Synthesis and antitumor potential of new arylidene ursolic acid derivatives via caspase-8 activation

Article abstract of DOI:10.1002/ardp.202000448

Continuing our studies on NO-donating ursolic acid–benzylidene derivatives as potential antitumor agents, we designed and synthesized a series of new arylidene derivatives containing NO-donating ursolic acid and aromatic heterocyclic units. Compounds 5c and 6c showed a significant broad-spectrum antitumor activity. Compound 5c exhibited nearly three- to nine-fold higher cytotoxicity as compared with the parent drug in A549, MCF-7, HepG-2, HT-29, and HeLa cells, and it was also found to be the most potent apoptosis inducer of MCF-7 cells. More importantly, compound 5c arrested the MCF-7 cell cycle in the G1 phase, which was associated with caspase activation and a decrease of the Bcl-2/Bax ratio. Meanwhile, compound 5c caused changes in morphological features, dissipation of the mitochondrial membrane potential, and accumulation of reactive oxygen species. A docking study revealed that the nitroxyethyl moiety of compound 5c may form hydrogen bonds with caspase-8 amino acid residues (SER256 and HIS255). Together, these data suggest that NO-donating ursolic acid–arylidene derivatives are potent apoptosis inducers in tumor cells.

Full text of DOI:10.1002/ardp.202000448

Received: 30 November 2020
DOI: 10.1002/ardp.202000448
Revised: 22 January 2021
Accepted: 29 January 2021
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F U L L P A P E R
Synthesis and antitumor potential of new arylidene ursolic
acid derivatives via caspase‐8 activation
Baoen He1 | Zuchang Zhu1 | Fenglian Chen1 | Rong Zhang1
Weiqiang Chen2 | Te Zhang3 | Tao Wang1 | Jiamei Lei1
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1School of Pharmaceutical Sciences,
Guangzhou University of Chinese Medicine,
Guangzhou, China
Abstract
Continuing our studies on NO‐donating ursolic acid–benzylidene derivatives as
potential antitumor agents, we designed and synthesized a series of new arylidene
derivatives containing NO‐donating ursolic acid and aromatic heterocyclic units.
Compounds 5c and 6c showed a significant broad‐spectrum antitumor activity.
Compound 5c exhibited nearly three‐ to nine‐fold higher cytotoxicity as compared
with the parent drug in A549, MCF‐7, HepG‐2, HT‐29, and HeLa cells, and it was
also found to be the most potent apoptosis inducer of MCF‐7 cells. More im-
portantly, compound 5c arrested the MCF‐7 cell cycle in the G1 phase, which was
associated with caspase activation and a decrease of the Bcl‐2/Bax ratio. Mean-
while, compound 5c caused changes in morphological features, dissipation of the
mitochondrial membrane potential, and accumulation of reactive oxygen species. A
docking study revealed that the nitroxyethyl moiety of compound 5c may form
hydrogen bonds with caspase‐8 amino acid residues (SER256 and HIS255). To-
gether, these data suggest that NO‐donating ursolic acid–arylidene derivatives are
potent apoptosis inducers in tumor cells.
2School of Nursing, Guangzhou University of
Chinese Medicine, Guangzhou, China
3Department of Research and Development,
Shanghai Hequan Pharmaceutical Co. Ltd.,
Shanghai, China
Correspondence
Jiamei Lei and Tao Wang, School of
Pharmaceutical Sciences, Guangzhou
University of Chinese Medicine, Waihuan East
Rd., No. 232, Guangzhou Higher Education
Mega Center, 510006 Guangzhou, China.
Email: ljm123@gzucm.edu.cn and
wangtao@gzucm.edu.cn
Funding information
Postdoctoral research from Guangzhou
University of Chinese Medicine,
Grant/Award Number: AFD01820Z0373
K E Y W O R D S
antitumor activity, apoptosis inducer, NO donor prodrug, ursolic acid derivatives
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1
INTRODUCTION
In general, there is a balance between apoptosis and proliferation
in normal cells.[7] The balance depends on the Bcl‐2/Bax ratio. It is
Apoptosis (also called programmed cell death) was described to per-
form various functions in development.[1] It plays an important role
not only in embryogenesis but also in the regulation of cell numbers
and the elimination of unwanted cells. The previous literature pointed
out that irregularity in apoptosis may cause a number of human dis-
eases, such as neurodegenerative disorders, immunodeficiency, AIDS,
and cancer.[2–4] Subsequent studies confirmed that apoptosis is one of
the major pathways involved in the proliferation, migration, and in-
vasion of tumor cells. Most of the cytotoxic compounds were reported
to induce apoptosis in cancer cells. The developments of apoptosis
inducers have become effective methods for cancer therapy.[5,6]
worth noting that high levels of Bcl‐2/Bax ratio may increase the risk
of cancer. An excess of Bcl‐2 promotes migration and invasion of
tumor cells.[8] Morphological alterations, including cell shrinkage,
chromatin condensation, and nuclear fragmentation, are important
features of tumor cell apoptosis.[9] It has been proved that most of the
morphological changes are caused by caspase family proteases. Cas-
pases are divided into three subfamilies, I (caspase‐2, ‐8, ‐9, and ‐10),
II (caspase‐3, ‐6, and ‐7), and III (caspase‐1, ‐4, ‐5, ‐11, ‐12, ‐13,
and ‐14).[10] Caspases I and II are involved in the activation and ex-
ecution of apoptosis. During the process, initiator caspases activate
downstream caspases, which are responsible for the cleavage of
Baoen He and Zuchang Zhu contributed equally to this study.
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Arch Pharm. 2021;e2000448.
wileyonlinelibrary.com/journal/ardp
© 2021 Deutsche Pharmazeutische Gesellschaft
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specific substrates and eventually cause apoptosis.[11] Currently,
caspase‐8 is considered as a key activator of apoptosis,[12] which can
be activated by dimerization of procaspase‐8 monomers.[13,14] The
activation of caspase‐8 may lead to stimulate caspase‐3 and result in
caspase protein cascade reaction.[15,16] Therefore, caspase‐8 was
considered as one of the major targets for cancer treatment. It is
focused on by many pharmaceutical companies. However, the non-
apoptotic functions of caspase‐8 have also been gradually recognized
in recent years.[17]
compounds were investigated for their antitumor activity against a
panel of human tumor cell lines (A549, MCF‐7, HepG‐2, HT‐29, and
HeLa). Subsequently, a molecular docking study was carried out to
clarify the binding mode between the most active compound and
caspase‐8. Finally, we intend to discover a new UA derivative as
a potential apoptosis inducer via the caspase‐8 activation.
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2
RESULTS AND DISCUSSION
Chemistry
Ursolic acid (UA) is a ursane‐type pentacyclic triterpene. As a
ubiquitous chemical constituent derived from plants, UA exhibited an
antitumor‐promoting effect in vitro and in vivo.[18,19] Multiple mole-
cular targets of UA have been proposed in many studies of anti-
proliferative mechanism, and most of them are related to apoptosis.
Moreover, the reasonable modifications of the UA skeleton at posi-
tions 2, 3, and 28[20–22] have been found to increase antitumor activity
and apoptosis induction. All of the above suggested that UA is an ideal
starting point for the development of new antitumor agents.[23] The
introduction of the Michael acceptor group in the parent structure is
thought to improve the activity.[24,25] These active products are com-
monly obtained by using arylideneketones as convenient starting ma-
terials via Claisen–Schmidt condensation.[26] However, nitric oxide
produced in large amounts by NO‐releasing prodrug is known to
mediate cytotoxicity. Thus, the design of NO‐releasing prodrug has
been a major topic of medicinal chemistry.[27,28]
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2.1
Heterocyclic compounds have been reported to possess a wide range
of biological activities, including antibacterial, antitumor, anti‐
inflammatory, and anticonvulsant effects. The synthesis, the physical
properties, and the biological activities of hybrid compounds con-
taining different heterocyclic fragments have been widely in-
vestigated.[30–33] Motivated by these findings, we designed and
synthesized new arylidene UA derivatives incorporating oxygen‐,
sulfur‐, and nitrogen‐containing heterocycles. Furan, thiophene,
pyridine, and pyrimidine derivatives were reported to have anti-
tumor activity.[34–36] Therefore, we designed the synthetic route of
UA hybrid molecules containing heterocyclic moieties mentioned
above. The target compounds were prepared according to the re-
actions outlined in Scheme 1.
Previously, we found that the hybrid compounds containing the
NO‐releasing UA scaffold linked to the Michael acceptor group ex-
hibited a potent antitumor activity.[29] In this study, we describe
further optimization for the design of NO‐donating UA derivatives
combined with heterocyclic aldehydes. The newly synthesized
First, it is necessary to prepare an active methylene inter-
mediate 2 for the introduction of heterocyclic moieties. Second,
Claisen–Schmidt condensation of 2 with different heterocyclic
formaldehyde (furaldehyde, 2‐thenaldehyde, 4‐pyridylaldehyde,
and 4,6‐dimethoxypyrimidine‐5‐carbaldehyde) was carried out in
SCHEME 1 Synthesis of NO‐donating ursolic acid derivatives containing heterocyclic moieties 5a–d and 6a–d. Reagents and conditions:
(a) Dess–Martin periodinane, CH2Cl2, rt, 6 h; (b) aromatic aldehydes, EtOH, KOH, rt, 5–24 h; (c) Br(CH2)2Br, DMF, K2CO3, rt, 12 h; (d) AgNO3,
CH3CN, 50°C, 24 h; (e) NaBH4, THF + EtOH, rt, 2 h

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ethanol under base catalysis, as described in our previous paper.[29]
NO‐donating UA derivatives 5a–d containing heterocyclic moieties
were synthesized via two‐step nucleophilic substitution, which has
been widely used in the preparation of organic nitrate prodrugs.
Compounds 5a–d were reduced with sodium borohydride in tet-
rahydrofuran (THF) to give another series of NO‐donating UA de-
rivatives containing heterocyclic moieties 6a–d. All of the target
compounds were purified by silica gel column chromatography.
Then, their structures were confirmed by infrared (IR), nuclear
magnetic resonance (NMR), and high‐resolution mass spectro-
metry (HRMS).
used simultaneously to determine cell viability.[37] However, several
studies reported that MTT and CCK‐8 assay exhibited inconsistent
results.[38] It is important to note that we need to select the suitable
method of cell viability based on its advantages and scope of appli-
cation. Considering that MTT assay is generally applied to measure
the proliferation of adherent cells,[39] we chose this method for
evaluation of the antiproliferative activity.
All new compounds, 5a–d and 6a–d, were screened for their in
vitro antitumor activity against five human tumor cells: A549,
MCF‐7, HepG‐2, HT‐29, and HeLa cells. As control groups, a culture
medium, a culture medium containing 0.1% dimethyl sulfoxide
(DMSO), UA, and cisplatin were used in the experiment. We de-
termined cell viability after tumor cell lines were treated with var-
ious concentrations of test compounds for 48 h. The mean IC50
values of the compounds are shown in Table 1.
All spectral data were consistent with the proposed structure. The
characteristic IR absorption bands of OH, C═O, C═C, and C–O–C were
observed at ~3557, ~1669 to 1730, ~1637, and ~1280 cm−1, respec-
tively. Furthermore, the data of NMR provided strong support for the
existence of heterocyclic moieties and the formation of nitroxyethyl. The
proton signals at δ ~4.65 and ~4.28 ppm indicated the presence of two
methylene groups in nitroxyethyl. The proton signals of heterocyclic and
methyl groups in the UA ring were also found in the expected region. We
found that a new proton signal emerged at ~3.85 ppm after reduction of
the C‐3 carbonyl group. In addition, the chemical shifts of methylene
carbon in heterocyclic appeared at 101.2–167.6 ppm. The signals (~70.6
and ~60.2 ppm) were assigned to the methylene carbon of nitroxyethyl.
It is clear that UA displayed a considerable inhibitory activity in
vitro against all tested tumor cell lines, with IC50 values ranging from
40.28 μM to 66.53 μM. The viability of all cell lines was not inhibited
by thiophene‐containing NO‐donating UA derivatives 5b and 6b at
100 μM maximum concentration. Interestingly, furan‐containing and
pyrimidine‐containing derivatives 5a, 6a, and 6d were found to ex-
hibit selective inhibition against different tumor cell lines. Moreover,
pyridine‐containing NO‐donating UA derivatives 5c and 6c showed a
significant broad‐spectrum antitumor activity. The inhibitory effect
was exerted in a dose‐dependent manner. Compound 5c displayed
three‐ to nine‐fold greater inhibitory activity against five tumor cell
lines as compared with UA. Among all the tumor cell lines tested,
MCF‐7 was found to be most sensitive to compound 5c. Further
study of apoptosis was investigated using only this cell line. The
results of the cytotoxic evaluation indicated that the introduction of
pyridine structure into the UA skeleton may improve the antitumor
activity. In addition, the presence of the carbonyl group at the C3
position is necessary for the inhibitory effect.
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2.2
Antitumor activity
It is well known that sulforhodamine B (SRB), 3‐(4,5‐dimethylthiazol‐
2‐yl)‐2,5‐diphenyltetrazolium bromide (MTT), and cell‐counting kit‐8
(CCK‐8) assays are used infrequently for in vitro anticancer drug
screening. Previous studies demonstrated that the results of the
MTT test are well correlated with those of the SRB assay. To ensure
the reliability of the results, these two methods have always been
TABLE 1 IC50 values (μM) of UA derivatives 5a–d and 6a–d against different tumor cell lines
IC50 (μM)
Lung
Breast cancer
Liver cancer
Colon cancer
Cervical
Compound
cancer A549
MCF‐7
HepG‐2
HT‐29
cancer HeLa
5a
24.45 0.06
>100
>100
>100
>100
>100
5b
>100
>100
>100
>100
5c
7.96 0.61
>100
7.42 0.51
>100
13.35 0.18
>100
12.99 0.02
>100
17.05 0.06
>100
5d
6a
>100
14.48 0.44
>100
>100
43.88 0.86
>100
>100
6b
>100
>100
>100
6c
13.77 0.28
>100
12.74 1.10
13.96 0.05
66.53 1.03
23.06 0.12
20.13 0.44
>100
17.21 0.36
>100
33.44 1.08
>100
6d
1
45.36 0.84
22.23 0.04
43.25 0.45
22.42 0.12
40.28 0.56
13.44 0.10
56.30 0.44
8.67 0.03
Cisplatin
Note: IC50 values are the mean SD of three individual experiments determined after 48‐h treatment. Cisplatin is a positive control.

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2.3
Apoptosis detection
of compound 5c induced early apoptosis in MCF‐7 cells, whereas high
concentrations of compound 5c mainly induced late apoptosis.
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2.3.1
Morphological changes detection
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Apoptosis and cell cycle detection
Apoptosis is known as a common mode of death in many tumor cells.
As described previously, the apoptotic process of cells was accom-
panied by morphological changes.[40] Light microscopy, electron mi-
croscopy, and DNA electrophoresis analysis are powerful tools in the
analysis of morphological changes in apoptotic cells.[41] Fluorescent
staining techniques, which can identify apoptotic changes in the
nucleus, are widely applied to measure the apoptosis of tumor cells
in recent years.[42] Here, 4′,6‐diamidino‐2‐phenylindole (DAPI)
and acridine orange/ethidium bromide (AO/EB) staining were used
to observe morphological changes characteristic of apoptosis in the
MCF‐7 cells.
2.3.2
To quantitatively determine apoptosis of MCF‐7, annexin V‐fluorescein
isothiocyanate (FITC)/propidium iodide (PI) staining was performed
in our study. Annexin V has a high affinity for phosphatidylserine.
Translocation of membrane phosphatidylserine was found at the
early stage of apoptosis. Hence, FITC‐labeled annexin V can be used
for the detection of early‐stage apoptosis using flow cytometry.
Once cells entered the late‐stage apoptosis, apoptotic nuclei were
stained by PI due to the disappearance of cell membranes integrity.
It is indicated that the different stages of apoptotic cells can
be distinguished by the combination of annexin V‐FITC and PI.[44]
As shown in Figure 2a, the proportion of early and late apoptotic
cells was increased in a dose‐dependent manner. We also found
that late‐stage apoptosis predominated at 2 × IC50 concentration of
compound 5c.
The staining results are shown in Figure 1. DAPI is a DNA‐specific
fluorescent dye, which can stain nuclei of tumor cells as blue spots.
During apoptosis of tumor cells, cell membrane integrity was lost and
subsequent apoptotic characteristics appeared. Images of DAPI staining
revealed that the cells of the control group possess large‐volume nuclei
and intact membrane. After MCF7 cells were treated with compound 5c,
nuclei pyknosis and DNA fragmentation were found (Figure 1a). The
occurrence of apoptosis induced by compound 5c was preliminary
confirmed. The results were further substantiated by AO/EB staining.
AO/EB staining is another useful detection of changes in nuclei.[43] Many
publications have reported that live cells with intact membrane were
stained green by AO. Late apoptotic cells were stained orange by EB
when their membrane integrity was lost. The same results were found in
our study (Figure 1b). These findings suggested that low concentrations
Mammalian development and homeostasis are dependent on the
regulation of apoptosis and the cell cycle.[45] It has been believed
that progression through the different phases of the cell cycle is
governed by the cyclin‐dependent kinases (Cdks).[46] Different Cdks
functioned at specific stages during cell cycle progression.[47]
Therefore, targeting the cell cycle provides a valuable strategy for
the inhibition of tumor cell proliferation.[48] Previous evidence has
shown that antitumor agents mainly arrested cell cycle at S and G1
phases.[49] To investigate the effects of compound 5c on the cell
FIGURE 1 Cellular morphologic observation of MCF‐7 after treatment with different concentrations of compound 5c (0, IC50, and 2IC50).
(a) 4′,6‐Diamidino‐2‐phenylindole staining; (b) acridine orange/ethidium bromide staining

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FIGURE 2 Apoptosis and cell cycle assay of compound 5c on MCF‐7. (a) Cells were treated with different concentrations of compound 5c
(0, IC50, and 2 × IC50) for 24 h, stained with annexin V‐FITC/PI, and observed by flow cytometry; (b) cells were treated with different
concentrations of compound 5c (0, IC50 and 2 × IC50) for 24 h, stained with PI, and observed by flow cytometry. FITC, fluorescein
isothiocyanate; PI, propidium iodide
cycle of MCF‐7 cells, the cell cycle was determined by staining DNA
with PI and flow cytometry. Figure 2b indicated that compound 5c
caused an increase of the G1 phase cell population and a decrease of
the G2 phase cell population after 24 h of treatment. A dose‐
dependence effect on the cell cycle was also observed. Multiple
studies have confirmed that DNA replication is initiated in the G1
phase. The tumor suppressor p53 is involved in G1 phase arrest and
induction of apoptosis.[50] These facts give a good reason for us to
speculate that compound 5c induced G1 phase arrest via regulating
p53 expression in MCF‐7 cells.
membrane potential is one early hallmark of apoptosis.[52] In the
present study, ROS level and membrane potential were measured by
using flow cytometry simultaneously.
JC‐1 probe has been frequently used to monitor mitochondrial
potential due to its fluorescent properties depending on membrane
potential. When the mitochondrial membrane potential changed, the
transition between JC‐1 dye monomer and J‐aggregates resulted in a
fluorescence shift from green to red. The results of mitochondrial
membrane potential analysis are shown in Figure 3a and displayed as
an image composed of four quadrants. JC‐1 monomer accumulated in
control cells and exhibited a high ratio of green fluorescence. After
treatment with different concentrations of compound 5c, the
red/green ratio in Q3 quadrants increased to 14.9%, and 20.7%,
respectively. The results indicated that the apoptosis of MCF‐7 in-
duced by compound 5c was associated with mitochondrial mem-
brane depolarization and was carried out in a dose‐dependent
manner.
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2.3.3
Membrane potential and reactive oxygen
species (ROS) detection
To date, both ROS and mitochondria involved in apoptosis induction
have been recognized. ROS are considered as an effective activator
of mitochondrial permeability transition pore (MPTP) opening. The
loss of mitochondrial membrane potential is observed after the
irreversible opening of MPTP.[51] This is accompanied by the release
of cytochrome c, activation of caspase cascades, and ultimately
apoptosis. All this evidence suggests that the loss of mitochondrial
A previous study has pointed out that ROS are involved in dif-
ferent pro‐ and anticarcinogenic mechanisms.[53] A moderate in-
crease in ROS can promote cell proliferation and differentiation.
However, a high level of ROS tends to inhibit tumor cell growth and
induce apoptosis.[54] The overproduction of ROS may lead to

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FIGURE 3 Membrane potential and reactive oxygen species assay of compound 5c on MCF‐7. (a) Cells were treated with different concentrations
of compound 5c (0, IC50 and 2 × IC50) for 24 h, stained with JC‐1, and determined by flow cytometry; (b) cells were treated with different concentrations
of compound 5c (0, IC50, and 2 × IC50) for 24 h, stained with 2ʹ,7ʹ‐dichlorofluorescein diacetate, and analyzed by flow cytometry
oxidative stress, which can cause damages in DNA, proteins, and
lipids. Numerous investigations have documented that many che-
motherapeutic drugs induce apoptosis by increasing ROS production.
To confirm the effect of compound 5c on ROS in MCF‐7 cells, the
level of ROS was tested by 2ʹ,7ʹ‐dichlorofluorescein diacetate
(DCFH‐DA). After the penetration of DCFH‐DA into the cell mem-
brane, it was hydrolyzed by intracellular esterase to DCFH. Further
oxidization of DCFH by ROS produced DCF, which is a highly
fluorescent compound. Many studies have found that there is a
quantitative relationship between the DCF fluoresce intensity and
the amount of ROS.[55] Figure 3b showed that the level of ROS in
treated groups notably increased to 13.8% and 24.6% as compared
with that in the control group (3.42%). These results demonstrated
that compound 5c could increase the level of ROS in a dose‐
dependent manner. So, we infer that ROS production may be a
possible mechanism for apoptosis induction of MCF‐7 cells by com-
pound 5c.
mitochondria. A number of previous reports have indicated that the
loss of the mitochondrial membrane potential was associated with the
Bcl‐2 (antiapoptotic protein)/Bax (proapoptotic protein) ratio.[56,57]
Subsequently, the release of cytochrome c and activation of caspases
cascade were observed.[58] Therefore, we investigated a number of key
protein markers by the Western blot analysis. The results of the
Western blot analysis are shown in Figure 4. After treatment with
different concentrations of compound 5c, the expression of Bax, cas-
pase 3, and caspase 9 was remarkably upregulated. Besides, Bcl‐2 ex-
pression was downregulated in MCF‐7 cancer cells. It is reasonable to
presume that compound 5c induces apoptosis of MCF‐7 tumor cells
through regulating the Bcl‐2/Bax ratio, leading to the loss of the mi-
tochondrial membrane potential and activation of caspases.
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2.3.4
Western blot analysis
FIGURE 4 The results of Western blot analysis of compound 5c
on MCF‐7 cells. Cells were treated with different concentrations of
compound 5c (0, 0.5×IC50, IC50 and 2 × IC50) for 24 h and total
protein was extracted to determine the expression of caspase‐9,
caspase‐3, Bcl 2, and Bax
It is widely recognized that most anticancer drugs induce apoptosis
through the mitochondrial pathway. Mitochondrial membrane poten-
tial plays an important role in maintaining the normal function of

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2.4
Molecular docking studies
evaluation against five tumor cell lines (A549, MCF‐7, HepG‐2,
HT‐29, and HeLa cells) and molecular recognition were undertaken.
The MTT assay showed that pyridine‐containing NO‐donating
UA derivatives 5c and 6c exhibited a broad spectrum of antitumor
activity, with IC50 values better than those of the parent compound
(UA). Compound 5c was the most active derivative against MCF‐7
cells (IC50 7.42 μM). A preliminary mechanism of antitumor activity
demonstrated that the cytotoxicity of compound 5c against MCF‐7
cells may be related to the decrease of mitochondrial membrane
potential, the increase in the level of ROS, and cell cycle arrest in the
G1 phase to induce apoptosis. The expression of key protein markers
hinted that compound 5c induced MCF‐7 cells apoptosis in a
mitochondrial‐dependent manner. Moreover, compound 5c effec-
tively interacted with the active site of caspase 8 via hydrogen
bonds. Our results indicated that compound 5c may be a promising
anticancer agent worthy of further study.
Caspase‐8 is one of the critical initiators of apoptosis. Apoptosis
induced by anticancer drugs is mediated by the activation of caspase‐
8 in many tumor cells. It has also been found that caspase‐8 may
serve as an amplifier of apoptotic signals.[59] Thus, targeting caspase‐
8 to initiate apoptosis may offer a promising therapeutic opportunity
in cancer. Here, the further study on the docking interactions of
potent compound and caspase‐8 was performed by computational
analysis. The binding mode of compound 5c and caspase‐8 was stu-
died by docking and is presented in Figure 5. On the basis of the total
score of 7.31, compound 5c showed an excellent interaction within
the 2c2z protein pocket of caspase‐8. The two‐dimensional docking
interaction between compound 5c and caspase‐8 active site is pre-
sented in Figure 5a. It should be noted that pi–alkyl interactions
were colored in pink, whereas the hydrophobic interactions were
colored in green. As seen in Figure 5a, pi–alkyl interactions were
found between the UA skeleton of compound 5c and residues of
caspase‐8, including CYS360, VAL410, ARG258, HIS317, and ILE257,
wherein the pi–alkyl distances ranged from 2.06 Å to 4.01 Å. More-
over, SER256 and HIS255 residues of caspase‐8 are engaged in ad-
ditional hydrophobic interactions with nitroxyethyl moiety
(distances: 1.93 Å and 2.26 Å). Figure 5ab showed that nitroxyethyl
moiety of compound 5c docked into the hydrophobic pocket of
caspase‐8. It might be assumed that the 2c2z‐pocket of caspase‐8
participates in the apoptotic regulation by acting as a docking site for
substrates.
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4
EXPERIMENTAL
Chemistry
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4.1
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4.1.1
General
All chemicals and reagents were obtained from commercial suppliers.
When necessary, reagents were dried and purified before use. All
reactions were monitored by thin‐layer chromatography (TLC) on
silica gel GF254 plates and spots were visualized with 5%
vanillin–sulfuric acid. Melting points were measured on the X‐6 mi-
croscopic melting point apparatus and were uncorrected. IR spectra
were obtained on a Nicolet 330 FT‐IR spectrometer. NMR spectra
were determined in CDCl3 on a Bruker Avance spectrometer
(400 MHz for 1H NMR, 100 MHz for 13C NMR). High‐resolution mass
spectra were recorded on an AB SCIEX Triple TOF™ 5600+
LC‐MS/MS system. The original spectra can be found in the
Supporting Information.
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3
CONCLUSION
In recent years, pentacyclic triterpenoids have received much at-
tention as lead compounds for the development of the new anti-
tumor drug. Lupane‐, oleanane‐, and ursane triterpenoids are
particularly prominent among the known research. In view of the
antitumor effects of pentacyclic triterpenoids and their semisyn-
thetic derivatives, we synthesized eight new arylidene derivatives
containing NO‐donating UA and aromatic heterocyclic units. In vitro
The InChI codes of the investigated compounds, together with some
biological activity data, are provided as Supporting Information.
FIGURE 5 Docking interaction between compound 5c and caspase‐8 (PDB: 2c2z). (a) Two‐dimensional structure of compound 5c docked
into the caspase‐8 active site; (b) three‐dimensional conformation of compound 5c bound to caspase‐8 interface region

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177.3 (C28), and 207.4 (C3) ppm; HRMS (m/z) [M+H]+ calcd. for
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4.1.2
General procedure for the preparation of
intermediates 2, 3a–d, 4a–d, and compounds 5a–d
C37H52SNO6: 638.3510, found: 638.3504.
The intermediates were synthesized as described in Reference [29] .UA
1 (1 eq.) was dissolved in dichloromethane and subjected to oxidation
by Dess–Martin periodinane (1 eq.) to give compound 2. The con-
densation of compound 2 (1 eq.) with various aromatic aldehydes
(1–1.5 eq.), namely, furaldehyde, 2‐thenaldehyde, 4‐pyridylaldehyde,
and 4,6‐dimethoxypyrimidine‐5‐carbaldehyde, in ethanol and in
the presence of a base, afforded compounds 3a–d. Compounds 4a–d
were obtained by stirring of compounds 3a–d (1 eq.) with 1,
2‐dibromoethane (4–8 eq.) at room temperature for 12 h in 41–73%
yields. Finally, silver nitrate (10–12 eq.) was added to a solution of
compounds 4a–d (1 eq.) in acetonitrile. The mixture was heated at
50°C for 24 h. The products 5a–d were prepared as yellow or yellow‐
white or yellow‐green solids.
2‐[2‐(Pyridin‐4‐yl)vinyl]‐3‐oxo‐urs‐12‐en‐28‐oic acid‐(2‐nitrooxy)‐
ethyl ester (5c)
Yellow‐white solid; yield 55.5%; m.p. 113.9–114.5°C; IR (KBr):
~
v
= 2924 (vs), 2869 (m) (C–H, aliphatic), 1728 (s), 1669 (m) (2C═O),
1636 (vs) (C═C), 1455 (m), 1579 (m) (C═N, pyridinyl), and 1280 (vs)
(C–O–C, ester) cm−1; 1H NMR (400 MHz, CDCl3): δ = 0.77, 0.86, 1.12,
1.14, 1.16 (15H, s, 5× CH3), 0.90 (3H, d, J = 6.4 Hz, CH3), 0.94 (3H, d,
J = 6.1 Hz, CH3), 4.25 (2H, m, –OCH2), 4.63 (2H, m, CH2ONO2), 5.28
(1H, t, J = 7.0, 3.5 Hz, H‐12), 7.27 (2H, d, J = 5.8 Hz, 3’, 5’‐H), 7.38 (1H,
s, ═CHAr), and 8.64 (2H, d, J = 5.0 Hz, 2’, 6’‐H) ppm; 13C NMR
(100 MHz, CDCl3): δ = 15.6 (C29), 16.8 (C30), 17.2 (C23), 20.4 (C6),
21.2 (C24), 22.9 (C27), 23.5 (C26), 23.7 (C11), 24.2 (C22), 28.0 (C21),
29.6 (C25), 30.7 (C7), 32.2 (C16), 36.5 (C10), 36.7 (C15), 38.9
(C19), 39.2 (C20), 39.6 (C8), 42.4 (C17), 44.0 (C1), 45.3 (C18), 45.5
(C14), 48.5 (C4), 53.1 (C9), 53.3 (C5), 60.2 (COOCH2), 70.6
(CH2ONO2), 124.2, (C3’, C5’), 125.6 (C12), 134.2 (═CHAr), 138.0
(C4’), 138.2 (C13), 143.7 (C2), 150.0 (C2’, C6’), 177.3 (C28), and
207.5 (C3) ppm; HRMS (m/z) [M–H]– calcd. for C38H51N2O6:
631.3753, found: 631.3744.
2‐[2‐(Furan‐2‐yl)vinyl]‐3‐oxo‐urs‐12‐en‐28‐oic acid‐(2‐nitrooxy)‐
ethyl ester (5a)
~
Yellow solid; yield 90.3%; m.p. 123.3–123.9°C; IR (KBr):
v
= 2924
(vs), 2869 (m) (C–H, aliphatic), 1730 (s), 1672 (m) (2C═O), 1637 (vs)
(C═C), 1594 (m), 1542 (w), 1455 (m) (C═C, furanyl), and 1280 (vs)
(C–O–C, ester) cm−1; 1H NMR (400 MHz, CDCl3): δ = 0.80, 0.90, 1.10,
1.14, 1.16 (15H, s, 5× CH3), 0.92, 0.96 (6H, d, J = 6.4, 6.2 Hz, 2× CH3),
4.30 (2H, m, –OCH2), 4.65 (2H, m, CH2ONO2), 5.34 (1H, t, J = 7.0,
3.3 Hz, H‐12), 6.51 (1H, m, 4’‐H), 6.61 (1H, d, J = 3.4 Hz, 3’‐H), 7.33
(1H, s, ═CHAr), and 7.57 (1H, d, J = 1.6 Hz, 5’‐H) ppm; 13C NMR
(100 MHz, CDCl3): δ = 15.9 (C30), 16.8 (C29), 17.2 (C27), 20.5 (C6),
21.3 (C23), 22.7 (C26), 23.5 (C24), 23.8 (C11), 24.2 (C16), 28.0 (C21),
30.0 (C25), 30.7 (C15), 32.2 (C1), 35.8 (C10), 36.7 (C22), 39.0
(C20), 39.3 (C19), 39.5 (C8), 42.6 (C14), 44.4 (C7), 45.0 (C17),
45.3 (C9), 48.5 (C4), 52.9 (C18), 53.2 (C5), 60.2 (COOCH2), 70.7
(CH2ONO2), 112.4 (C3’), 115.7 (C4’), 124.4 (═CHAr), 125.9 (C12),
130.9 (C13), 138.2 (C2), 144.5 (C5’), 152.7 (C2’), 177.3 (C28), and
207.3 (C3) ppm; HRMS (m/z) [M+H]+ calcd. for C37H52NO7:
622.3738, found: 622.3733.
2‐[2‐(4,6‐Dimethoxypyrimidin‐5‐yl)vinyl]‐3‐oxo‐urs‐12‐en‐28‐oic
acid‐(2‐nitrooxy)ethyl ester (5d)
Yellow‐green solid; yield 88.2%; m.p. 105.7–106.2°C; IR (KBr):
~
v
= 2953 (s), 2869 (w) (C–H, aliphatic), 1729 (s), 1680 (s) (2C═O),
1637 (vs) (C═C), 1565 (vs) (C═N, pyrimidinyl), and 1280 (vs) (C–O–C,
ester) cm−1 1H NMR (400 MHz, CDCl3): δ = 0.77, 0.89, 1.08, 1.14,
;
1.16 (15H, s, 5× CH3), 0.84 (3H, d, J = 6.4 Hz, CH3), 0.93 (3H, d,
J = 6.1 Hz, CH3), 3.98 (6H, s, 2× OCH3), 4.28 (2H, m, –OCH2), 4.64
(2H, m, CH2ONO2), 5.24 (1H, t, J = 6.8, 3.7 Hz, H‐12), 7.24 (1H, d,
J = 2.5 Hz, 2’‐H), and 8.41 (1H, s, ═CHAr) ppm; 13C NMR (100 MHz,
CDCl3): δ = 15.3 (C29), 16.9 (C30), 17.1 (C23), 20.3 (C6), 21.2
(C24), 21.2 (C27), 23.2 (C26), 23.5 (C11), 24.3 (C22), 28.0 (C21), 29.3
(C25), 30.7 (C7), 32.4 (C16), 36.3 (C10), 36.7 (C15), 38.9 (C19), 39.2
(C20), 39.6 (C8), 42.7 (C17), 43.8 (C1), 45.2 (C14), 45.6 (C18), 48.5
(C4), 53.1 (C9), 53.8 (C5), 54.4 (2×OCH3), 60.2 (COOCH2), 70.7
(CH2ONO2), 101.1 (C5’), 125.8 (C12), 127.3 (═CHAr), 137.6 (C13), 138.1
(C2), 156.5 (C2’), 167.6 (C4’, C6’), 177.3 (C28), and 206.6 (C3) ppm;
HRMS (m/z) [M+H]+ calcd. for C39H56N3O8: 694.4062, found: 694.4059.
2‐[2‐(Thiophene‐2‐yl)vinyl]‐3‐oxo‐urs‐12‐en‐28‐oic acid‐(2‐nitrooxy)
ethyl ester (5b)
~
Yellow solid; yield 70.3%; m.p. 127.4–128.2°C; IR (KBr):
v
= 2949
(vs), 2869 (m) (C–H, aliphatic), 1729 (s), 1669 (m) (2C═O), 1636 (vs)
(C═C), 1579 (m), 1455 (m) (C═C, thiopheneyl), and 1280 (vs)
(C–O–C, ester) cm‐1; 1H NMR (400 MHz, CDCl3): δ = 0.80, 0.91, 0.94,
0.96, 1.10, 1.14, 1.18 (21H, s, 7× CH3), 4.31 (2H, m, –OCH2), 4.65
(2H, m, CH2ONO2), 5.36 (1H, t, J = 7.0, 3.3 Hz, H‐12), 7.14 (1H, q,
J = 5.1, 3.7 Hz, 4’‐H), 7.32 (1H, d, J = 3.5 Hz, 5’‐H), 7.51 (1H, d, J = 5.1 Hz,
3’‐H), and 7.74 (1H, s, ═CHAr) ppm; 13C NMR (100 MHz, CDCl3):
δ = 16.0 (C29), 16.8 (C30), 17.2 (C23), 20.6 (C6), 21.3 (C24), 22.7 (C27),
23.6 (C26), 23.7 (C11), 24.3 (C22), 28.0 (C21), 30.1 (C25), 30.7 (C7),
32.2 (C16), 36.2 (C10), 36.7 (C15), 39.0 (C19), 39.3 (C20), 39.6 (C8),
42.4 (C17), 44.8 (C1), 45.0 (C14), 45.6 (C18), 48.5 (C4), 52.8 (C9), 53.1
(C5), 60.2 (COOCH2), 70.7 (CH2ONO2), 125.9 (C12), 127.7 (C3’), 129.8
(C4’), 130.5 (C5’), 130.8 (C2’), 132.8 (═CHAr), 138.1 (C13), 139.6 (C2),
|
4.1.3
General procedure for the preparation of
compounds 6a–d
To a solution of compounds 5a–d (1 eq.) in dry THF (25 ml) and
absolute ethanol (5 ml), sodium borohydride (10 eq.) was added at
0°C. The mixture was stirred in the dark for 2 h. At the end of the
reaction, 20 ml of distilled water was added and the pH was adjusted
to 7 with 1 M hydrochloric acid. The mixture was then extracted with
ethyl acetate (20 ml × 3), dried over anhydrous MgSO4, and con-
centrated under reduced pressure. The crude product was purified

HE ET AL.
9 of 12
|
by silica gel column chromatography (PE/EtOAc, 25:1) to give cor-
3.90 (1H, s, H‐3), 4.24–4.27 (2H, m, –OCH2), 4.62–4.64 (2H, m,
CH2ONO2), 5.23 (1H, t, J = 6.8, 3.4 Hz, H‐12), 6.69 (1H, s, ═CHAr),
7.18 (2H, d, J = 5.56 Hz, 3’, 5’‐H), and 8.53 (2H, d, J = 4.88 Hz, 2’, 6’‐
H) ppm; 13C NMR (100 MHz, CDCl3): δ = 15.7 (C30), 15.9 (C29), 17.0
(C27), 17.1 (C25), 18.5 (C6), 21.3 (C24), 23.4 (C11), 23.6 (C23), 24.3
(C16), 28.0 (C21), 28.9 (C26), 30.7 (C1), 32.9 (C15), 36.8 (C22), 38.9
(C20), 39.2 (C19), 39.9 (C10), 40.6 (C8), 42.1 (C14), 42.2 (C7), 42.3
(C17), 47.0 (C9), 48.4 (C4), 52.9 (C18), 55.8 (C5), 60.2 (COOCH2),
70.6 (–CH2ONO2), 81.0 (C3), 120.6 (C3’), 120.6 (C5’), 124.2 (C12),
125.5 (═CHAr), 138.1 (C13), 145.4 (C4’), 147.5 (C2), 148.6 (C2’),
148.6 (C6’), and 177.3 (C28, C═O) ppm; HRMS (m/z) [M+Na]+ calcd.
for C38H54N2O6Na: 657.3874, found 657.3870.
responding pure compounds 6a–d.
2‐[2‐(Furan‐2‐yl)vinyl]‐3β‐hydroxy‐urs‐12‐en‐28‐oic acid‐(2‐
nitrooxy)ethyl ester (6a)
~
Yellow solid; yield 66.4%; m.p. 198.4–198.9°C; IR (KBr):
v
= 3557 (s)
(–OH), 2926 (s), 2875 (s) (C–H, aliphatic), 1729 (vs) (C═O), 1643 (vs)
(C═C), 1457 (s), 1488 (m) (C═C, furanyl), and 1277 (s) (C–O–C, ester)
cm−1; 1H NMR (400 MHz, CDCl3): δ = 0.71, 0.74, 0.85 (9H, s, 3× CH3),
0.87 (3H, d, J = 6.44 Hz, CH3), 0.94 (3H, d, J = 6.20 Hz, CH3), 1.12 (6H,
d, J = 2.08 Hz, 2× CH3), 2.24 (1H, d, J = 11.32 Hz, 18‐H), 3.83 (1H, s,
3‐H), 4.23–4.32 (2H, m, –OCH2), 4.65 (2H, dd, J = 3.96, 5.28 Hz,
CH2ONO2), 5.31 (1H, t, J = 3.36, 4.20 Hz, H‐12), 6.23 (1H, d,
J = 3.24 Hz, 3‘‐H), 6.37 (1H, dd, J = 3.24, 1.88 Hz, 4’‐H), 6.44 (1H,
s, ═CHAr), and 7.35 (1H, d, J = 1.44 Hz, 5’‐H) ppm; 13C NMR
(100 MHz, CDCl3): δ = 15.4 (C30), 15.7 (C29), 17.0 (C27), 17.1 (C25),
18.6 (C6), 21.3 (C24), 23.5 (C11), 23.7 (C23), 24.3 (C16), 28.1 (C21), 28.7
(C26), 30.7 (C1), 33.0 (C15), 36.7 (C22), 38.9 (C20), 39.2 (C19), 39.9
(C10), 40.7 (C8), 42.3 (C14), 43.0 (C7), 42.3 (C17), 46.9 (C9), 48.4 (C4),
53.0 (C18), 56.2 (C5), 60.2 (COOCH2), 70.7 (–CH2ONO2), 81.3 (C3),
108.6 (C3’), 111.0 (C4’), 111.3 (═CHAr), 125.9 (C12), 138.1 (C13), 140.0
(C2’), 141.4 (C5’), 153.5 (C2), and 177.4 (C28, C═O) ppm; HRMS (m/z)
[M+H]+ calcd. for C37H54NO7: 624.3895, found: 624.3893.
2‐[2‐(4,6‐Dimethoxypyrimidin‐5‐yl)vinyl]‐3β‐hydroxy‐urs‐12‐en‐28‐
oic acid‐(2‐nitrooxy)ethyl ester (6d)
~
White solid; yield 84.8%; m.p. 105.6–106.5°C; IR (KBr):
v
= 3523 (br)
(–OH), 2952 (vs), 2871 (s) (C–H, aliphatic), 1729 (s) (C═O), 1639 (vs)
(C═C), 1567 (vs) (C═N, pyrimidinyl), and 1279 (s) (C–O–C, ester)
;
cm−1 1H NMR (400 MHz, CDCl3): δ = 0.62, 0.65, 0.78, 1.08, 1.13
(15H, s, 5× CH3), 0.84 (3H, d, J = 6.44 Hz, CH3), 0.93 (3H, d,
J = 6.04 Hz, CH3), 2.18 (1H, d, J = 11.28 Hz, 18‐H), 3.90 (1H, s, 3‐H),
3.96 (6H, s, 2× OCH3), 4.24 (2H, dd, J = 3.32, 2.92 Hz, –OCH2), 4.61
(2H, t, J = 5.24, 4.00 Hz, –CH2ONO2), 5.17 (1H, t, J = 3.40, 3.36 Hz,
12‐H), 6.15 (1H, s, ═CHAr), and 8.39 (1H, s, 2’‐H) ppm; 13C NMR
(100 MHz, CDCl3): δ = 14.8 (C30), 15.8 (C29), 16.9 (C27), 17.1 (C25),
18.6 (C6), 21.2 (C24), 23.4 (C11), 23.7 (C23), 24.2 (C16), 28.0 (C21), 28.6
(C26), 30.7 (C1), 32.9 (C15), 36.7 (C22), 38.9 (C20), 39.2 (C19), 39.8
(C10), 39.9 (C8), 41.8 (C14), 42.2 (C17), 43.9 (C7), 47.0 (C9), 48.4 (C4),
52.8 (C18), 55.7 (C5), 60.2 (COOCH2), 70.6 (–CH2ONO2), 81.1 (C3),
102.4 (C5’), 110.6 (C12), 125.7 (═CHAr), 138.0 (C13), 144.4 (C2), 154.9
(C2’), 167.5 (C4’), 167.5 (C6’), and 177.3 (C28, C═O) ppm; HRMS (m/z)
[M+H]+ calcd. for C39H58N3O8: 696.4218, found 696.4217.
2‐[2‐(Thiophene‐2‐yl)vinyl]‐3β‐hydroxy‐urs‐12‐en‐28‐oic acid‐(2‐
nitrooxy)ethyl ester (6b)
Pale‐yellow solid; yield 81.8%; m.p. 182.5–182.8°C; IR (KBr):
~
v
= 3555 (br) (–OH), 2924 (s), 2871 (s) (C–H, aliphatic), 1726 (s)
(C═O), 1641 (vs) (C═C), 1454 (s), 1509 (m) (C═C, thiopheneyl), and
1279 (s) (C–O–C, ester) cm−1 1H NMR (400 MHz, CDCl3): δ = 0.73
;
(6H, d, J = 5.12 Hz, 2× CH3), 0.87 (6H, d, J = 3.64 Hz, 2× CH3), 0.94
(3H, d, J = 6.12 Hz, CH3), 1.12 (6H, d, J = 5.88 Hz, 2× CH3), 2.23 (1H,
d, J = 11.24 Hz, 18‐H), 3.85 (1H, s, 3‐H), 4.27 (2H, m, –OCH2), 4.64
(2H, m, CH2ONO2), 5.31 (1H, t, J = 3.24, 3.40 Hz, H‐12), 6.81 (1H,
s, ═CHAr), 6.98 (2H, d, J = 3.52 Hz, 3’, 4’‐H), and 7.19 (1H, t, J = 2.96,
3.48 Hz, 5’‐H) ppm; 13C NMR (100 MHz, CDCl3): δ = 15.4 (C30), 15.8
(C29), 17.0 (C27), 17.1 (C25), 18.5 (C6), 21.3 (C24), 23.5 (C11), 23.7
(C23), 24.2 (C16), 28.1 (C21), 28.8 (C26), 30.7 (C1), 33.0 (C15), 36.7
(C22), 38.9 (C20), 39.2 (C19), 39.9 (C10), 40.9 (C8), 42.3 (C14), 42.3
(C17), 43.3 (C7), 46.8 (C9), 48.4 (C4), 53.0 (C18), 56.2 (C5), 60.2
(COOCH2), 70.7 (–CH2ONO2), 81.4 (C3), 115.8 (C3’), 124.1 (C4’),
125.9 (═CHAr), 126.7 (C12), 127.8 (C5’), 138.1 (C13), 139.2 (C2’),
140.7 (C2), and 177.4 (C28, C═O) ppm; HRMS (m/z) [M+H]+ calcd.
for C37H54NO6S: 640.3666, found 640.3617.
|
4.2
Biological assays
|
4.2.1
Cell lines and culture
Five human cancer cell lines consisting of the human lung cancer cell
line A549, human breast cancer cell line MCF‐7, human liver cancer
cell line HepG‐2, human colon cancer cell line HT‐29, and the human
cervical cancer cell line HeLa were purchased from the Shanghai
Institute of Biochemistry and Cell Biology (China). Cell lines were
cultured in RPMI 1640 medium supplemented with 10% FBS at 37°C
in a humidified 5% CO2 atmosphere. The medium was refreshed
every 2 days and the growth of cells was monitored regularly under
the inverted microscope.
2‐[2‐(Pyridin‐4‐yl)vinyl]‐3β‐hydroxy‐urs‐12‐en‐28‐oic acid‐(2‐
nitrooxy)ethyl ester (6c)
~
White solid; yield 57.5%; m.p. 205.9–206.7°C; IR (KBr):
v
= 3416 (br)
(–OH), 2926 (s), 2871 (m) (C–H, aliphatic), 1727 (m) (C═O), 1638 (vs)
(C═C), 1453 (w), 1545 (w), 1599 (m) (C═N, pyridinyl), and 1280 (s)
;
(C–O–C, ester) cm−1 1H NMR (400 MHz, CDCl3): δ = 0.69, 1.10, 1.15
|
4.2.2
MTT assay
(9H, s, 3× CH3), 0.76 (3H, d, J = 3.68 Hz, CH3), 0.84 (3H, d, J = 6.36 Hz,
CH3), 0.93 (3H, d, J = 6.24 Hz, CH3), 2.20 (1H, d, J = 11.08 Hz, 18‐H),
The tested tumor cell lines were harvested and seeded at a density of
1 × 103 cells/well into 96‐well plates. After 24 h of incubation in a

10 of 12
HE ET AL.
|
humidified atmosphere with 5% CO2 at 37°C, the cells were exposed
to various concentrations of compounds 5a–d and 6a–d in a medium
containing 0.1% DMSO for 48 h. Then, 10 μl of MTT (5 mg/ml, in
phosphate‐buffered saline [PBS]) was added to each well. Next, the
cells were incubated for another 4 h. The supernatant was discarded
and 150 μl of DMSO was added to dissolve the formazan. The ab-
sorbance was determined at 490 nm by a Multiskan FC Microplate
Reader (Thermo Fisher Scientific). The IC50 value was calculated by
GraphPad Prism Software. Negative control (cells+medium), blank
control (only medium), and positive control (cisplatin) were
co‐assayed.
and resuspended with 50 µg/ml PI and 0.1 mg/ml RNase A for 30 min
at room temperature. Cell cycle analysis was performed on the
FACSCanto II flow cytometer (BD Biosciences).
|
4.2.6
Membrane potential detection
MCF‐7 cells were harvested (2 × 104 cells/well) and incubated in
6‐well plates overnight. Then, the cells were treated with indicated
concentrations of compound 5c for 24 h. After treatment, the cells
were washed with PBS, resuspended in 500 µl of culture medium,
and incubated with 10 µg/ml JC‐1 for 20 min. Immediately after
centrifugation at 600g for 4 min, the cells were rinsed twice with
1× incubation buffer. JC‐1 fluorescence was determined by the
FACSCanto II flow cytometer (BD Biosciences).
|
4.2.3
AO/EB and DAPI staining
MCF‐7 cells were harvested (2 × 104 cells/well) and incubated in
6‐well plates overnight. The cells were treated with indicated
concentrations of compound 5c for 24 h. After treatment, the cells
were washed three times with PBS. Next, 500 µl of 1 × dilution buffer
containing 20 µl of a 1:1 (v/v) AO/EB mixture was added to the cells.
Then the cells were stained with AO/EB for 5 min at room
temperature in the dark, and 5 µl of cell suspension was transferred
to a sterile glass slide and covered with a coverslip. The morphology
of cells was imaged using a fluorescence microscope (Olympus) with
488‐nm excitation and 525‐nm emission.
|
4.2.7
ROS detection
MCF‐7 cells were harvested (2 × 104 cells/well) and incubated in
6‐well plates overnight. The cells were treated with indicated con-
centrations of compound 5c for 24 h. Thereafter, the medium was
removed, the cells were washed with PBS, and collected by cen-
trifugation (1000 r/min, 5 min). Then, 10 μM of DCFH‐DA was added
to the cells and incubated for 20 min at 37°C. Finally, the cells were
washed three times in a serum‐free medium and resuspended in PBS.
The generation of ROS was measured with excitation at 488 nm and
emission at 525 nm by the FACSCanto II flow cytometer (BD
Biosciences).
Cells treatment of DAPI staining was similar to that of AO/EB
staining. After washing with PBS three times, MCF‐7 cells were stained
with 1 ml of 300 nM DAPI in PBS for 20–30 min at 37°C. Subsequently,
the cells were collected, washed twice by PBS, and resuspended in 1 ml
of PBS. Next, 5 µl of cell suspension was transferred to a sterile glass
slide and covered with a coverslip. The morphology of cells was in-
vestigated using a fluorescence microscope.
|
4.2.8
Western blot
MCF‐7 cells were seeded (1 × 106 cells/dish) in 60‐mm culture
dishes. After overnight incubation, the cells were treated with in-
dicated concentrations of compound 5c for 24 h. The cells were lysed
with an ice‐cold radioimmunoprecipitation assay containing 1 mM
phenylmethylsulfonyl fluoride for 30 min. The mixture was cen-
trifuged at 1.2 × 105 r/min at 4°C for 10 min. Then, loading buffer
was added to the supernatant and the samples were treated in
boiling water for 5 min, and 5 μl of protein samples were separated
by 12% sodium dodecyl sulfate‐polyacrylamide gel electrophoresis
and then transferred to a polyvinylidene fluoride (PVDF) membrane.
The PVDF membrane was blocked by 5% nonfat powdered milk and
washed three times with Tris‐buffered saline and Tween 20 (TBST)
(10 min/time). After overnight incubation at 4°C in the primary an-
tibody diluted with TBST, the membrane was washed with TBST.
A secondary antibody, containing 5% nonfat powdered milk was
added, the membrane was incubated for 1 h at room temperature,
and then the membrane was washed with PBST (phosphate‐buffered
saline with Tween 20) again. The relative protein band was detected
using enhanced chemiluminescence Western blot analysis substrate
with Chemiluminescent Imaging System (Tanon).
|
4.2.4
Apoptosis assay (annexin V‐FITC/PI)
MCF‐7 cells were harvested (2 × 104 cells/well) and incubated in
six‐well plates overnight. After treatment with indicated concentra-
tions of compound 5c for 24 h, MCF‐7 cells were washed twice with
PBS and collected by centrifugation. The resulting cells were then
resuspended in 1× incubation buffer and incubated with annexin
V‐FITC/PI kit (Dalian Meilun Biotech) in the dark at room tempera-
ture for 15 min. The fluorescent signal was detected by the FACS-
Canto II flow cytometer (BD Biosciences).
|
4.2.5
Cell cycle analysis
After being harvested and seeded in six‐well plates (2 × 104 cells/well),
MCF‐7 cells were treated with indicated concentrations of compound
5c for 24 h. The cells were washed with PBS, collected by centrifugation
at 1000 r/min for 5 min, and fixed in 70% ethanol at 4°C overnight.
Next, the cells were centrifuged (1000 r/min, 3 min), washed with PBS,

HE ET AL.
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|
|
4.2.9
Statistical analysis
[17] R. Mandal, J. Compte, I. Kostova, S. Becker, K. Strebhardt, Biochim.
Biophys. Acta, Rev. Cancer 2020, 1873, 188357.
[18] R. E. D. Angel, S. M. Smith, R. D. Glickman, S. N. Perkins,
S. D. Hursting, Nutr. Cancer 2010, 62, 1074.
The data are presented as the mean standard deviation. No sta-
tistical method was used for data analysis.
[19] L. S. Bergamin, F. Figueiró, F. Dietrich, F. D. M. Manica, E. C.
Filippi‐Chiela, F. B. Mendes, E. H. F. Jandrey, D. V. Lopes,
F. H. Oliveira, I. C. Nascimento, H. Ulrich, A. M. O. Battastini,
Eur. J. Pharmacol. 2017, 811, 268.
|
4.3
Docking methodology
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CONFLICTS OF INTEREST
The authors declared that there are no conflicts of interest.
ORCID
Jiamei Lei
https://orcid.org/0000-0001-8224-5682
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