Exploring the Changes of Peroxisomal Polarity in the Liver of Mice with Nonalcoholic Fatty Liver Disease

Article abstract of DOI:10.1021/acs.analchem.1c01776

Peroxisome proliferator-activated receptor alpha (PPAR-a) is a crucial nuclear transcription regulator of lipid metabolism, which is closely associated with the initiation and development of nonalcoholic fatty liver disease (NAFLD). Because PPAR-a can directly decide the level of peroxisomal metabolic enzymes, its changes might directly cause variations in peroxisomal polarity. Therefore, we developed a new two-photon fluorescence imaging probe, PX-P, in which the triphenylamine and cyanide moieties can real-time sense peroxisomal polarity changes. Using PX-P, we observed a prominent decrease in the peroxisomal polarity in the liver of mice with NAFLD for the first time. More importantly, we discovered that intracellular excessive peroxynitrite (ONOO^-) and hydrogen peroxide (H₂O₂) underwent nitrification and oxidation, respectively, with various sites of PPAR-a. Interestingly, the key site of PPAR-a was nitrated by a low concentration of ONOO^-rather than being oxidized by the high level of H₂O₂. These drastically reduced the activity of PPAR-a, accelerating the occurrence of NAFLD. Moreover, through activating PPARs with pioglitazone, peroxisomal polarity markedly increased compared with that of NAFLD. Altogether, our work presents a new approach for the early diagnosis of NAFLD and identifies potential therapeutic targets.

Full text of DOI:10.1021/acs.analchem.1c01776

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Article
Exploring the Changes of Peroxisomal Polarity in the Liver of Mice
with Nonalcoholic Fatty Liver Disease
Yongqing Zhou, Chuanchen Wu, Xin Wang, Ping Li,* Nannan Fan, Wei Zhang, Zhenzhen Liu,
Wen Zhang, and Bo Tang*
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ABSTRACT: Peroxisome proliferator-activated receptor alpha (PPAR-a) is
a crucial nuclear transcription regulator of lipid metabolism, which is closely
associated with the initiation and development of nonalcoholic fatty liver
disease (NAFLD). Because PPAR-a can directly decide the level of
peroxisomal metabolic enzymes, its changes might directly cause variations
in peroxisomal polarity. Therefore, we developed a new two-photon
fluorescence imaging probe, PX-P, in which the triphenylamine and cyanide
moieties can real-time sense peroxisomal polarity changes. Using PX-P, we
observed a prominent decrease in the peroxisomal polarity in the liver of
mice with NAFLD for the first time. More importantly, we discovered that
intracellular excessive peroxynitrite (ONOO−) and hydrogen peroxide
(H2O2) underwent nitrification and oxidation, respectively, with various
sites of PPAR-a. Interestingly, the key site of PPAR-a was nitrated by a low
concentration of ONOO− rather than being oxidized by the high level of
H2O2. These drastically reduced the activity of PPAR-a, accelerating the occurrence of NAFLD. Moreover, through activating
PPARs with pioglitazone, peroxisomal polarity markedly increased compared with that of NAFLD. Altogether, our work presents a
new approach for the early diagnosis of NAFLD and identifies potential therapeutic targets.
oxygen species/nitrogen (ROS/N).12,13 Then, these ROS/N
INTRODUCTION
■
will cause oxidative damage to lipids, proteins, and DNA,
Nonalcoholic fatty liver disease (NAFLD) is a kind of
eventually accelerating the formation of NAFLD.14,15 Hitherto,
metabolic syndrome, characterized by hepatocellular steatosis
and lipid deposition.1,2 Because the specific mechanism of
the existing studies indicate that PPAR-a has been used as a
key targeting protein for the NAFLD treatment.16,17 However,
NAFLD is undefined, there is no effective treatment strategy at
the moment.3 Continuous progression and exacerbation of
under intracellular oxidative stress, the exact mechanism of
depressing the protein content of PPAR-a in the initiation and
progression of NAFLD is unclear. Furthermore, it is
remarkable that abnormal lipid metabolism directly impacted
the change of the peroxisomal microenvironment,18 especially
its polarity. Conspicuously, monitoring peroxisomal polarity
behaviors is fundamental to warn about the beginning of
NAFLD formation and investigate the reasons for the
dysfunction of PPAR-a, and even unravel the specific
mechanisms of NAFLD, which are conducive to provide the
corresponding therapeutic targets and strategies to improve the
cure rate of NAFLD.
NAFLD will greatly accelerate the incidence of liver hepatitis,
cirrhosis, and hepatocellular carcinoma.4−6 Hence, the timely
and accurate diagnosis of NAFLD can effectively contribute to
its treatment and save lives. The present commonly used
diagnostic techniques of NAFLD mainly rely on conventional
medical methods, which mainly include liver biopsy,7 serum
biochemistry, and ultrasonic examination.8 However, these
strategies are mostly applied in the late stage of NAFLD. Based
on these facts, we can say that there is an urgent need to
develop a straightforward visualization method that will
effectively conduce to the early intervention and improvement
of the cure rate of NAFLD.
Peroxisome proliferator-activated receptor alpha (PPAR-a)
plays a critical role in regulating hepatic metabolizing
enzymes,9 especially peroxisomal metabolizing enzymes, such
as acyl-CoA oxidase 1 and L-bifunctional enzyme.10 These will
be deposited in hepatocytes when lipids exceed the
peroxisomal β-oxidation capacity.11 Subsequently, excessive
lipid accumulation gives rise to an explosion of free reactive
Received: April 26, 2021
Accepted: June 15, 2021
Published: June 30, 2021
https://doi.org/10.1021/acs.analchem.1c01776
© 2021 American Chemical Society
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Scheme 1. Structure of PX-P and the Proposed Response Mechanism to Polarity
Currently, fluorescence imaging techniques are serving as a
superior approach to monitor microenvironmental changes in
cells and in vivo.19−22 In particular, two-photon fluorescence
probes have attracted great interests in the field of disease
diagnosis in view of their momentous merits, including high
spatial resolution and deep penetration.23−25 So far, some two-
photon fluorescent probes have been developed to detect the
polarity alteration of subcellular organelles.26−29 Unfortunately,
two-photon fluorescent probes for monitoring peroxisomal
polarity in the liver of mice with NAFLD are still lacking.
On the basis of these circumstances, a novel molecular
fluorescent probe (PX-P) with a donor−π−acceptor structure
was ingeniously constructed to detect peroxisomal polarity in
vivo. In the probe skeleton (Scheme 1), the triphenylamine
group was used as the donor (D), the furan moiety as the π
bridge, and the cyanide unit as the electron acceptor (A),
causing the excited-state twisted intramolecular charge transfer
(TICT). The D and A were linked with a furan segment,
forming a tremendous conjugate system, which could
effectively shift the fluorescence emission to the red region.
Moreover, the SKL peptide chain was introduced as the
peroxisomal-targeted moiety because it could tightly bind to
the Pex5p receptor on the surface of the peroxisomal
membrane and then was transported into peroxisomes.30 PX-
P displayed a weak fluorescence in a high polar environment
because the excited-state energy can be dissipated by
nonradiative relaxation pathways, while a strong fluorescence
emission was obtained in a nonpolar medium. We proved that
PX-P could selectively accumulate in peroxisomes and indicate
the change of polarity. Moreover, PX-P was successfully
applied in exploring the relationship between peroxisomal
polarities and NAFLD. Furthermore, through proteomic
analysis, we first revealed that the underlying mechanism of
the PPAR-a protein is associated with the occurrence of
NAFLD.
solution was removed and dimethyl sulfoxide (DMSO) (150
μL) was added to each well to dissolve the dark blue formazan
crystals. Absorbance was measured at 490 nm in a Triturus
microplate reader.
Cell Imaging. These cells were detached and replanted on
glass-bottomed dishes before HL-7702 cell imaging. For
monitoring the polarity in living cells, these cells were first
exposed to oleic acid (OA) (5.0 μM, 30 min); then, the cells
were treated with PX-P (5.0 μM, 30 min) before being washed
three times with PBS. For the control assay, cells were treated
with PX-P (5.0 μM, 30 min). To study PX-P specifically
targeted in peroxisomes, first, the cells were stained with
Peroxisome-GFP (3.0 μL, 108 particle·mL−1, 26 h) or other
commercial dyes (100 nM MitoTracker Deep Red, 100 nM
LysoTracker Deep Red, 50 nM ER-Tracker Red, 50 nM Golgi-
Track Red) and then treated with PX-P (5.0 μM, 30 min)
before being washed three times with PBS. The time-
dependent fluorescence imaging of the HL-7702 cells
incubated with OA (5.0 μM) for 30 min was performed and
then the cells were treated with PX-P (5.0 μM) for 30 min. To
verify peroxisomal polarity regulated by PPAR-a using gene-
silencing PPAR-a with small interfering RNA (siRNA) in HL-
7702 cells, we designed three types of siRNAs for inhibiting
the expression of PPAR-a. These cells were stably transfected
with the three types of siRNAs obtained from Shanghai
GenePharma Co., Ltd.
Cell Breaking Extraction. 1.0 × 105 cells were cultured on
a 6-well plate overnight. Culture medium was discarded and
cells were rinsed three times with PBS. For experimental cells,
these were first exposed to OA (5.0 μM, 30 min) or three types
of siRNAs. Then, every cell was treated with 200 μL of native
lysis buffer (2 min) before being washed three times with PBS.
These cells were fully lysed and centrifuged at 12,000 rpm.
Ultimately, the supernatant of the cell-breaking solvents was
taken for subsequent experiments. For the control assay, the
cells were treated with 200 μL of native lysis buffer (2 min)
before being washed three times with PBS.
Ethics Statement. All animal experiments were strictly
performed in accordance with the relevant Laws and
Guidelines issued and approved by the Ethics Committee of
Shandong Normal University for the Care and Use of
Laboratory Animals.
In Vivo Toxicity Test. C57 mice (female, 5 weeks old, 16−
18 g) were purchased from the Laboratory Animal Center of
Shandong University (Jinan, China). C57 mice were randomly
divided into three groups, one control group and two
experimental groups. The two experimental groups were
treated with different concentrations of PX-P (100 μL, 10−4
and 2.0 × 10−4 M) by intravenous administration, respectively.
The control group was treated with 100 μL of saline. The body
EXPERIMENTAL SECTION
■
Cytotoxicity Assay. The cytotoxicity was measured by the
3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide
(MTT) assay. HL-7702 cells were seeded in a 96-well plate
at a concentration of 1 × 105 cell/well in 100 μL of Dulbecco’s
modified Eagle’s medium with 10% fetal bovine serum, 1%
penicillin, and 1% streptomycin and maintained at 37 °C in a
5% CO2 incubator for 12 h. Then, the cells were exposed to
different concentrations of PX-P (1 × 10−4, 1 × 10−5, 1 × 10−6,
1 × 10−7, 1 × 10−8, and 1 × 10−9 M) for 24 h. The total
volume of 96-well plates is 200 μL. These cells were washed
with 37 °C phosphate-buffered saline (PBS), and MTT
solution (5 mg mL−1, 20 μL) was added to each well and
continuously incubated for 4 h at 37 °C. After 4 h, the MTT
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Scheme 2. Synthetic Route for PX-P
weight changes of mice were measured every day. Moreover,
all the rats showed no obvious symptoms for 2 weeks, n = 5.
Mouse Models with NAFLD. C57 mice were randomly
divided into three groups, a control group, an experimental
group, and a treatment group. For the experimental group and
the treatment group, the mice were fed with high-fat forage (60
kcal % fat) for 7 weeks. Also, the treatment group mice were
treated with the same dose of pioglitazone (4 mg/kg) by
intragastric administration every 2 days after 2 weeks. Normal
mice were fed with normal diet every day. Additionally, the
body weights of all these mice were measured every 3 days.
In Vivo Fluorescence Imaging of NAFLD Mice. The
NAFLD mice model was successful established after 7 weeks.
C57 mice were fasted for 12 h to avoid possible food
fluorescence interference with the dye fluorescence before
imaging. Experimental and treatment groups were treated with
PX-P (100 μL, 10−4 M) by intravenous administration. The
control group was treated with 100 μL of saline. In vivo
imaging of PX-P was done in livers of mice with NAFLD after
30 min. λex = 480 nm and λex = 600 20 nm. In vivo imaging
of PX-P was performed in livers of mice with NAFLD after 30
min. λex = 800 nm, λex = 550−650 nm, and n = 5.
mmol, 140 mg), K2CO3 aqueous solution (2 M, 1.2 mL), and
Pd(PPh3)4 (0.1 mmol, 116 mg) was dissolved in 10 mL 1,4-
dioxane (1,4-Dio). Then, the reaction solution was stirred at
90 °C for 8.0 h. After that, the solvent was cooled to room
temperature, then 100 mL of ice water was added, and the
mixture was extracted three times with CH2Cl2. The organic
phase was separated, washed with brine, and dried with
anhydrous Na2SO4. The solvent in the collected filtrate was
evaporated, and the product was purified by silica gel column
chromatography (CH2Cl2/MeOH = 10:1, v/v) to give pure
compound 1 as a white solid (236 mg, 70%). 1H NMR
(DMSO-d6, 400 MHz): δ 9.553 (s, 1H), 7.748 (d, J = 8.8 Hz,
2H), 7.643 (d, J = 2.0 Hz, 1H), 7.736 (t, J = 7.6 Hz, 1H), 6.91
(m, 7H), 7.13 (d, J = 8.4 Hz, 2H). 13C NMR (DMSO-d6, 100
MHz): δ 105.015, 121.886, 122.056, 124.637, 125.532,
130.255, 146.95, 149.145, 151.798, 159.106, 177.640. HRMS
(ESI): calcd for C23H17NO2, [M + H]+ = 340.1339; found,
340.1340.
Compound 1 (1.0 mmol, 340 mg) was dissolved in 3.0 mL
of acetonitrile, then added with cyanoacetic acid (1.0 mmol, 85
mg) and one drop of piperidine, and stirred at room
temperature for 4 h. The solvent was removed, and the
resulting crude product was purified by column chromatog-
raphy (CH2Cl2/MeOH = 8:1, v/v) to give pure compound 2
Immunohistochemistry and Biochemical Measure-
ments. Fresh liver organs were quickly removed from these
mice after euthanasia. Then, a part of the liver tissue was fixed
in 10% formalin and paraffin. Paraffin-embedded liver tissues
were used for H&E as well as oil red O staining. The other part
was prepared into a liver tissue homogenate for the detection
of other markers in the process of NAFLD and kept in a
freezer at −80 °C.
1
(325 mg, 80%). H NMR (DMSO-d6, 400 MHz): δ 9.403 (s,
1H), 2.56 (d, J = 9.2 Hz, 3H), 7.356 (t, J = 8.0 Hz, 4H), 7.204
(d, J = 3.2 Hz, 1H), 7.119 (m, 7H), 6.987 (d, J = 8.4 Hz, 2H).
13C NMR (DMSO-d6, 100 MHz): δ 108.123, 119.854,
121.013, 122.482, 123.141, 124.436, 125.284, 126.136,
130.282, 133.618, 146.985, 148.233, 148.633, 156.433,
164.597. HRMS (ESI): calcd for C26H18N2O3, [M − H]− =
405.1227; found, 405.1284.
Proteomic Analysis. The PPAR-a protein (1.0 mg/mL)
was incubated at 37 °C in buffer (10 mM sodium phosphate,
pH 7.4, 0.1 mM diethylenetriamine pentaacetate) for 2 h
supplemented with ONOO− (100 μM) or H2O2 (2.0 mM).
Then, the PPAR-a protein was subjected to trypsin digestion,
and the peptides were isolated from the hydrolysate by solid-
phase extraction on a C-18 column. Proteomic analysis was
then performed through liquid chromatography with tandem
mass spectrometry (LC−MS/MS).
PX-P was provided by Sangon Biotech. Theoretical calcd for
C41H46N6O7, [M] = 734.34; found, [M] = 734.35.
The final structures of compounds 2, 3, and PX-P were well
1
characterized by HNMR, 13C NMR, and high-resolution MS
(HRMS). PX-P was well characterized by HRMS and high-
performance liquid chromatography (HPLC).
RESULTS AND DISCUSSION
Optical Properties of PX-P. First, the photophysical
properties of PX-P were examined in various polarity solvents.
Synthesis of PX-P. The PX-P was synthesized, as
illustrated in Scheme 2. First, the mixture of 4-bromotriphenyl-
amine (1.0 mmol, 324 mg), 5-formylfuran-2-boronic acid (1.0
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Figure 1. Optical properties of PX-P. (a) Fluorescence spectra of PX-P (5.0 μM) in various 1,4-Dio−H2O solvent systems. (b) Linear relationship
between fluorescence intensities of PX-P (5.0 μM) and Δf in various 1,4-Dio−H2O solvent systems. (c) Fluorescence time (τ) of PX-P (5.0 μM) in
various 1,4-Dio−H2O solvent systems. Linear relationship between τ of PX-P (5.0 μM) and Δf in various 1,4-Dio−H2O solvent systems. λex = 460
nm and λem = 586 nm.
As could be seen from Figure S1a, the main absorption peak of
PX-P was concentrated at about 460 nm. However, PX-P
showed significant fluorescence in some solvents with lower
polarity and higher hydrophobicity (Figure S1b). Moreover,
the maximum emission peak exhibited an obvious positive
solvatochromism from 1,4-Dio to MeOH with a 65 nm
bathochromic shift, an obvious decrease of fluorescence
intensity accordingly. Compared with other solvents, PX-P
had a higher quantum yield in 1,4-Dio (Table S1). The
fluorescence intensities of PX-P in two solvents with similar
viscosities and different polarities were measured. The
fluorescent signal of PX-P in tetrahydrofuran (THF) was
prominently stronger than that in MeOH (Figure S2). This
proves that PX-P is not sensitive to viscosity.
To reveal the polarity determination ability of PX-P, the
fluorescence responses of PX-P to various polarity mixtures
were estimated in a class of 1,4-dio−water systems with
different ratios. The Lippert-Mataga polarity parameter (Δf)
was used to represent the solvent polarity.31 With the increase
of the ratio of 1,4-Dio, the Δf of mixed media gradually
decreased. As illustrated by Figures S3 and 1a, both the
maximum absorption peak and the fluorescence peak of PX-P
increased dramatically when the solvent polarity decreased.
Notably, there was an excellent linear relationship (R2 = 0.994)
between the fluorescence intensity of PX-P and Δf (Figure 1b).
Additionally, the fluorescence lifetime (τ) is also a crucial index
to investigate the performance of polarity probes.32 Hence, the
τ of PX-P in different polarity media were measured. The τ of
PX-P gradually increased along with decreasing media polarity
(Figure 1c). As illustrated in Figure 1d, the favorable linearity
relationship (R2 = 0.999) between the τ of PX-P and Δf could
be obtained by the corresponding equation.33
All of these experimental results manifest that the photo-
physical properties of PX-P are strongly dependent on the
solvent polarity. Next, we interpreted the strong solvatochrom-
ism mechanism of PX-P by density functional theory
calculations.34 The process of intramolecular charge transfer
(ICT) occurs from the triphenylamine to cyanide unit when
photoexcitation transfer occurs from the S0 to S1 state of PX-P
(Figure S4). Because PX-P possessed a freely rotatable single
bond between the donor and acceptor moieties, the activation
of the ICT process is likely accompanied by a critical molecular
geometry change, resulting in the formation of a TICT state.
Hence, the solvatochromism mechanism and the large Stokes
shift of PX-P both attribute to the TICT effect. Moreover, the
fluorescence intensity of PX-P markedly decreased in the high
polar environments, owing to the rapid consumption of the
energy of the TICT state through nonradiative relaxation
pathways.
To test the specificity of PX-P for polarity, we examined
whether PX-P possessed the essential ability to compete
against the interferences of intracellular relevant bioactive
molecules, such as ROS, RNS, reactive sulfur species, metal
ions, and enzymes. As shown in Figure S5, the fluorescence of
PX-P could be ignored except in 1,4--Dio, which produced a
remarkable fluorescent signal because of the lower-polarity
environments. Therefore, PX-P did not respond to the
bioactive species in complex biological systems and had a
high specificity ability of environmental polarity. To study the
stability of PX-P, we tested the fluorescent signal of PX-P
under different pH (5.0−9.0) conditions. As displayed in
Figure S6, the fluorescence intensity of PX-P remains
unchanged in PBS, indicating that the fluorescent signal of
PX-P is independent of pH. To investigate the photostability of
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Figure 2. Fluorescence images of PX-P and Peroxisome-GFP in HL-7702 cells. The HL-7702 cells were stained with Peroxisome-GFP (2.0 μL, 108
particle mL−1) for 26 h and then with PX-P (5.0 μM) for 30 min. (a) Fluorescence image of Peroxisome-GFP (λex = 488 nm and λem = 500−550
nm). (b) Fluorescence image of PX-P (λex = 488 nm and λem = 550−650 nm). (c) Overlay imaging of (a,b). (d) Intensity profile of the blue line in
image (c). Scale bar = 50 μm.
PX-P, we evaluated the fluorescence variations of PX-P in 1,4-
Dio and PBS for 1.0 h, respectively. The fluorescent signals of
PX-P remained stable in two different solvents (Figure S7).
Overall, the above results show that PX-P has the advantages of
high selectivity, specificity, and photostability toward environ-
mental polarity.
The PX-P was constructed as a D−π−A system, which had
distinctive TICT features. Therefore, PX-P was expected to
exhibit a higher two-photon absorption (2PA) cross section in
different solvents. Using rhodamine 6G as the standard, the
2PA cross section of PX-P was obtained in different solvents by
a two-photon fluorescence microscope.35 Under an excitation
of 800 nm, compared with other solvents, PX-P had the
maximum two-photon cross-sectional area value of 26.6 GM in
MeOH (Figure S8). Therefore, PX-P can be used as a good
two-photon imaging tool for studying the polarity under
complicated biological conditions.
Peroxisomal Localization of PX-P. Before applying PX-P
to biological imaging, we evaluated its toxicity in HL-7702 cells
by using the MTT assay.36 The IC50 value was calculated to be
81.3 M using the corresponding formula,37 demonstrating that
PX-P has low cytotoxicity (Figure S9). To test the in vivo
toxicity of PX-P, C57 mice were injected with various
concentrations of PX-P for 2 weeks.38 These mice did not
show any obvious symptoms, such as death, calvities, anorexia,
or inflammation. Moreover, the average body weight of the
two experimental mice was close to that of normal mice
(Figure S10), indicating that PX-P possesses low toxicity in
vivo. Therefore, PX-P has promising application potential in
biological experiments. To investigate the subcellular distribu-
tion of PX-P in living cells, we employed the commercial
staining dye (Peroxisome-GFP) with PX-P for co-localization
imaging in HL-7702 cells. The same patterns were observed
solely by using two dyes (Figure 2), with a decent co-
localization coefficient of 0.89. These data demonstrate that
PX-P can entirely accumulate in peroxisomes. To further prove
that PX-P accumulated only in peroxisomes rather than in
other organelles, the co-localization experiments of PX-P with
other organelle commercial dyes were also further carried out.
It could be seen from these figures that PX-P rarely appeared
in the golgi apparatus, endoplasmic reticula, mitochondria, or
lysosomes (Figure S11). Altogether, these results testify that
PX-P have a prominent peroxisomal-targeting ability.
Fluorescence Imaging of Peroxisomal Polarity in HL-
7702 Cells. Next, we intended to detect the local polarity in
living cells with PX-P. Previous reports have shown that the
organelles of cancer cells have a lower polarity environment
than those of normal cells, such as mitochondria and golgi
apparatus.39,40 Herein, we examined the peroxisomal polarity
in normal and cancer cells. As shown in Figure S12, compared
to HL-7702 cells, SMMC-7721 cells elicited a strong
fluorescence, suggesting that the peroxisomes of liver cancer
cells possessed a lower-polarity environment. This correspond-
ing result highlights the fact that PX-P can real-time track the
variation of peroxisomal polarity at the cellular level. We then
used PX-P to monitor polarity when the amount of
intracellular lipid level increased significantly in the steatotic
hepatocyte model. We chose OA as a stimulus to incubate HL-
7702 cells to induce the NAFLD cell model, which represented
the steatotic cell type.41 The stimulated HL-7702 cells gave out
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Figure 3. Two-photon fluorescence images of PX-P in NAFLD cells. (a−c) Confocal fluorescence images of HL-7702 cells incubated with PX-P
(5.0 μM) for 30 min. (e−g) Confocal fluorescence images of cells incubated with PX-P (5.0 μM) after treatment with OA (5.0 μM) for 30 min. (d)
Relative fluorescence intensity and (h) relative PPAR-a activity of the control cells. λex = 800 nm, λem = 550−650 nm, and scale bar = 20 μm. These
values are the mean s.d. ★★★P < 0.001, for n = 5.
obviously brighter fluorescent signals (about 2.7 times)
compared with the control cells (Figure S13). These data
confirm that peroxisomal polarity obviously decreases during
NAFLD. Subsequently, we also performed an experiment to
observe the fluorescence change of PX-P with time in the
NAFLD cell model. The fluorescence intensity of PX-P
gradually increased within 40 min (Figure S14), demonstrating
that the peroxisomal polarity gradually attenuated with the
process of NAFLD. Therefore, through these experimental
results, the established imaging approach of PX-P can
successfully monitor the decrease of peroxisomal polarity in
the NAFLD cell model.
Because of the fascinating two-photon properties of PX-P,
we next evaluated whether PX-P was suitable for the two-
photon fluorescence imaging peroxisomal polarity variations in
the NAFLD cell model. Similarly, OA was also selected as an
inducer to induce lipid accumulation in HL-7702 cells. Figure
3 shows that HL-7702 cells stimulated by OA emitted a
pronounced fluorescence (2.8 times) compared with the
control cells, indicating that peroxisomal polarity reduced
significantly during abnormal lipid metabolism. Hence, these
persuasive data confirm that PX-P is appropriate for two-
photon imaging peroxisomal polarity changes. Moreover, the
PX-P-based polarity imaging method can be used to
distinguish cells with abnormal lipid metabolism from normal
cells by polarity change. Consequently, we also tested the key
protein expression level of PPAR-a in HL-7702 cells under OA
stimulation. These experimental results prove that the level of
PPAR-a in NAFLD cells was significantly lower than that in
normal cells (Figure 3h), manifesting that PPAR-a is closely
involved in the pathogenesis of NAFLD.
by excess lipid accumulation in the mouse liver,42,43 C57 mice
were fed with a high-fat diet (HFD) to establish the model of
NAFLD.44 As reported earlier, pioglitazone is a highly selective
agonist of PPARs, which can activate PPARs to regulate lipid
metabolism, thereby reducing lipid and triglyceride contents in
hepatocytes.45,46 Here, pioglitazone was selected to relieve the
mice with NAFLD (Figure 4A). The hematoxylin and eosin
and oil red O staining were used to stain the hepatic tissue
after C57 mice were fed with HFD for 7 weeks.47,48 According
to the abovementioned stained results, the degree of hepatic
steatosis and fat deposition in mice with NAFLD are
apparently higher than those in normal mice, as well as
slightly higher than those in pioglitazone-treated mice (Figure
4B,C), suggesting that pioglitazone can serve as an effective
drug to alleviate NAFLD. Moreover, mice fed with HFD got
more body weight compared with other groups (Figure S15).
At the same time, we also tested the NAFLD marker levels by
using commercial kits, alanine aminotransferase (ALT) and
gamma-glutamyltransferase (GGT).49,50 These consequences
illuminated that the levels of ALT/GGT in the livers of mice
with NAFLD were significantly increased than those of normal
mice and slightly higher than those of pioglitazone-treated
mice (Figure 4D,E). Furthermore, malondialdehyde (MDA)
has been commonly used as a biomarker of lipid peroxidation51
because accumulating lipids generated more oxidative stress
and damage. Here, we next evaluated the concentrations of
MDA in fresh liver extracts. These results showed that the
MDA levels of fresh liver extracts were found to be markedly
enhanced in mice with NAFLD in comparison to the treatment
and control mice (Figure 4F). Collectively, all these
experimental data imply that the NAFLD mice model was
successfully established.
Fluorescence Imaging of PX-P in Livers of Mice with
NAFLD. Inspired by these reputable performances of PX-P in
living cells, we next investigated whether it could be used to
distinguish the differences in liver polarity between NAFLD
mice and normal mice. Considering that NAFLD is triggered
Next, we sought to use PX-P to study the underlying
relationships between peroxisomal polarity and mice with
NAFLD. As illustrated in Figure 5A, the livers of mice with
NAFLD emitted considerable fluorescent signals than those of
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Figure 4. Establishment and analysis of the mice model with NAFLD. (A) Establishment process of the mice model with NAFLD. (B) H&E and
(C) oil red “O” staining of liver tissues. (D) Relative ALT activity, (E) relative GGT activity, and (F) relative MDA activity to the control cells. a1,
b1: normal mice; a2, b2: NAFLD mice; and a3, b3: NAFLD mice + pioglitazone. Scale bar: 100 μm. These values are the mean s.d. ★★★P <
0.001, for n = 5.
normal mice (at about 3.3 times), proving that the liver
polarity of NAFLD mice was lower than those of normal mice
and pioglitazone-treated mice. These phenomena further verify
that fluorescence enhancements in the liver of mice with
NAFLD are induced by decreased peroxisomal polarity.
To real-time and in situ measure the polarity change in the
deep liver tissues, we used a two-photon fluorescence imaging
technique to monitor peroxisomal polarity. As shown in Figure
5B, fluorescence in the liver of mice with NAFLD was
significantly enhanced than that of the normal group (at about
3.0 times), indicating that peroxisomal polarity decreases in the
liver of mice with NAFLD. Compared with NAFLD mice, the
liver of pioglitazone-treated mice showed a weaker fluores-
cence, demonstrating that the peroxisomal polarity of
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Figure 5. In vivo fluorescence imaging in livers of mice. (A) (a) Liver fluorescence imaging of PX-P (100 μL, 10−4 M) in vivo: a1: normal mice +
PX-P, a2: NAFLD mice + pioglitazone + PX-P, a3: NAFLD mice + PX-P; (b) relative fluorescence intensity and (c) relative PPAR-a content of the
control mice, respectively. λex = 480 nm and λem = 600 20 nm. (B) Two-photon imaging of PX-P in livers of mice with NAFLD: (b1): normal
mice + PX-P, (b2): NAFLD mice + PX-P, (b3): NAFLD mice + pioglitazone + PX-P, and (b4): relative fluorescence intensity of control mice. λex
= 800 nm and λem = 550−650 nm. Scale bar: 200 μm, Z = 500 μm. These values are mean s.d. ★★P < 0.01, ★★★P < 0.001, for n = 5.
pioglitazone-treated mice was higher than that of NAFLD
mice. These two-photon imaging results were consistent with
the in vivo fluorescence imaging results. Therefore, the imaging
detection method based on PX-P can be used to distinguish
NAFLD mice from normal mice by monitoring the polarity
change.
formation of NAFLD. Among RNS, peroxynitrite (ONOO−)
is a stronger adduct of nitric oxide and superoxide anion
radicals. Hydrogen peroxide (H2O2), as an intracellular ROS
present in high concentrations, reacts directly with amino acids
containing a sulfhydryl group and inactivates the related
proteins. In particular, the concentrations of ONOO− and
H2O2 were found to be significantly increased with the
progression of NAFLD.53,54 We first measured the levels of
PPAR-a in the presence of excessive ONOO− or H2O2 in living
HL-7702 cells. As results displayed, these PPAR-a contents
reduced significantly when cells were added a mass of ONOO−
or H2O2 (Figure S18). We next explored the definite
interaction of decreasing PPAR-a protein activity with the
excessive ONOO− or H2O2 from the perspective of oxidative
stress. To fully understand the product profile, we then
performed the proteomic analysis by LC−MS/MS to
investigate the corresponding reaction residues. The results
found that six sites in PPAR-a protein were nitrated and
oxidized by the overexpression of ONOO− and H2O2, Y155,
Y311, Y314, Y334, H242, and C248, respectively (Figure 6).
Meanwhile, Y314 is one of the key amino acids for protein
activation which plays a critical role in lipid metabolism.
Hence, these results provide strong evidence that the
intracellular excess ONOO− caused structural and functional
damage to the PPAR-a protein, leading to the downregulation
of the related metabolic enzymes, eventually accelerating the
formation of NAFLD.
Key Role of PPAR-a during the NAFLD Process. After
the in vivo imaging experiments, we further measured the key
protein concentration of PPAR-a in the liver extracts by a
specific kit. The results illuminated that the PPAR-a level in
NAFLD mice was significantly lower than that of normal and
treatment mice groups (Figure 5c). These findings are
consistent with the previously reported significant reductions
in the PPAR-a level during NAFLD.52 Therefore, using gene-
silencing PPAR-a with siRNA in HL-7702 cells, we next
verified whether the decrease of the PPAR-a level led to
peroxisomal polarity alterations. We first screened out this
sequence with the best inhibitory effect on the expression of
PPAR-a. As illustrated in Figure S16, cells with sequence
siRNA-1234 possess the lowest expression of PPAR-a,
manifesting that it has a good inhibition effect for intracellular
PPAR-a. Then, we observed a significant increase in the
fluorescence intensity and a distinct reduction in the level of
PPAR-a from silencing cells by using this sequence relative to
the control cells (Figure S17), indicating that the reduced
peroxisomal polarity might result from the low level of
intracellular PPAR-a protein.
Oxidation Modifications of PPAR-a during the NAFLD
Process. As previously reported, the level of ROS/N increased
remarkably when there was lipid overload in hepatocytes,
following oxidative damage. This eventually accelerated the
NAFLD is an epidemic metabolic syndrome of hepatocel-
lular excessive fat accumulation, which has the potential to
gradually progress to hepatic fibrosis and end-stage liver
disease. Currently, the “two hit” theory has been commonly
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Figure 6. Proteomic analysis by LC−MS/MS. (a,b) Proteomic analysis of the reaction of PPAR-a (1.0 mg/mL) with H2O2 (2.0 mM) and (c,d)
proteomic analysis of the reaction of PPAR-a (1 mg/mL) with ONOO− (100 μM).
accepted in the pathophysiology of NAFLD, although this
pathogenetic mechanism is still controversial.12,55 Never-
theless, peroxisomal dysfunction, oxidative stress, and hepatic
steatosis were widely considered to be the main contrib-
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utors.56,57 Considering that peroxisomal dysfunction may
directly lead to the change in its microenvironment, in this
study, we developed a two-photon fluorescent probe, PX-P, for
the highly sensitive and selective detection of peroxisomal
polarity changes. Utilizing the PX-P, we first found that
peroxisomal polarity significantly decreased with the pro-
gression of NAFLD. Furthermore, we unprecedentedly
unmasked the concrete molecular mechanism of reducing
PPAR-a activity under oxidative stress. Therefore, these
findings further highlight the evidence that oxidative stress
causes lipid peroxidation and then increases cytokine
production and inflammation, ultimately resulting in NAFLD.
According to the environmental polarity and corresponding
fluorescence spectra of PX-P, the emission wavelength of PX-P
gradually red-shifts with the increase of solvent polarity.
Moreover, such properties, with a large Stokes shift of up to
180 nm, suggest that PX-P can effectively avoid the overlap of
absorption and emission spectra, as well as the self-absorption
phenomenon. Remarkably, PX-P possessed an excellent two-
photon cross-sectional area in different polarity solvents,
especially in THF solvents. Therefore, PX-P holds great
potential for being used as a peroxisome-specific two-photon
fluorescence tool for the dynamic monitoring of polarity in
cells and in vivo.
PX-P is not cytotoxic under the concentration of 81.3 M,
which is 16 times higher than that required for the peroxisomal
polarity imaging in HL-7702 cells in our experiments. At the
same time, PX-P also shows low toxicity in vivo. Thus, the
toxicity of PX-P is negligible in our biological experiments.
Additionally, PX-P exhibits a pronounced fluorescence
emission in silenced and OA-stimulated cells, as well as in
the liver of mice with NAFLD induced by a HFD. These
results indicate that PX-P not only can be useful for
peroxisomal polarity imaging in living cells but also for sensing
polarity in the livers of mice with NAFLD. These phenomena
confirm that peroxisomal polarity was altered during NAFLD.
To our knowledge, PPAR-a is also found to be closely
involved in steatosis. However, the specific mechanism of
decreased PPAR-a activity is still unclear in the pathogenesis of
NAFLD. To identify the pathogenesis of NAFLD, we next
meticulously investigated the reason for decreasing PPAR-a
protein activity in terms of oxidative stress. Oxidative
dysfunction has long been implicated in diverse biological
disorders in the metabolism process because the increased
concentration of ROS/N causes cumulative damage to many
protein structures. Recent research has illustrated that the
levels of both H2O2 and ONOO− increased significantly in the
initiation and development of NAFLD.53,54 To fully clarify the
mechanism of the dysfunction of PPAR-a under excessive
H2O2 and ONOO−, the corresponding products are analyzed
by the post-translational modification of PPAR-a by proteomic
analysis. According to LC−MS/MS data, the two sites (H242
and C248) of the PPAR-a protein were oxidized in the
presence of H2O2. Importantly, we found that the four tyrosine
residues (Y155, Y311, Y314, and Y334) of the PPAR-a protein
were nitrated under excessive ONOO−, including a key active
site Y314, which is the activation site of most agonists.58−60
Also, it was confirmed that decreasing PPAR-a protein activity
is related to site Y314. That is, this Y314 site will act as a
screen to novel PPAR-a agonists, drugs, and therapeutic targets
with NAFLD. The decrease of PPAR-a activity is caused by the
downregulation of the metabolic enzyme levels regulated by
PPAR-a, ultimately leading to a reduction in the peroxisomal
polarity. Altogether, these results clearly suggest that the
PPAR-a protein is inactivated by ONOO− under oxidative
stress conditions, thereby leading to NAFLD. In the meantime,
this further confirms that PPAR-a is a reliable drug target for
the treatment of NAFLD.
CONCLUSIONS
■
In summary, we reported a two-photon fluorescent probe
based on the D−π−A structure for peroxisomal polarity
imaging in living cells and in vivo for the first time. Using this
probe, we found that the liver peroxisomal polarity of NAFLD
mice was significantly decreased than that of normal mice and
treated mice, due to the reduction in the intracellular level of
PPAR-a. Furthermore, we uncovered that intracellular
excessive ONOO− caused the inactivation of PPAR-a.
Subsequently, the inactivated PPAR-a accelerated the
occurrence of NAFLD. Therefore, these also boost the new
roles of ONOO− in the pathogenesis of NAFLD. Altogether,
this PX-P-based polarity detection method can not only be
used in the early warning of the NAFLD but also contribute to
explicate the pathogenesis of NAFLD.
ASSOCIATED CONTENT
* Supporting Information
The Supporting Information is available free of charge at
https://pubs.acs.org/doi/10.1021/acs.analchem.1c01776.
■
sı
General materials and instruments, optical properties,
biocompatibility tests, cell imaging, weight of mice,
1
monitoring PPAR-a activity, and H NMR, 13C NMR,
and MS spectra (PDF)
AUTHOR INFORMATION
Corresponding Authors
■
Ping Li − College of Chemistry, Chemical Engineering and
Materials Science, Key Laboratory of Molecular and Nano
Probes, Ministry of Education, Collaborative Innovation
Center of Functionalized Probes for Chemical Imaging in
Universities of Shandong, Institute of Biomedical Science,
Shandong Normal University, Jinan 250014, People’s
Republic of China; orcid.org/0000-0002-2356-0962;
Email: lip@sdnu.edu.cn
Bo Tang − College of Chemistry, Chemical Engineering and
Materials Science, Key Laboratory of Molecular and Nano
Probes, Ministry of Education, Collaborative Innovation
Center of Functionalized Probes for Chemical Imaging in
Universities of Shandong, Institute of Biomedical Science,
Shandong Normal University, Jinan 250014, People’s
Republic of China; orcid.org/0000-0002-8712-7025;
Email: tangb@sdnu.edu.cn
Authors
Yongqing Zhou − College of Chemistry, Chemical Engineering
and Materials Science, Key Laboratory of Molecular and
Nano Probes, Ministry of Education, Collaborative
Innovation Center of Functionalized Probes for Chemical
Imaging in Universities of Shandong, Institute of Biomedical
Science, Shandong Normal University, Jinan 250014,
People’s Republic of China
Chuanchen Wu − College of Chemistry, Chemical Engineering
and Materials Science, Key Laboratory of Molecular and
Nano Probes, Ministry of Education, Collaborative
Innovation Center of Functionalized Probes for Chemical
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(5) Nakagawa, H.; Umemura, A.; Taniguchi, K.; Font-Burgada, J.;
Dhar, D.; Ogata, H.; Zhong, Z.; Valasek, M.; Seki, E.; HiDalgo, J.;
Koike, K.; Kaufman, R.; Karin, M. Cancer Cell 2014, 26, 331−343.
(6) Natarajan, Y.; Kramer, J.; Yu, X.; Li, L.; Thrift, A.; EI-Serag, B.;
Kanwai, H. Hepatology 2020, 72, 1242−1252.
Imaging in Universities of Shandong, Institute of Biomedical
Science, Shandong Normal University, Jinan 250014,
People’s Republic of China
Xin Wang − College of Chemistry, Chemical Engineering and
Materials Science, Key Laboratory of Molecular and Nano
Probes, Ministry of Education, Collaborative Innovation
Center of Functionalized Probes for Chemical Imaging in
Universities of Shandong, Institute of Biomedical Science,
Shandong Normal University, Jinan 250014, People’s
Republic of China
Nannan Fan − College of Chemistry, Chemical Engineering
and Materials Science, Key Laboratory of Molecular and
Nano Probes, Ministry of Education, Collaborative
Innovation Center of Functionalized Probes for Chemical
Imaging in Universities of Shandong, Institute of Biomedical
Science, Shandong Normal University, Jinan 250014,
People’s Republic of China
Wei Zhang − College of Chemistry, Chemical Engineering and
Materials Science, Key Laboratory of Molecular and Nano
Probes, Ministry of Education, Collaborative Innovation
Center of Functionalized Probes for Chemical Imaging in
Universities of Shandong, Institute of Biomedical Science,
Shandong Normal University, Jinan 250014, People’s
Republic of China
Zhenzhen Liu − College of Chemistry, Chemical Engineering
and Materials Science, Key Laboratory of Molecular and
Nano Probes, Ministry of Education, Collaborative
Innovation Center of Functionalized Probes for Chemical
Imaging in Universities of Shandong, Institute of Biomedical
Science, Shandong Normal University, Jinan 250014,
People’s Republic of China
Wen Zhang − College of Chemistry, Chemical Engineering and
Materials Science, Key Laboratory of Molecular and Nano
Probes, Ministry of Education, Collaborative Innovation
Center of Functionalized Probes for Chemical Imaging in
Universities of Shandong, Institute of Biomedical Science,
Shandong Normal University, Jinan 250014, People’s
Republic of China
(7) Hashimoto, E.; Tokushige, K.; Ludwig, J. J. Hepatol. Res. 2015,
45, 20−28.
(8) Cleveland, E.; Bandy, A.; Vanwagner, L. B. Clin. Liver Dis. 2018,
11, 98−104.
(9) Anstee, Q. M.; Goldin, R. D. Int. J. Exp. Pathol. 2006, 87, 1−16.
(10) Liss, K. H. H.; Finck, B. N. Biochimie 2017, 136, 65−74.
(11) Latruffe, N.; Malki, M. C.; Nicolas-Frances, V.; Clemencet, M.-
C.; Jannin, B.; Berlot, J.-P. Biochem. Pharmacol. 2000, 60, 1027−1032.
(12) Tessari, P.; Coracina, A.; Cosma, A.; Tiengo, A. Nutr., Metab.
Cardiovasc. Dis. 2009, 19, 291−302.
(13) Harano, Y.; Yasui, K.; Toyama, T.; Nakajima, T.; Mitsuyoshi,
H.; Mimani, M.; Hirasawa, T.; Itoh, Y.; Okanoue, T. Liver Int. 2006,
26, 613−620.
(14) Masarone, M.; Rosato, V.; Dallio, M.; Gravina, A.; Aglitti, A.;
Loguercio, A.; Federico, C.; Persico, M. Oxid. Med. Cell. Longevity
2018, 2018, 9547613.
(15) Takaki, A.; Kawai, D.; Yamamoto, K. Int. J. Mol. Sci. 2013, 14,
20704−20728.
(16) Ye, G.; Gao, H.; Wang, Z.; Lin, Y.; Liao, X.; Zhang, H.; Chi, Y.;
Zhu, S.; Dong, J. Cell Death Dis. 2019, 10, 39.
(17) Michelet, X.; Dyck, L.; Hogan, A.; Loftus, R. M.; Duquette, D.;
Wei, K.; Beyaz, S.; Tavakkoli, A.; Foley, C.; Donnelly, R.; O’Farrelly,
C.; Raverdeau, M.; Vernon, A.; Pettee, W.; O’Shea, D.; Nikolajczyk,
B. S.; Mills, K. H. G.; Brenner, M. B.; Finlay, D.; Lynch, L. Nat.
Immunol. 2018, 19, 1330−1340.
(18) Zhou, Y.; Li, P.; Wang, X.; Wu, C.; Fan, N.; Liu, X.; Wu, L.;
Zhang, W.; Zhang, W.; Liu, Z.; Tang, B. Chem. Sci. 2020, 11, 12149−
12156.
(19) Li, X.; Li, X.; Ma, H. Chem. Sci. 2020, 11, 1617−1622.
(20) Cheng, D.; Xu, W.; Gong, X.; Yuan, L.; Zhang, X.-B. Acc. Chem.
Res. 2020, 54, 403−415.
(21) Guo, L.; Tian, M.; Zhang, Z.; Lu, Q.; Liu, Z.; Niu, G.; Yu, X. J.
Am. Chem. Soc. 2021, 143, 3169−3179.
(22) Wolstenholme, C. H.; Hu, H.; Ye, S.; Funk, B. E.; Jain, D.;
Hsiung, C.-H.; Ning, G.; Liu, Y.; Li, X.; Zhang, X. J. Am. Chem. Soc.
2020, 142, 17515−17523.
(23) Murfin, L. C.; Weber, M.; Park, S. J.; Kim, W. T.; Lopez-Alled,
C. M.; McMullin, C. L.; Pradaux-Caggiano, F.; Lyall, C. L.; Kociok-
Köhn, G.; Wenk, J.; Bull, S. D.; Yoon, J.; Kim, H. M.; James, T. D.;
Lewis, S. E. J. Am. Chem. Soc. 2019, 141, 19389−19396.
(24) Ding, Q.; Tian, Y.; Wang, X.; Li, P.; Su, D.; Wu, C.; Zhang, W.;
Tang, B. J. Am. Chem. Soc. 2020, 142, 20735−20743.
(25) Wang, X.; Li, P.; Ding, Q.; Wu, C.; Zhang, W.; Tang, B. Angew.
Chem., Int. Ed. 2019, 58, 4674−4678.
(26) Jiang, M.; Gu, X.; Lam, J. W. Y.; Zhang, Y.; Kwok, R. T. K.;
Wong, K. S.; Tang, B. Z. Chem. Sci. 2017, 8, 5440−5446.
(27) Yin, J.; Peng, M.; Lin, W. Sens. Actuators, B 2019, 288, 251−
258.
Complete contact information is available at:
https://pubs.acs.org/10.1021/acs.analchem.1c01776
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
This work was supported by the National Natural Science
Foundation of China (21927811 and 22074083), the Key
Research and Development Program of Shandong Province
(2018YFJH0502), and the National Science Foundation of
Shandong Province of China (ZR2020ZD17).
(28) Yuan, Z.; Chen, J.; Zhou, Q.; Liu, A.; Qiang, Z.; Fang, M.;
Chen, M.; Feng, Y.; Yu, H.; Yang, X.; Meng, X. Chin. Chem. Lett.
2021, 32, 1803−1808.
(29) Xiao, H.; Li, P.; Tang, B. Coord. Chem. Rev. 2021, 427, 213582.
(30) Jankowski, A.; Kim, J. H.; Collins, R. F.; Daneman, R.; Walton,
P.; Grinstein, S. J. Biol. Chem. 2001, 276, 48748−48753.
(31) Fan, L.; Wang, X.; Ge, J.; Li, F.; Wang, X.; Wang, J.; Shuang, S.;
Dong, C. Chem. Commun. 2019, 55, 4703−4706.
(32) Yoshihara, T.; Maruyama, R.; Shiozaki, S.; Yamamoto, K.; Kato,
S.-i.; Nakamura, Y.; Tobita, S. Anal. Chem. 2020, 92, 4996−5003.
(33) Li, M.; Fan, J.; Li, H.; Du, J.; Long, S.; Peng, X. Biomaterials
2018, 164, 98−105.
(34) Zeng, X.; Chen, Z.; Tang, L.; Yang, H.; Liu, N.; Zhou, H.; Li,
Y.; Wu, J.; Deng, Z.; Yu, Y.; Deng, H.; Hong, X.; Xiao, Y. Chem.
Commun. 2019, 55, 2541−2544.
REFERENCES
■
(1) Zhou, J.; Zhou, F.; Wang, W.; Zhang, X.; Ji, Y.; Zhang, P.; She,
Z.; Zhu, L.; Cai, J.; Li, H. Hepatology 2020, 71, 1851−1864.
(2) Paik, J. M.; Golabi, P.; Younossi, Y.; Mishra, A.; Younossi, Z.
Hepatology 2020, 72, 1605−1616.
(3) Stefan, N.; Häring, H.-U.; Cusi, K. Lancet Diabetes Endocrinol.
2019, 7, 313−324.
(4) Fekry, B.; Ribas-Latre, A.; Baumgartner, C.; Mohamed, A. M. T.;
Kolonin, M. G.; Sladek, F. M.; Younes, M.; Eckel-Mahan, K. L. Cancer
Res. 2019, 79, 5860−5873.
9619
https://doi.org/10.1021/acs.analchem.1c01776
Anal. Chem. 2021, 93, 9609−9620

Analytical Chemistry
pubs.acs.org/ac
Article
(35) Li, Y.; Xie, X.; Yang, X. e.; Li, M.; Jiao, X.; Sun, Y.; Wang, X.;
Tang, B. Chem. Sci. 2017, 8, 4006−4011.
(36) Zhou, Y.; Li, P.; Fan, N.; Wang, X.; Liu, X.; Wu, L.; Zhang, W.;
Zhang, W.; Ma, C.; Tang, B. Chem. Commun. 2019, 55, 6767−6770.
(37) Xie, X.; Tang, F.; Liu, G.; Li, Y.; Su, X.; Jiao, X.; Wang, X.;
Tang, B. Anal. Chem. 2018, 90, 11629−11635.
(38) Ye, M.; Wang, X.; Tang, J.; Guo, Z.; Shen, Y.; Tian, H.; Zhu,
W.-H. Chem. Sci. 2016, 7, 4958−4965.
(39) Jiang, N.; Fan, J.; Xu, F.; Peng, X.; Mu, H.; Wang, J.; Xiong, X.
Angew. Chem., Int. Ed. 2015, 54, 2510−2514.
(40) Li, P.; Guo, X.; Bai, X.; Wang, X.; Ding, Q.; Zhang, W.; Zhang,
W.; Tang, B. Anal. Chem. 2019, 91, 3382−3388.
(41) Ziamajidi, N.; Khaghani, S.; Hassanzadeh, G.; Vardasbi, S.;
Ahmadian, S.; Nowrouzi, A.; Ghaffari, S. M.; Abdirad, A. Food Chem.
Toxicol. 2013, 58, 198−209.
(42) Heffern, M. C.; Park, H. M.; Au-Yeung, H. Y.; Van de Bittner,
G. C.; Ackerman, C. M.; Stahl, A.; Chang, C. J. Proc. Natl. Acad. Sci.
U.S.A. 2016, 113, 14219−14224.
(43) Kucera, O.; Cervinkova, Z. World J. Gastroenterol. 2014, 20,
8364−8376.
(44) Kanuri, G.; Bergheim, I. Int. J. Mol. Sci. 2013, 14, 11963−
11980.
(45) Athyros, V. G.; Alexandrides, T. K.; Bilianou, H.; Cholongitas,
E.; Doumas, M.; Ganotakis, E. S.; Goudevenos, J.; Elisaf, M. S.;
Germanidis, G.; Giouleme, O.; Karagiannis, A.; Karvounis, C.; Katsiki,
N.; Kotsis, V.; Kountouras, J.; Liberopoulos, E.; Pitsavos, C.; Polyzos,
S.; Rallidis, L. S.; Richter, D.; Tsapas, A. G.; Tselepis, A. D.; Tsioufis,
K.; Tziomalos, K.; Tzotzas, T.; Vasiliadis, T. G.; Vlachopoulos, C.;
Mikhailidis, D. P.; Mantzoros, C. Metabolism 2017, 71, 17−32.
(46) Schreuder, T. C.; Nieuwkerk, C.; Mulder, C. World J.
Gastroenterol. 2008, 14, 2474−2486.
(47) Zhao, P.; Sun, X.; Chaggan, C.; Liao, Z.; Wong, K.; He, F.;
Singh, S.; Loomba, R.; Karin, M.; Witztum, J. L.; Saltiel, A. R. Science
2020, 367, 652−660.
(48) Gong, J.; Fang, C.; Zhang, P.; Wang, P. X.; Qiu, Y.; Shen, L. J.;
Zhang, L.; Zhu, X. Y.; Tian, S.; Li, F.; Wang, Z.; Huang, Z.; Wang, A.;
Zhang, X. D.; She, Z. G. Hepatology 2019, 69, 524−544.
(49) Oh, H. J.; Kim, T. H.; Sohn, Y. W.; Kim, Y. S.; Oh, Y. R.; Cho,
E. Y.; Shim, S. Y.; Shin, S. R.; Han, A. L.; Yoon, S. J.; Kim, H. C.
Hepatology 2011, 17, 27−36.
(50) Valenti, L.; Fracanzani, A.; Dongiovanni, P.; Rovida, S.;
Rametta, R.; Fatta, E.; Pulixi, E.; Maggioni, M.; Fargion, A. World J.
Gastroenterol. 2014, 20, 3002−3010.
(51) Zelber-Sagi, S.; Ivancovsky-Wajcman, D.; Fliss-Isakov, N.;
Hahn, M.; Webb, M.; Shibolet, O.; Kariv, R.; Tirosh, O. Antioxidants
2020, 9, 578.
(52) Rakhshandehroo, M.; Hooiveld, G.; Muller, M.; Kerten, S. PloS
One 2009, 4, No. e6796.
(53) Cheng, D.; Gong, X.; Wu, Q.; Yuan, J.; Lv, Y.; Yuan, L.; Zhang,
X. Anal. Chem. 2020, 92, 11396−11404.
(54) Pafili, K.; Roden, M. Mol. Metabol. 2020, 53, 101122.
(55) Chavez-Tapia, N. C.; Rosso, N.; Tiribelli, C. BMC Gastro-
enterol. 2012, 12, 20.
(56) Bai, J.; Xia, M.; Xue, Y.; Ma, F.; Cui, A.; Sun, Y.; Han, Y.; Xu,
X.; Zhang, F.; Hu, Z.; Liu, Z.; Liu, Y.; Cai, G.; Su, W.; Sun, X.; Wu,
H.; Yan, H.; Chang, X.; Hu, X.; Bian, H.; Xia, P.; Gao, J.; Li, Y.; Gao,
X. EBioMedicine 2020, 57, 102849.
(57) Azzimato, V.; Jager, J.; Chen, P.; Morgantini, C.; Levi, L.;
Barreby, E.; Sulen, A.; Oses, C.; Craige, S.; Aouadi, M. Sci. Transl.
Med. 2020, 12, No. eaaw9709.
(58) Oswal, D. P.; Alter, G. M.; Rider, S. D.; Hostetler, H. A. J. Mol.
Graph. Model. 2014, 51, 27−36.
(59) Dhoke, G. V.; Gangwal, R. P.; Sangamwar, A. T. J. Mol. Struct.
2012, 1028, 22−30.
(60) Xu, H. E.; Lambert, M. H.; Montana, V. G.; Plunket, K. D.;
Moore, L. B.; Collins, J. L.; Oplinger, J. A.; Kliewer, S. A.; Gampe, R.
T.; McKee, D. D.; Moore, J. T.; Willson, T. M. Proc. Natl. Acad. Sci. U.
S. A. 2001, 98, 13919−13924.
9620
https://doi.org/10.1021/acs.analchem.1c01776
Anal. Chem. 2021, 93, 9609−9620