characteristics of CPD and CPDDM. (a) Normalized
absorption and fluorescence of CPD and CPDDM in PBS buffer. (b) Fluorescence
responses of 5 μM CPD to different concentrations of redox regulating reagents
2 2
DTT and H O (0–5 mM).
Fig. 4 Selectivity of CPD as shown by SDS-PAGE. The labeling of proteins by
CPD occurred only in the presence of hydrogen peroxide or N,N,N’,N’-tetra-
2 2
methylazodicarboxamide (TMAD). DTT: dithiothreitol, Dim: dimedone, H O :
hydrogen peroxide, Dia: TMAD. “+”: the molecule was present in the labeling
system; “−”: the molecule was not present in the labeling system.
in living cells. In order to investigate whether the diethylamino-
coumarin fluorophore and piperazine alkyl ether linker unit
induce a significant bias in probe localization and protein
labeling, the CPD was introduced into the labeling system
before washing. Both CPD and CPDDM penetrated into Chang
liver cells, and showed a similar and wide pattern in cells (ESI
Fig. 3 CPD do not interfere with cell viability and the intracellular ROS level. (a)
Cell viability test of CPD. (b) Effect of CPD on cellular ROS detected by 5 μM
H DCFDA.
2
Fig. 4a and c†). This result suggested that CPD had no bias.
After the cells were washed to remove free probes, the CPDDM
and hydrogen peroxide (H
2
O
2
), which confirmed that CPD labeled cells had a very weak signal left (ESI Fig. 4c,d and
exhibited stable fluorescence characteristics in various redox Fig. 5b,f†). In contrast, marked fluorescence signal was still
environments. observed in cells stained with CPD (ESI Fig. 4a,b and Fig. 5a,
Next, cell cytotoxicity experiments were conducted. CPD f†), which further confirmed that CPD has no bias and the
showed minimal cytotoxicity as shown by the CellTiter 96 labeled fluorescence signal was mainly from endogenous sulfe-
AQueous Non-Radioactive Cell Proliferation Assay (Promega) nic acids in proteins. We then tracked the dynamic changes of
2
2,28
(Fig. 3a). As was reported in the literature,
high concen- protein sulfenic acids by using CPD based live-cell-labeling
tration of dimedone (millimolar level) perturbed the cellular technology. The fluorescence intensity of CPD-labeled cellular
redox equilibrium, which may be due to the formation of sulfenylated protein decreased notably with increasing
2
9
4
-peroxydimedone radical. In contrast, CPD in the micro- amounts of dimedone (Fig. 5c and g), owing to the competitive
molar range showed a negligible effect on the intracellular binding of dimedone with the de novo formed sulfenic acid in
ROS level and cellular redox balance (Fig. 3b). This was also the cells. Co-incubation of the cells with DTT, which could
2
2
consistent with reported results. In subsequent experiments, reduce the sulfenic acids to thiols, also decreased the fluore-
μM CPD was used for live cell labeling. scent staining signal by CPD (Fig. 5d, h and i). On the other
Then the specific assay of CPD for sulfenic acid was also hand, oxidants such as and TMAD significantly
5
2 2
H O
tested. We used a mono-thiol mutant of thioredoxin (Trx-M, enhanced CPD staining in a dose-dependent manner, by oxi-
human thioredoxin-2 with C69G mutation, the detailed amino dizing free thiols and increasing the cellular protein sulfenyla-
acid sequence in ESI†) as a model protein for in vitro sulfenyla- tion level (Fig. 5e, j, k, l and m). In these cases, the newly
tion labeling. The electrophoresis result demonstrated that formed sulfenic acid was trapped and labeled by CPD upon
CPD could label sulfenylated protein selectively in the pres- oxidation (Fig. 5k and m). The increased CPD staining under
ence of hydrogen peroxide or TMAD, both of which were able oxidative conditions disappeared when dimedone was present,
to oxidize thiols to sulfenic acids (Fig. 4). Interestingly, CPD due to the competitive binding of sulfenic acid by CPD and
did not stain the Trx-M dimer, which was also formed under dimedone (Fig. 5j). We also evaluated protein sulfenylation in
these oxidative conditions and contained intersubunit di- HL60 cells, the human promyelocytic leukemia cells in suspen-
sulfide bonds. These in vitro labeling data confirmed the sion (ESI Fig. 5†). It was interesting that TMAD was much
selectivity of CPD to sulfenic acid in proteins.
more efficient than H O to promote sulfenic acid formation
2 2
Imaging of protein sulfenic acid with CPD was tested in in the HL60 cells, while these two oxidants induced similar
Chang liver cells. Marked fluorescence signal was observed in sulfenylation levels in Chang liver cells, suggesting different
cells stained with CPD (Fig. 5a), but none for cells stained with redox environments in different cell lines. All these results
cell permeable CPDDM (Fig. 5b and 5f), suggesting the demonstrated that the cell-permeable CPD could be used for
specific labeling of endogenous sulfenylated proteins by CPD imaging and tracking endogenous sulfenylated proteins by the
7
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Fig. 5 Live cell imaging of protein sulfenylation in Chang liver cells responding to redox regulations. (a–e) Fluorescent images of cells labeled with CPD (5 μM, a),
CPDDM (5 μM, b), the coumarin fluorophore, CPD (5 μM) co-labeled with 1 mM dimedone (c), CPD (5 μM) co-labeled with 2 mM DTT (d), and CPD (5 μM) labeled
in the presence of 1 mM H O (e). Cellular responses of CPD labeled sulfenylated protein in Chang liver cells to redox regulation (f–m). (f) The fluorescence intensity
2 2
of CPD/CPDDM labeled cells. (g) Effect of different concentrations of dimedone (co-incubation). (h) Effect of different concentrations of DTT. (i) Kinetics of 2 mM
DTT reduction (co-incubation). ( j) Effect of different concentrations of TMAD on protein sulfenylation. (Black: co-incubated with CPD, red: co-incubated with CPD
and 1 mM dimedone). (k) Kinetics of 1 mM TMAD oxidation (co-incubation). (l) Effect of different concentrations of H
reduction (co-incubation). Error bars represent ±S.E.M. of six measurements.
2 2
O (co-incubation). (m) Kinetics of 1 mM
2 2
H O
variation of fluorescence intensity in living cells, and protein scattered yellow fluorescence (Fig. 6a). Under the oxidative con-
sulfenic acid in different cells had different responses toward ditions, the signal of CPD staining turned out to be much
oxidative conditions.
brighter in the focused eccentric perinuclear zone where mito-
We next investigated the localization of endogenous protein chondria were located (Fig. 6b) with no obvious fluorescent
sulfenic acids by confocal microscopy and co-staining with change of the network pattern (Fig. 6d), showing the increase
organelle-specific labels, such as the Mito-Tracker Deep Red of sulfenylated proteins within mitochondria during the oxi-
staining for mitochondria and DsRed-2 fluorescent protein tar- dation. These results suggested that the protein sulfenylation
geted to ER. The fluorescence signal of the CPD-labeled sulfe- occurred mainly in the ER and mitochondria, and the thiols in
nylated proteins showed mainly a smooth perinuclear and mitochondria proteins were significantly oxidized to sulfenic
peripheral pattern of a lace-like network, suggesting mainly acid under the oxidative stress.
the endoplasmic reticulum (ER) location (Fig. 5a and 6c).
In recent years, different detection methodologies to
There was a weak signal indicated by the punctuated pattern protein sulfenylation have been highlighted. Poole et al. syn-
2
in the perinuclear zone suggesting mitochondria staining thesized fluorescent or biotinylated derivatives of dimedone.
(Fig. 5a). The merged image of CPD and mitochondria showed Carroll et al. designed a series of azide-based analogs of
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2
dimedone specific antibodies, the immuno-fluorescence
signal was shown to be located in ER as well. Protein sulfenyla-
tion could happen spontaneously and nonenzymatically
during protein folding processes in aqueous solutions contain-
8
ing dissolved oxygen. As the main location for protein syn-
thesis and oxidative folding, ER maintains the most oxidative
2
9
status in the cell. Therefore, sulfenic acid could be an impor-
tant intermediate during protein disulfide bond formation in
ER. It was interesting that sulfenic acids were also found to be
enriched in mitochondria, particularly under oxidative con-
ditions. As the key organelle for respiration, mitochondria are
the major locations for many redox-related biochemical pro-
cesses. We previously reported that in mammalian cells, mito-
chondria maintained a substantial level of disulfide proteome
under resting conditions, which markedly increased upon oxi-
2
4
dative stimulation. Similarly, S-nitrosoproteins also mainly
existed in the mitochondria and perimitochondrial compart-
2
4
ment. Protein vicinal dithiols, the reduced end of the thiol
redox form, were also found to be concentrated in mitochon-
3
0
dria.
Furthermore, both S-nitrosothiol and disulfide
formation were determined by mitochondrial respiration and
the generation of reactive oxygen species. All these results
suggested that mitochondria were the central participants in
thiol redox regulation and might have profound effects on cel-
lular protein function.
Fig. 6 Co-localization of sulfenylated proteins with mitochondria and ER. (a)
Co-localization of sulfenylated proteins labeled with CPD with mitochondria
labeled with Mito-Tracker Deep Red. (b) Co-localization of sulfenylated proteins
Conclusions
In summary, we reported the in situ visualization and detec-
tion of protein sulfenylation responses in living cells through
trapped by CPD during 1 mM H
2 2
O co-incubation with mitochondria labeled
with Mito-Tracker Deep Red. (c) Co-localization of sulfenylated proteins labeled
with CPD with ER-targeted DsRed-2. (d) Co-localization of sulfenylated proteins the direct fluorescence readout with dimedone-based fluore-
trapped by CPD during 1 mM H
2
O
2
co-incubation with ER-targeted DsRed-2.
scent probes CPD. The control probe CPDDM labeled cells dis-
played no fluorescence signal and further demonstrated that
CPD could be applied for dynamic tracking of protein sulfeny-
2
,20
dimedone,
and combined the usage of isotope-coded lation without bias. By using this live-cell-labeling technology,
2
dimedone and iododimedone to quantify the sulfenic acid in we could track dynamic changes of protein sulfenylation and
proteins. Furthermore, they also developed an immunohisto- obtained more information about the subcellular distribution
2
,7
chemistry approach for differential analysis of samples with of protein sulfenic acid in situ, as well as its potential roles in
various protein sulfenylation levels by using specific anti- cellular function.
bodies that identify the epitope conjugated by protein sulfenic
acid and dimedone. Herein, we focused on the development of
new live-cell-labeling technology by using a small-molecule
fluorescent probe. This noninvasive method can be applied to
monitor the protein sulfenylation in living cells through direct
Experimental
Synthesis of CPD and CPDDM
readout of CPD fluorescent staining signal with minimal per-
turbation to sulfenylated proteins and less interference with chromene-3-carbonyl)piperazine-1-carboxylate (CPD). To a
cellular function.
solution of CSO1 (66 mg, 0.12 mmol) in THF–DCM (3/1, v/v,
4-(2,4-Dioxocyclohexyl)butyl-4-(7-(diethylamino)-2-oxo-2H-
By using this versatile live-cell-labeling technology, we 4 mL) was added 4 N HCl (4 mL) in an ice bath. The
found that the basal level of protein sulfenylation already mixture was stirred at room temperature for about 4 h until
existed in resting cells, suggesting that it was an important TLC shows that the material CSO1 disappeared. Then the reac-
physiological regulator of protein activity and the global signal- tion mixture was diluted with water (5 mL) and extracted with
7
ing mechanism, rather than just defense against ROS stress. DCM (3 × 15 mL). The organic phases were combined and
In resting cells, protein sulfenylation appeared to be abundant dried with anhydrous MgSO . The organic phase was reduced
4
in ER as demonstrated by CPD labeling of living cells. In a to dryness and the crude product was purified by column
recent work in which protein sulfenylation was detected by chromatography on silica gel (with DCM–EtOAc–MeOH,
7
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6 : 4 : 1) to obtain a yellow solid (43 mg, 70%). Melting point Preparation of reduced forms of Trx-M
Paper
1
1
(
mp) 130–135 °C. H NMR (CDCl , 400 MHz) δ 7.89 (s, 1H),
3
For the single thiol model of Trx-M, 10 mM DTT was used to
reduce the cysteine in the protein. After reduction, excess
reductant was removed by gel filtration on Sephadex G-25
against PBSE buffer (PBS buffer prepared by DPBS powder,
7
1
4
1
2
1
2
1
.27 (d, 1H, J = 9.2 Hz), 6.62 (dd, 1H, J = 8.8, 2.0 Hz), 6.49 (d,
H, J = 1.6 Hz), 4.14 (t, 2H, J = 6.4 Hz), 3.74 (br, 2H), 3.58 (br,
H), 3.48–3.39 (m, 8H), 2.75–1.90 (m, 6H), 1.72–1.40 (m, 5H),
1
3
3
.24 (t, 6H, J = 7.2 Hz); C NMR (CDCl , 100 MHz) δ 204.4,
5
mM EDTA, pH 7.4).
03.8, 165.3, 159.1, 157.3, 155.4, 151.8, 129.9, 115.7, 109.4,
07.7, 104.5, 96.9, 65.3, 58.2, 49.2, 44.9, 39.6, 29.6, 29.0, 28.8,
The status of the thiols in model proteins was verified
using 5,5′-dithiobis-(2-nitrobenzoic acid) (DTNB) after incubat-
ing proteins with an excess of DTNB in PBSE buffer for 30 min
in the dark at room temperature. An absorption coefficient at
−
1
4.5, 23.5, 12.4; IR (KBr, cm ): 3475, 2925, 2850, 1710, 1620,
595, 1518, 1456, 1412, 1354, 1231, 1190, 1125, 1077; HRMS
+
(ESI+) calcd for C H N O [M + H] 540.2710, found
−1
−1
2
9
38 3 7
4
12 nm of 14.12 mM cm was used to quantify the 5-thio-3-
5
R
40.2703; HPLC t = 25.20 min (purity >95%, monitored both
nitrobenzoate anion, with the absorbance of the DTNB solu-
tion and the intrinsic low absorbance of proteins at this wave-
length accounted for. The molar ratio of thiol quantified by
DTNB and of protein quantified by absorbance at 280 nm was
about 0.9–1.1.
at 254 and 415 nm).
-(3,3-Dimethyl-2,4-dioxocyclohexyl)butyl 4-(7-(diethylamino)-
-oxo-2H-chromene-3-carbonyl)piperazine-1-carboxylate
CPDDM). To a solution of CPD (20 mg, 0.04 mmol) in
acetone (4 mL) was added potassium carbonate (11 mg,
4
2
(
0
.08 mmol). After refluxing for 30 min, CH
3
I (14 mg,
SDS-PAGE and fluorescence imaging of gels
0
.09 mmol) was added, and then the reaction mixture was con-
The selectivity of CPD was verified by 15% SDS-PAGE electro-
phoresis. Samples were labeled in the PBS buffer at 37 °C for
tinued to reflux for about 12 h. TLC shows that the material
CPD disappeared, the reaction mixture was cooled to room
temperature and chloroform was added, left for 30 min, and
filtrated to get a clean solution. After removing the solvent
under reduced pressure, a yellow crude solid was obtained.
The crude product was purified by column chromatography on
silica gel (with DCM–MeOH, from 100 : 1 to 50 : 1) to obtain a
3
1
0 min, with a final concentration of protein at 50 μM, CPD at
00 μM and different concentrations of chemicals. After label-
ing, the samples were desalted using Bio-Rad Micro Bio-Spin
columns with Bio-Gel P-6 in SSC buffer (cat. no. 732-6201).
Then, the desalted samples were mixed with SDS-PAGE
loading buffer without β-mercaptoethanol. Protein concen-
trations were quantified by the Bradford method. Different
volumes of samples were loaded to make sure the same quan-
tity of proteins was loaded. Then, electrophoresis was started
immediately. The gel was imaged using the Carestream In Vivo
Imaging FX System (excitation: 420 nm, emission: 480 nm).
The same gel was also stained using Coomassie brilliant blue
pale yellow solid (13 mg, 62%). Melting point (mp)
1
1
1
2
3
55–160 °C. H NMR (CDCl
3
, 400 MHz) δ 7.91 (s, 1H), 7.36 (d,
H, J = 8.8 Hz), 6.63 (dd, 1H, J = 8.8, 2.4 Hz), 6.49 (d, 1H, J =
.4 Hz), 4.15 (t, 2H, J = 6.4 Hz), 3.76 (br, 2H), 3.59 (br, 4H),
.49–3.43 (m, 6H), 2.88–1.89 (m, 6H), 1.71–1.41 (m, 5H), 1.38
1
3
(s, 6H), 1.27–1.23 (m, 6H); C NMR (CDCl , 100 MHz) δ 210.8,
3
2
1
2
2
10.6, 171.3, 159.1, 157.3, 155.4, 151.9, 129.7, 115.9, 109.6,
08.2, 105.3, 97.1, 65.4, 61.0, 46.0, 45.1, 37.0, 29.7, 29.3, 27.2,
5.6, 24.1, 23.6, 22.6, 19.2, 14.1, 12.3; IR (KBr, cm− ): 3462,
(
CBB) after the fluorescent image was obtained.
1
Cell culture
928, 2856, 1700, 1625, 1599, 1517, 1407, 1345, 1247, 1140;
+
Chang liver and HL60 cells were obtained from the American
Type Culture collection. Chang liver/HL60 cells were grown in
DMEM (high glucose) medium/RPMI 1640 medium sup-
plemented with 10% FBS. Chang liver cells were typically pas-
saged with a sub-cultivation ratio of 1 : 4 every two days. HL60
cells were typically passaged with a sub-cultivation ratio of
HRMS (ESI+) calcd for C31
found 590.2842; HPLC t = 31.26 min (purity >95%, monitored
both at 254 and 415 nm).
H
41
N
3
O
7
Na [M + Na] 590.2842,
R
Expression and purification of thioredoxin 2 mutant (Trx)
Human thioredoxin 2 (signal peptide removed) gene with an
N-terminal hexahistidine tag was constructed on pET-28a
vector (Novogen). To obtain a single thioredoxin mutant,
Cys69 was mutated to glycine on pET-28a vector. Thioredoxin 2
C69G mutation (Trx-M) protein was expressed in Escherichia
coli strain BL21 (DE 3). After 4 hours of induction with 0.1 mM
1
: 10 every two days. Cells were incubated in a 5% CO
2
humidi-
fied incubator at 37 °C.
Detection of Chang liver cellular protein sulfenylation
response to different chemicals
IPTG, E. coli cells were harvested by centrifugation. Lysis was Cells were seeded on a GE 96-well glass matriplate (cat. no.
obtained by ultrasound, and then centrifuged at 10 000g for 28-9323-99) the day before detection with about 80% intensity.
3
0 min at 4 °C. Trx-M was purified from the supernatant using During detection, the cells were washed twice using DPBS
affinity chromatography on a Histrap HP column (GE Health- (containing Ca, Mg) buffer at 100 μL per well each time. Then,
care) according to the manufacturer’s instructions. The buffer the cells were co-incubated with 5 μM CPD and different con-
for storage was inter-changed with DPBS using an Amicon centrations of chemicals (100 μL per well in all) in DPBS (con-
Ultrafiltration device from Millipore. The purity of the protein taining 1% DMSO) for 30 min in a 5% CO
2
humidified
was verified by 15% SDS-PAGE to reach 95%. The purified incubator at 37 °C. After that, the cells were washed for six
protein was stored at −80 °C until further use.
times using PBS (containing Ca, Mg) buffer at 100 μL per well
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each time. Then 100 μL of PBS was added into each well for using a Promega CellTiter 96® AQueous Non-Radioactive Cell
detection. Then, the fluorescent intensity was quantified using Proliferation Assay Kit.
a Biotek Synergy 2 microplate reader with a 420 nm excitation
filter (bandpass: 10 nm) and a 485 nm emission filter (band-
pass: 20 nm).
H
2
DCFDA test
Cells were seeded on a GE 96 well glass matriplate (cat. no.
8-9323-99) the day before detection with about 80% intensity.
During detection, cells were labeled with different concen-
2
In situ imaging
Chang liver cells were labeled with 5 μM CPD in DPBS (con- trations of dimedone for 30 min as described before. Then
taining 1% DMSO) at 37 °C. For the H
, DTT, or dimedone 100 μL of H DCFDA (5 μM) in PBS solution was added to each
co-incubated sample, 1 mM H
, 2 mM DTT or 1 mM dime- well. After 30 min of incubation and two washes, the
2
O
2
2
2 2
O
done was introduced into the labeling system. For the CPDDM fluorescent intensity was measured using a Biotek Synergy
sample, the same concentration of CPDDM was used instead 2 Microplate Reader (ex: 485 nm, em: 528 nm (bandpass:
of CPD. After 30 min, the cells were washed six times to 20 nm)).
remove unbounded CPD before in situ imaging with a Nikon
A1R confocal laser scanning microscope using a Plan Apochro-
mat violet corrected (VC) 60× WI (N.A. 1.20; W.D. 0.27) objec-
tive, with excitation by a 405 nm laser, and 425–475 nm
Acknowledgements
We acknowledge financial support from the National Natural
Science Foundation of China (grants 21076077, 21236002 and
31071260), the National Basic Research Program of China
emission light was collected.
Colocalization
(
973 Program, 2013CB733700 and 2010CB126100), the
For CPD co-localization with Mito-Tracker sample: cells were
pre-washed twice, and labeled with 50 nM Mito-Tracker Deep
Red for 15 min at first and then with 5 μM CPD as described
above.
National High Technology Research and Development
Program of China (863 Program, 2012AA061601 and
2
011AA10A207), Shanghai Pujiang Program, Fok Ying Tung
Education Foundation (grant no. 111022), the Program for Pro-
fessor of Special Appointment (Eastern Scholar) at Shanghai
Institutions of Higher Learning and the Fundamental
Research Funds for the Central Universities, the Innovative
Program of Shanghai Normal University (SK201331), the
Science and Technology Innovation Foundation for College
Students (B-7062-12-001114).
For CPD co-localization with ER-tracker sample: pDsRed-2-
ER vector was transfected into Chang liver cells using a
Calcium Phosphate Cell Transfection Kit (Beyotime, cat. no.
C0508) the day before detection. Then, the cells were labeled
by 5 μM CPD as described above.
To exclude the interference with the fluorescent signal: the
excitation of CPD was at 405 nm, and 425–475 nm emission
light was collected. The excitation of DsRed-2 was at 561 nm,
and 570–620 nm emission light was collected. The excitation
of Mito-Tracker Deep Red was at 638 nm, and >650 nm emis-
sion light was collected.
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