Redox Biology 72 (2024) 103158
Contents lists available at ScienceDirect
Redox Biology
journal homepage: www.elsevier.com/locate/redox
PM2.5-induced iron homeostasis imbalance triggers cardiac hypertrophy
through ferroptosis in a selective autophagy crosstalk manner
Tianyu Li a, b, Mengqi Sun a, b, Qinglin Sun a, b, Xiaoke Ren a, b, Qing Xu c, Zhiwei Sun a, b, **,
Junchao Duan a, b, *
a
b
c
Department of Toxicology and Sanitary Chemistry, School of Public Health, Capital Medical University, Beijing, 100069, PR China
Beijing Key Laboratory of Environmental Toxicology, Capital Medical University, Beijing, 100069, PR China
Core Facilities for Electrophysiology, Core Facilities Center, Capital Medical University, Beijing, 100069, PR China
A R T I C L E I N F O
A B S T R A C T
Keywords:
PM2.5
Cardiac hypertrophy
Ferroptosis
NCOA4
MitoQ
Exposure to PM2.5 is correlated with cardiac remodeling, of which cardiac hypertrophy is one of the main clinical
manifestations. Ferroptosis plays an important role in cardiac hypertrophy. However, the potential mechanism of
PM2.5-induced cardiac hypertrophy through ferroptosis remains unclear. This study aimed to explore the mo­
lecular mechanism of cardiac hypertrophy caused by PM2.5 and the intervention role of MitoQ involved in this
process. The results showed that PM2.5 could induce cardiac hypertrophy and dysfunction in mice. Meanwhile,
the characteristics of ferroptosis were observed, such as iron homeostasis imbalance, lipid peroxidation, mito­
chondrial damage and abnormal expression of key molecules. MitoQ treatment could effectively mitigate these
alternations. After treating human cardiomyocyte AC16 with PM2.5, ferroptosis activator (Erastin) and inhibitor
(Fer-1), it was found that PM2.5 could promote ferritinophagy and lead to lipid peroxidation, mitochondrial
dysfunction as well as the accumulation of intracellular and mitochondrial labile iron. Subsequently, mitophagy
was activated and provided an additional source of labile iron, enhancing the sensitivity of AC16 cells to fer­
roptosis. Furthermore, Fer-1 alleviated PM2.5-induced cytotoxicity and iron overload in the cytoplasm and
mitochondria of AC16 cells. It was worth noting that during the process of PM2.5 caused ferroptosis, abnormal
iron metabolism mediated the activation of ferritinophagy and mitophagy in a temporal order. In addition,
NCOA4 knockdown reversed the iron homeostasis imbalance and lipid peroxidation caused by PM2.5, thereby
alleviating ferroptosis. In summary, our study found that iron homeostasis imbalance-mediated the crosstalk of
ferritinophagy and mitophagy played an important role in PM2.5-induced ferroptosis and cardiac hypertrophy.
1. Introduction
Epidemiological studies have shown that PM2.5 exposure is corre­
lated with the increased risk of CVD [5,6]. Cardiac hypertrophy is a
leading cause of various PM2.5-induced CVD [7]. Evidence from a pop­
ulation study indicated that for every 5 μg/m3 elevation of PM2.5 con­
centration, the left ventricular mass index (LVMI) increased by 4.0 g/m2
[8]. Iron homeostasis is essential for maintaining normal cardiac
structure and function [9]. Recently, as a programmed cell death related
to iron homeostasis imbalance, ferroptosis has been proven to be an
important trigger of cardiac hypertrophy [10]. However, there is no
report on the potential mechanism of PM2.5 leading to cardiac hyper­
trophy through ferroptosis.
As the form of iron in circulation, Fe3+ is reduced to Fe2+ and
released into the labile iron pool (LIP) after entering the cell. Excessive
Air pollution is considered as the main environmental health risk. Air
pollution contributed to 7 million premature deaths worldwide ac­
cording to the World Health Organization (WHO) [1]. Besides, 4.14
million global premature deaths were attributed to long-term exposure
to PM2.5, which was the largest driver of air pollution’s disease burden
worldwide [2]. Based on the data from World Health Statistics 2023,
premature deaths caused by cardiovascular disease (CVD) were No.1
among the four major non-communicable diseases [3]. The results of the
Global Burden of Disease (GBD) showed that CVD accounts for 51.5 % of
the total DALYs ascribed to PM2.5 [4]. Therefore, PM2.5 is the most
consistent and reliable predictor of CVD mortality.
* Corresponding author. Department of Toxicology and Sanitary Chemistry, School of Public Health, Capital Medical University, Beijing, 100069, PR China.
** Corresponding author. Department of Toxicology and Sanitary Chemistry, School of Public Health, Capital Medical University, Beijing, 100069, PR China.
E-mail addresses: zwsun@ccmu.edu.cn (Z. Sun), jcduan@ccmu.edu.cn (J. Duan).
https://doi.org/10.1016/j.redox.2024.103158
Received 10 January 2024; Received in revised form 27 March 2024; Accepted 10 April 2024
Available online 12 April 2024
2213-2317/© 2024 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY-NC license (http://creativecommons.org/licenses/bync/4.0/).
T. Li et al.
Redox Biology 72 (2024) 103158
labile iron is stored by ferritin complexes (composed of ferritin heavy
and light chains) [11]. Subsequently, ferritin could be transported to
lysosomes for degradation after being specifically recognized by NCOA4
(a ferritinophagy-related molecule). The accumulation of Fe2+ released
from ferritin promoted the production of phospholipid hydroperoxides
(PL-OOH) by mediating the Fenton reaction, which in turn triggered
ferroptosis [12]. A recent study has shown that ferritinophagy played an
important role in airway injury induced by ferroptosis after exposure to
PM2.5 [13]. Notwithstanding, there is still a lack of evidence on whether
ferritinophagy is the inducement of ferroptosis in PM2.5-triggered car­
diac hypertrophy.
Mitochondria are the most abundant organelles in heart. When iron
overload occurred in cytoplasm, excess labile iron was stored by mito­
chondrial ferritin (FTMT) to maintain iron homeostasis [14]. Recently,
Sandoval et al. reported that after the disruption of iron homeostasis,
labile iron accumulated into mitochondria and caused redox imbalance,
ultimately resulting in mitophagy [15]. After activation of mitophagy, a
large amount of ferrous iron is released and further accumulated in
cytoplasm, exacerbating cell sensitivity to ferroptosis [16]. Recent
studies have shown that ferritinophagy and mitophagy might contribute
to ferroptosis [17,18]. Nevertheless, the regulatory relationship between
iron homeostasis imbalance and the two important selective autophagy
(ferritinophagy and mitophagy) in PM2.5-induced ferroptosis remains
unclear.
We attempted to investigate the potential mechanism of PM2.5induced cardiac hypertrophy through ferroptosis in this study. Firstly,
C57BL/6J mice were used to observe the cardiac hypertrophy and fer­
roptosis due to PM2.5. Subsequently, the regulatory process of PM2.5triggered ferroptosis was identified using AC16 cells. Lastly, the precise
mechanisms of PM2.5-induced iron homeostasis imbalance-mediated the
crosstalk of ferritinophagy and mitophagy were examined using a time
gradient exposure model and the stable NCOA4 knockdown AC16 cell
line. In addition, the intervention effect of MitoQ on cardiac toxicity
induced by PM2.5 was also evaluated. The data exhibited in this study
provided molecular evidence for elucidating the pathogenesis of heart
disease caused by PM2.5, which is of great significance in reducing the
risk and burden of CVD related to PM2.5.
respiratory volume of adult mice weighing 25 g is about 0.15 mL/time,
and the respiratory rate is 163 times/min), combined with the concen­
tration of PM2.5 (the interim target-I 35 μg/m3) in the WHO global air
quality guidelines, and the 100-fold uncertainty coefficient according to
toxicology textbooks.
2.3. Echocardiographic assessment
After the final treatment, the cardiac function and structure were
assessed with the Vevo 2100 small animal ultrasound imaging system
(FUJIFILM VisualSonics, United States). The mice were anesthetized
with 350 mg/kg tribromoethanol injection. The chest of mice was
shaved and placed in the supine position. Mitral valve Doppler echo­
cardiography and left ventricular short-axis M-mode images were
recorded by averaging the data during 5 consecutive heartbeats.
The left ventricular systolic and diastolic wall thickness, volume and
mass were assessed by M-mode. Mitral valve Doppler echocardiography
was employed to determine diastolic function by measuring E/A ratio,
IVCT and IVRT. All parameters were determined at least 3–5 cardiac
cycles.
2.4. Histopathological analysis
The heart tissue was isolated and washed with precooled PBS three
times, followed by paraformaldehyde (4%) fixation and paraffin
embedding. The heart sections (5 μm thick) were stained with Wheat
Germ Agglutinin (WGA) (Sigma, USA) as well as H&E for histological
analysis and measurement of the cross-sectional area of cardiomyocytes.
An automated slide scanning system (3Dhistech, Hungary) was used to
image the slices.
2.5. Ultrastructural observation by TEM
2. Materials and methods
The ultrastructure of myocardium was assessed with transmission
electron microscope (TEM). In brief, fresh heart tissues were perfused
with pre-chilled PBS and then rapidly fixed in glutaraldehyde (2.5%) for
6 h. Subsequently, the fully fixed tissue was washed, dehydrated,
embedded and cut into ultrathin sections (about 50–60 nm). The ultrathin sections were imaged by a TEM (JEOL JEM2100, Tokyo, Japan).
2.1. PM2.5 samples preparation
2.6. Assessment of lipid peroxidation
The PM2.5 samples used in this experiment were collected from
Capital Medical University in Beijing with a high-flow atmospheric
particulate sampler (TH-1000C II, Wuhan Tianhong, China). Subse­
quently, the collected PM2.5 was eluted, lyophilized, and subsequently
sterilized with UV. The physicochemical characterization of the PM2.5
samples including morphology, particle size and composition has been
described in our previous study [19].
The level of 4-HNE in heart tissues and AC16 cells was detected with
ELISA kits (mlbio, Shanghai, China). The contents of GSH and MDA in
heart tissues and AC16 cells were measured by the corresponding kits
(Nanjing Jiancheng Bioengineering Institute, China).
2.7. Cell culture and treatment
The human cardiomyocytes AC16 cell line was provided by Shanghai
Institutes for Biological Sciences (SIBS) (Shanghai, China). The AC16
were grown in DMEM/F12 (VivaCell, China) containing 1% strepto­
mycin & penicillin mixture (key GEN, China) and 10% fetal bovine
serum (Corning, USA), which was incubated at 37 ◦ C in a 5% CO2
incubator (Thermo, USA). AC16 were inoculated and converged to
70–80% density before treatment. All experiments were performed with
five biological replicates per group. The stable NCOA4 knockdown
AC16 cell line was established with short hairpin RNA (shRNA) inter­
ference. 293T cells were inoculated and cultured in an incubator with
5% CO2 at 37 ◦ C for 24 h. The cell culture medium was changed with 10
mL of the complete medium without antibiotics and added the mixed
reagent containing 5 μg Plvx-shRNA-Zsgreen-T2a-puro recombinant
vector, 1.25 μg pMD2.G vector, 3.75 μg pspAX2 vector and 25 μL Lip­
ofectamine 2000 per 1 mL Opti-MEM to transfect 293T cell for 8 h. After
that, 293T cells were cultured for 16 h in newly replaced complete
medium (without antibiotics), followed by 24 h in complete medium
2.2. Experimental animals and treatments
The Animal Ethics Committee of Capital Medical University
approved the animal experiment in this study (Ethics No. AEEI-2016076). Male C57BL/6J mice (Seven weeks old) were obtained from the
Vital River Laboratory Animal Technology Co., Ltd. (Beijing, China) and
maintained under standard conditions (12/12 h light/dark cycle,
20–24 ◦ C, 40%–60% humidity). Following a week of adaptive feeding,
C57BL/6J mice were split into four groups (the Control group, the
MitoQ group, the PM2.5 group and the PM2.5+MitoQ group) at random.
Anhydrous ethanol dilution and MitoQ (5 mg/kg⋅bw) were given
through gavage daily. Saline and PM2.5 suspension (5 mg/kg⋅bw) were
given through intratracheal instillation every three days for four weeks.
The basis for PM2.5 exposure dose was described in our previous study
[20]. The determination of PM2.5 exposure dose (5 mg/kg⋅bw) was
based on the respiratory physiological parameters of mice (the average
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Redox Biology 72 (2024) 103158
Fig. 1. PM2.5 promoted cardiac hypertrophy and ferroptosis in C57BL/6J mice. (A) M-mode echocardiography. (B) Mitral valve doppler echocardiography. (C)
Representative images of H&E staining and WGA staining in heart sections (Scale bar: 20 μm). (D) Representative TEM images of myocardial tissue (Scale bar: 2 μm).
(E-J) Left ventricular wall thickness and internal diameter. (K-M) Left ventricular diastolic function. (N) Quantitative analysis of cardiomyocyte cross-sectional area.
(O-Q) The levels of 4-HNE, MDA and GSH (nmol/mgprot). (R–S) The contents of Fe2+ and total iron. All data were expressed as mean ± S.D. * represents compared
to the control group, # represents compared to the PM2.5 group. *p < 0.05, **p < 0.01, #p < 0.05.
containing penicillin & streptomycin. After collection and concentra­
tion, the biological titer of lentivirus was measured. The concentrated
lentivirus was used to infect AC16 to establish the stable NCOA4
knockdown AC16 cell lines. The puromycin was adopted to screen virusinfected AC16 cells. The sh-RNA targeting NCOA4 sequence was 5′TGAACAGGTGGACCTTATTTA-3’.
2.10. ROS and lipid ROS assay
The intracellular Lipid ROS was assessed with BODIPYTM 581/591
C11 probe. After being stained with C11-BODIPY581/591 (5 μM) for 30
min and Hoechst (1 μM) for 10 min at 37 ◦ C, cells were imaged with a
laser scanning confocal microscope. Image J was used to measure the
relative fluorescence intensity. Besides, the level of ROS was assessed
with CM-H2DCFDA probe (5 μM). The relative fluorescence intensity of
FITC and PE-Texas channels was quantified by flow cytometry.
2.8. Assessment of cell viability
Does gradient experiments were performed using PM2.5 suspended in
DMEM at different concentrations (0, 12.5, 25, 50 μg/mL) for 24 h.
Besides, AC16 cells were pretreated with 0.5 μM MitoQ for 2 h prior to
being exposed to PM2.5. Furthermore, 5 μM Ferrostatin-1 (Fer-1) or 5 μM
Erastin was used to pretreat AC16 cells for 2 h, followed by exposure to
PM2.5 for 24 h. The cell viability was determined by CCK-8 (Tongren,
Japan).
2.11. Immunofluorescence staining
Rewarmed Frozen sections of heart and AC16 cells were per­
meabilized in Triton X-100 (0.5%) for 30 min and blocked with 10%
fetal bovine serum at 37 ◦ C for 1.5 h after being fixed with para­
formaldehyde for 30 min. Subsequently, the heart sections and AC16
cells were co-incubated with FTH1 (1:100) & NCOA4 (1: 50), DHODH
(1: 100) & COX-2 (1: 50) and PINK1 (1:100) & Parkin (1:50) overnight
at 4 ◦ C, followed by incubation with secondary antibody (1:200) at 37 ◦ C
for 1.5 h. Following the addition of DAPI to the heart sections and AC16
cells, confocal laser microscope images were captured. The semiquantitative methods with Image J were similar with the Iron and
Lipid ROS assay.
2.9. The level and mobilization of ferrous iron
The levels of cardiac total iron and ferrous iron were measured using
the iron assay kit (Abcam, 83366). A microplate reader was used to
measure the absorbance at 593 nm. The content of intracellular and
mitochondrial ferrous iron in AC16 was measured by FerroOrange (1
μM) and Mito-FerroGreen (5 μM) probes (DojinDo, Japan). After being
stained with probes for 30 min at 37 ◦ C, cells were collected for fluo­
rescence intensity assessment by flow cytometry. To evaluate the
mobilization of ferrous iron, cells were co-incubated with FerroOrange
& LysoTracker (1 μM) and FerroOrange & MitoTracker (1 μM) for 30
min. Moreover, a confocal laser microscope was adopted to acquire
images after being stained with 1 μM Hoechst (Solarbio) for 10 min.
Image J was used to measure the relative fluorescence intensity.
2.12. Assessment of mitochondrial function
Mitochondrial ATP production, OCR and the activity of mitochon­
drial complexes were separately detected by ATP determination kit
(Invitrogen, USA), OCR Assay Kit (Abcam, UK) and Mitochondrial
complex I - III Activity Assay Kits (Solarbio, China). Extracellular flux
analyzer (Seahorse XFe24, Agilent, USA) was adopted to detect mito­
chondrial stress, which was expressed by Maximal Respiration, ATP
production, Spare Respiratory Capacity, Basal Respiration, Non3
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Redox Biology 72 (2024) 103158
Fig. 2. PM2.5 resulted in cardiac ferritinophagy and mitophagy in C57BL/6J mice. (A) Representative immunofluorescence images of COX-2 and DHODH
proteins in heart (scale bar: 50 μm). (B) Representative immunofluorescence images of FTH1 and NCOA4 proteins in heart (scale bar: 50 μm). (C) Representative
immunofluorescence images of PINK1 and Parkin proteins in heart (scale bar: 25 μm). (D) Representative Western blot pictures. (E-J) The quantitative analysis of
COX-2, DHODH, FTH1, NCOA4, PINK1 and Parkin fluorescence intensity. (K–V) The quantitative analysis of ANP, BNP, DHODH, COX-2, NCOA4, FTH1, FTMT, LC3,
p62, Mfn2, Drp1, OPA1. All data were expressed as mean ± S.D. * represents compared to the control group, # represents compared to the PM2.5 group. *p < 0.05,
**p < 0.01, #p < 0.05. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)
Mitochondrial Oxygen Consumption and Proton leak. After inoculating
and converging to 70–80% density in a XFe24-well microplate (Agilent,
USA), AC16 cells were treated with PM2.5 suspended in DMEM at
different concentrations. Meanwhile, the probe plate was hydrated
overnight at 37 ◦ C in a non-CO2 environment. Cells were equilibrated in
the XF DMEM (Agilent, USA) for 1 h, with a final volume of 500 μL for
each well in a 37 ◦ C non-CO2 incubator. After that, oligomycin (1 μM),
FCCP (1 μM) and Rotenone/antimycin A (0.5 μM) were sequentially
added in the specific holes on the Seahorse XFe24 plate to detect the
relevant OCR. Ultimately, the OCR values were analyzed using Seahorse
Wave software and normalized through the cell densities.
Odyssey ® CLx Infrared Imaging System were employed to detect the
signals of protein bands. Then, the Image Studio™ Software was used for
visualization. Image J was used to analyze the density of the detected
protein bands.
2.14. Statistical analysis
All experimental data were analyzed by SPSS 22.0 and shown as
means ± standard deviations (SD). The Shapiro-Wilk test was used for
testing normality of data. F-test was adopted to test equal variance of
data. And Tukey’s multiple comparisons test was used for ANOVA. In
addition, the Welch’s test, Tamhane’s T2 test and Dunnett’s T3 test were
used when normality or equal variance test failed. GraphPad Prism 7
was used to make the statistical chart. Significance levels were defined
as *p < 0.05, **p < 0.01 (2-sided).
2.13. Western blotting
The protein of heart tissue and AC16 cells was extracted using whole
Cell Lysis Assay Kit (KeyGEN Biotech, China) and then the concentration
was detected by BCA kit (Dingguo Changsheng Biotech, China). The
proteins were loaded in equal amounts, separated by SDS-PAGE gels and
transferred to NC membranes. The membranes were incubated at 4 ◦ C
overnight with primary antibodies after blocking in TBS with skim milk
(5%), which include ANP, PINK1 (NOVUS, USA); FTH1, Mfn2, LC3A/B,
Drp1, Parkin, GAPDH (CST, USA); DHODH (Proteintech, USA); and
BNP, FTMT, p62, COX-2, NCOA4, OPA1, 4-HNE, MDA (Abcam, UK). The
next day, secondary antibody was used to incubate with the membranes.
The ChemiDocTM Imaging System (Bio-Rad, USA) or the LI-COR
3. Results
3.1. PM2.5 promoted the cardiac hypertrophy and ferroptosis
Echocardiographic analysis was used to evaluate the effects of PM2.5
on the cardiac function of mice. Firstly, M-mode echocardiography
showed that mice exposed to PM2.5 exhibited significant rises of LVAWs,
LVPWs, LVAWd and LVPWd compared with the control group, as well as
decreases in LVIDs and LVIDd (Fig. 1A and 1E-J). These results
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Redox Biology 72 (2024) 103158
Fig. 3. PM2.5 caused ferroptosis and mitochondrial dysfunction in cardiomyocytes. (A-C) Representative fluorescence images of FerroOrange probe staining,
Mito-FerroGreen probe staining and BODIPY™ 581/591 C11 staining (scale bar: 20 μm). (D) The cell viability of AC16 cells. (E-F) The levels of 4-HNE, MDA. (G-H)
The activity of mitochondrial complex I and complex II. (I–K) The fluorescence intensity analysis of intracellular Fe2+, mitochondrial Fe2+ and Lip ROS content. (L)
Curves of oxygen consumption rate. (M) The quantitative analysis of mitochondrial respiration. (N) Representative fluorescence images of Mito-FerroGreen probe
staining (scale bar: 20 μm). (O) Representative fluorescence intensity image obtained from flow cytometry of FerroOrange probe staining. (P-Q) The fluorescence
intensity analysis of intracellular Fe2+, mitochondrial Fe2+. All data were expressed as mean ± S.D. * represents compared to the control group. *p < 0.05, **p
< 0.01.
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Redox Biology 72 (2024) 103158
Fig. 4. MitoQ attenuated PM2.5-induced iron homeostasis imbalance and cardiac ferroptosis in cardiomyocytes. (A-C) Representative fluorescence images of
FerroOrange probe staining, Mito-FerroGreen probe staining and BODIPY™ 581/591 C11 staining (scale bar: 20 μm). (D) Representative immunofluorescence
images of COX-2 and DHODH proteins (scale bar: 20 μm). (E) Representative Western blot pictures. (F–I) The quantitative analysis of ANP, BNP, DHODH and COX-2.
(J) The levels of OCR. (K) The ATP production. (L-M) The activity of mitochondrial complex II and complex III. (N–P) The levels of 4-HNE, MDA and GSH. (Q-S) The
fluorescence intensity analysis of intracellular Fe2+, mitochondrial Fe2+ and Lip ROS content. (T-U) The quantitative analysis of DHODH and COX-2 fluorescence
intensity. (V–W) The quantitative analysis of intracellular Fe2+ by flow cytometry. (X–Y) The quantitative analysis of mitochondrial Fe2+ by flow cytometry. All data
were expressed as mean ± S.D. * represents compared to the control group, # represents compared to the PM2.5 group. *p < 0.05, **p < 0.01, #p < 0.05.
suggested that exposure to PM2.5 was correlated with increased wall
thickness and decreased inner diameter of left ventricles, accompanied
by systolic dysfunction. Left ventricular diastolic function was assessed
through mitral valve doppler echocardiography (Fig. 1B). It was
observed that IVCT, IVRT, and E/A ratios were reduced in the PM2.5
group (Fig. 1K-M), indicating the extension of left ventricular systolic/
diastolic time and alternation of diastolic function. In addition, the
higher left ventricular mass and heart/body weight ratio, as well as
lower left ventricular volume (systolic and diastolic) in response to
PM2.5 collectively implied the occurrence of cardiac hypertrophy
(Figure S1A-C and S3). The pathological analysis of cardiac tissue by
H&E staining displayed that compared with the control group, PM2.5
exposure resulted in enlargement and disordered arrangement of car­
diomyocytes as well as interstitium expansion (Fig. 1C). Meanwhile,
significant cardiac hypertrophy was identified through WGA staining in
the PM2.5 group (Fig. 1C and N). Furthermore, ANP and BNP were
upregulated after PM2.5 exposure (Fig. 2D). In summary, PM2.5 exposure
was closely related to cardiac systolic and diastolic dysfunction, which
was able to promote the progression of cardiac hypertrophy. MitoQ
effectively alleviated various cardiac dysfunctions and structural in­
juries such as cardiac hypertrophy caused by PM2.5.
To further explore the potential reason for PM2.5-induced cardiac
hypertrophy, the cardiac ultrastructural changes were observed under
TEM. It was found that PM2.5 exposure impaired the mitochondrial
structure, manifested by mitochondrial shrinkage, disappearance of
cristae and membrane rupture, as illustrated in Fig. 1D. In addition, our
data indicated that the accumulation of lipid peroxidation products (4HNE, MDA) and the consumption of GSH (reflecting the lipid
peroxidation repair capacity) occurred in the PM2.5 group (Fig. 1O-Q
and S4). Moreover, as shown in Fig. 1R and S, the elevated content of
total iron and ferrous iron was associated with PM2.5 exposure. The
evidence of above morphological characteristics and abnormal expres­
sion of ferroptosis markers all revealed the occurrence of ferroptosis. To
further validate our finding, western blotting was adopted to analyze the
expression of ferroptosis-related molecules. The Western blot evidence
showed that PM2.5 resulted in abnormal expression of DHODH and COX2 (ferroptosis marker) (Fig. 2A and D). The above results revealed that
PM2.5 triggered cardiac ferroptosis in mice, and MitoQ played an
effective intervention role.
3.2. PM2.5 activated the cardiac ferritinophagy and mitophagy in vivo
Multiple studies suggested that activation of selective autophagy
driven by iron homeostasis imbalance can promote ferroptosis. Ac­
cording to the clues provided above, Western blot and immunofluores­
cence were used to further explore the underlying mechanism of PM2.5induced ferroptosis. It was obtained that the ferritinophagy-related
molecules NCOA4 and FTH1 exhibited abnormal expression in the
PM2.5 group (Fig. 2B and D). It is worth noting that the level of FTMT (an
iron storage protein located in mitochondria) was downregulated due to
PM2.5 exposure (Fig. 2Q). Meanwhile, the LC3II/I ratio and p62 (also
known as SQSTM1) were significantly upregulated in mice exposed to
PM2.5 (Fig. 2R and S). Subsequently, it was observed in immunofluo­
rescence results that PM2.5 exposure was positively correlated with the
elevated expression of PINK1 and Parkin (mitophagy key molecules)
(Fig. 2C). In addition, PM2.5 affected the expression of mitochondrial
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Redox Biology 72 (2024) 103158
Fig. 5. PM2.5 disturbed the expression of ferritinophagy and mitophagy related molecules. (A) Representative immunofluorescence images of NCOA4 and FTH1
proteins (scale bar: 20 μm). (B) Representative fluorescence images of Fe2+ and lysosomes colocalization (scale bar: 20 μm). (C) Representative immunofluorescence
images of PINK1 and Parkin proteins (scale bar: 10 μm). (D) Representative Western blot pictures. (E-H) The quantitative analysis of NCOA4, FTH1, PINK1 and
Parkin fluorescence intensity. (I–P) The quantitative analysis of NCOA4, FTH1, FTMT, LC3, p62, Mfn2, Drp1, OPA1. All data were expressed as mean ± S.D. *
represents compared to the control group, # represents compared to the PM2.5 group. *p < 0.05, **p < 0.01, #p < 0.05. (For interpretation of the references to colour
in this figure legend, the reader is referred to the Web version of this article.)
dynamics-related proteins, manifested by downregulation of Mfn2 and
OPA1 and upregulation of Drp1, which indicated the existence of
mitochondrial fission and fusion dysfunction (Fig. 2T–V). MitoQ alle­
viated the ferritinophagy, mitophagy and mitochondrial dynamics
imbalance triggered by PM2.5.
mitochondrial oxygen consumption. The results in Fig. S2A suggested
that 5 μM of Erastin (a ferroptosis specific activator) significantly
reduced the cell viability of AC16. Meanwhile, the ferroptosis inhibitor
Fer-1 (5 μM) has been observed to effectively reverse the cytotoxicity of
PM2.5. In addition, it was observed that Fer-1 significantly reduced the
PM2.5-caused ferrous iron accumulation and mitochondrial distribution
through fluorescence probe-based confocal laser imaging and flow
cytometry (Fig. 3N-Q). All in all, these above results indicated that PM2.5
triggered ferroptosis in AC16.
3.3. PM2.5 induced ferroptosis and mitochondrial dysfunction in vitro
To validate PM2.5-induced ferroptosis and the activation of two se­
lective autophagy (ferritinophagy and mitophagy) observed in vivo and
further explore the specific regulatory processes, AC16 cells were
adopted to establish the PM2.5 exposure model in vitro. After being
treated with various doses of PM2.5, the cell viability of AC16 was
measured by CCK-8. The results indicated that the cytotoxicity of PM2.5
was dose-dependent (Fig. 3D). Furthermore, 50 μg/mL PM2.5 signifi­
cantly reduced the cell viability of AC16. Therefore, 0–50 μg/mL was
selected for this part, and 50 μg/mL was adopted for subsequent
mechanism exploration and reverse validation. Firstly, the effect of
PM2.5 on the accumulation and mobilization of ferrous iron was evalu­
ated. As shown in Fig. 3A and B, with the rise of PM2.5 concentration, the
labile iron gradually accumulated in the cytoplasm and further distrib­
uted into mitochondria. In addition, the level of ROS, Lipid ROS, 4-HNE
and MDA was elevated after exposure to PM2.5 (Fig. 3C, 3E-F and S5DE). The capture of these characteristic events of ferroptosis suggested the
importance of it in PM2.5-induced cardiac hypertrophy. Notably,
consistent with the mitochondrial injury found in vivo, it has been
detected that the activity of mitochondrial complexes I and II in AC16
was negatively associated with the concentration of PM2.5 (Fig. 3G and
H). Subsequently, Mitochondrial stress was analyzed through extracel­
lular flux analyzer. As shown in Fig. 3L-M, mitochondrial respiration
was remarkably inhibited by PM2.5, such as ATP production, maximum
respiration, basal respiration, spare respiration capacity and non-
3.4. MitoQ alleviated PM2.5-induced iron homeostasis imbalance and
cardiac ferroptosis
MitoQ, a mitochondrial-targeted antioxidant, can provide substrates
for the ferroptosis defense system to inhibit lipid peroxidation. As shown
in Figs. S2B–C, MitoQ (0.5 μM) significantly reversed the diminished cell
viability of AC16 caused by PM2.5, which was used in subsequent ex­
periments to intervene the cytotoxicity of PM2.5. With fluorescent probebased confocal microscopy imaging and flow cytometry, it was indicated
that MitoQ attenuated the accumulation and mitochondrial mobiliza­
tion of ferrous iron caused by PM2.5 (Fig. 4A and B and 4V–Y). Mean­
while, it was observed that MitoQ ameliorated PM2.5-induced lipid
peroxidation such as the elevated content of ROS, Lipid ROS, 4-HNE,
MDA, and the consumption of GSH, as shown in Fig. 4C, 4N–P and S5.
Immunofluorescence and Western blot results indicated that PM2.5
upregulated markers of cardiac hypertrophy (ANP & BNP) and ferrop­
tosis (COX-2), as well as downregulated DHODH. MitoQ alleviated the
expression abnormalities of the above markers (Fig. 4D and E). These
data demonstrated that MitoQ reduced the sensitivity of AC16 cells to
PM2.5-induced ferroptosis. Additionally, MitoQ significantly reversed
the lessening of OCR and ATP synthesis in AC16 cells, as well as restored
the activity of mitochondrial complexes II and III (Fig. 4J-M), indicating
7
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Redox Biology 72 (2024) 103158
Fig. 6. The crosstalk between ferritinophagy and mitophagy was mediated by PM2.5-triggered iron homeostasis imbalance. (A) Representative fluorescence
images of Fe2+ and lysosomes colocalization (scale bar: 20 μm). (B) Representative fluorescence images of Fe2+ and mitochondria colocalization (scale bar: 20 μm).
(C) Representative Western blot pictures. (D-J) The quantitative analysis of NCOA4, FTH1, FTMT, PINK1, Parkin, p62, LC3. (K) Comparison of expression patterns of
NCOA4, FTH1 and FTMT. All data were expressed as mean ± S.D. * represents compared to the control group, *p < 0.05, **p < 0.01. (For interpretation of the
references to colour in this figure legend, the reader is referred to the Web version of this article.)
its capacity to alleviate mitochondrial dysfunction caused by redox
imbalance.
upregulation of LC3II/I ratio and p62 was observed in AC16 cells
exposure to PM2.5. Additionally, the expression changes of Mfn2, Drp1
and OPA1 induced by PM2.5 exposure suggested the imbalance of
mitochondrial dynamics in AC16 cells. Consistent with the in vivo re­
sults, MitoQ exerted intervention effect on ferritinophagy, mitophagy
and mitochondrial dynamics imbalance caused by PM2.5.
3.5. Ferritinophagy and mitophagy activated by PM2.5 in vitro
Consistent with findings from in vivo studies, exposure to PM2.5
resulted in abnormal expression of ferritinophagy-related molecules
NCOA4 and FTH1 in AC16 cells (Fig. 5A and D). Meanwhile, the
colocalization of ferrous iron and lysosomes was observed in AC16 cells
exposed to PM2.5 (Fig. 5B). These results collectively demonstrated that
the activation of ferritinophagy is associated with PM2.5 exposure.
Subsequently, confocal imaging of immunofluorescence indicated that
PINK1 and Parkin were significantly activated in the PM2.5 group
(Fig. 5C). As shown in Fig. 5D, it could be seen that the expression of
FTMT is negatively correlated with PM2.5, which revealed the occur­
rence of mitochondrial iron homeostasis imbalance. Besides, significant
3.6. PM2.5-triggered iron homeostasis imbalance mediated the crosstalk of
ferritinophagy and mitophagy
To investigate the molecular mechanisms of iron homeostasis dis­
rupted by PM2.5 mediating the crosstalk of ferritinophagy and mitoph­
agy, AC16 cells were exposed to PM2.5 for 0, 3, 6, 12, 24 h to establish a
time series exposure model. It is worth noting that the mobilization of
ferrous iron into lysosomes and mitochondria exhibited different time
patterns. As shown in Fig. 6A, intracellular ferrous iron began to co8
T. Li et al.
Redox Biology 72 (2024) 103158
Fig. 7. NCOA4-mediated iron homeostasis imbalance is essential for PM2.5-triggered cardiomyocyte ferroptosis. (A) Representative immunofluorescence
images of FTH1 and NCOA4 proteins (scale bar: 10 μm). (B) Representative immunofluorescence images of PINK1 and Parkin proteins (scale bar: 10 μm). (C)
Representative immunofluorescence images of COX-2 proteins (scale bar: 20 μm). (D) Representative immunofluorescence images of DHODH proteins (scale bar: 10
μm). (E) Representative fluorescence images of FerroOrange probe staining (scale bar: 10 μm). (F) The mRNA expression of NCOA4. (G-L) The quantitative analysis
of NCOA4, FTH1, PINK1, Parkin, COX-2 and DHODH fluorescence intensity. (M) The fluorescence intensity analysis of intracellular Fe2+ content. All data were
expressed as mean ± S.D. * represents compared to the control group, *p < 0.05, **p < 0.01. (For interpretation of the references to colour in this figure legend, the
reader is referred to the Web version of this article.)
localize with lysosomes at 6 h PM2.5 exposure. What’s different is that
the colocalization of ferrous irons with mitochondria occurred at 12 h,
and gradually enhanced over time (Fig. 6B). It reminded us that the
activation of ferritinophagy preceded mitophagy, which was triggered
by PM2.5. The WB results showed that the expression of NCOA4 in AC16
cells was elevated with prolonged exposure to PM2.5 (Fig. 6C and D).
Interestingly, both FTH1 and FTMT were compensatory upregulated in
the first 6 h of PM2.5 exposure, followed by significant downregulation
(Fig. 6E and F), which indicated that iron homeostasis imbalance
appeared in both cytoplasm and mitochondria. Coincidentally, PINK1
and Parkin were significantly upregulated after 12 h of PM2.5 exposure,
consistent with the time that ferrous iron appeared co-localization with
mitochondria (Fig. 6G and H). Meanwhile, the LC3II/I ratio and p62
expression gradually increased with prolonged exposure to PM2.5 in
AC16 cells (Fig. 6I and J). Theses above evidence collectively
demonstrated that PM2.5 triggered iron homeostasis imbalance, which
mediated the crosstalk of ferritinophagy and mitophagy, and gradually
exacerbated iron overload in AC16 cells. In addition, during continuous
PM2.5 exposure, with the gradual elevation of NCOA4 in AC16 cells, the
expression of FTH1 and FTMT displayed a pattern of first increasing and
then decreasing (Fig. 6K). It reminded us that NCOA4 played an
important role in iron homeostasis imbalance caused by PM2.5.
3.7. NCOA4-mediated iron homeostasis imbalance is responsible for
PM2.5-triggered ferroptosis
To further explore the regulatory role of NCOA4 in iron metabolism
mediated the crosstalk of ferritinophagy and mitophagy, the stable
NCOA4 knockdown AC16 cell lines were established through shRNA
transfection. As shown in Fig. 7F, the qRT-PCR results exhibited 92%
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Redox Biology 72 (2024) 103158
knockdown efficiency of NCOA4 mRNA. The results of immunofluo­
rescence showed that NCOA4 knockdown significantly improved FTH1
degradation caused by PM2.5 (Fig. 7A). Meanwhile, it was observed that
the upregulation of PINK1 and Parkin in response to PM2.5 exposure was
significantly alleviated by knocking down NCOA4 (Fig. 7B). Subse­
quently, the effects of NCOA4 knockdown on iron distribution and
ferroptosis-related molecules in AC16 cells were determined. Unlike the
effects observed in WT AC16 cells, it was found that the abnormal
expression of DHODH and COX-2 were significantly attenuated in
NCOA4 knockdown AC16 cells exposed to PM2.5 (Fig. 7C and D). In
addition, laser confocal imaging of FerroOrange probe staining also
demonstrated effective inhibition of iron homeostasis imbalance
through knockdown of NCOA4 (Fig. 7E).
mitochondrial injury were aggravated after exposure to PM2.5 in this
study (Fig. 1). The increased content of ferrous iron enhanced the
accumulation of intracellular lipid hydroperoxides by Fenton chemistry
and subsequently triggered ferroptosis [33]. Consistent with our find­
ings, Menon et al. observed iron overload in mice with cardiac ferrop­
tosis [34]. Meanwhile, the accumulation of 4-HNE and MDA, as well as
the consumption of GSH were detected in the hearts of mice undergoing
ferroptosis [35,36], which were also observed in this study. Under TEM,
typical ultrastructural features of ferroptosis (mitochondrial shrinkage,
disappearance of cristae and membrane rupture) were captured. Addi­
tionally, exposure to PM2.5 resulted in abnormal expression of DHODH
(key molecule of ferroptosis defense system) and COX-2. The above
evidence suggested that ferroptosis was triggered in PM2.5-induced
cardiac hypertrophy. Furthermore, the potential mechanism was
explored through the detection of relevant molecules. PM2.5 disrupted
the expression of various iron metabolism-related proteins in mice heart,
such as NCOA4, FTH1 and FTMT (Fig. 2). Alterations of iron metabolism
determined the sensitivity to ferroptosis by promoting the generation of
lipid hydroperoxides, which were able to injure the phospholipid bilayer
membrane of cells. Meanwhile, cardiac mitophagy was activated in mice
exposed to PM2.5, accompanied by mitochondrial dynamics disorders.
Mitochondria are the most abundant organelles in heart, which are
involved in the storage of redox-active Fe2+. Mitochondrial damage
exacerbated the accumulation of labile iron in the cytoplasm, thereby
increasing the sensitivity of myocardial cells to ferroptosis [37]. The
above results collectively indicated that the two important selective
autophagy (ferritinophagy & mitophagy) contributed to cardiac fer­
roptosis triggered by PM2.5.
Consistent with in vivo experiments, PM2.5 also triggered ferroptosis
in AC16 cells (Fig. 3). Meanwhile, ferritinophagy and mitophagy were
activated in the PM2.5-treated group. Furthermore, Fer-1 alleviated the
decreased cell viability caused by PM2.5, while declining the intracel­
lular accumulation and intramitochondrial mobilization of ferrous iron
(Fig. 3). In fact, it has been proved that PM2.5 promoted ferroptosis in
human retinal microvascular endothelial cells (HRMEC), leading to
vascular dilatation and injury to the inner blood-retinal barrier (iBRB)
[38]. All of the above evidence provided support for our findings.
Dihydroorotate dehydrogenase (DHODH) is a key molecule in the
CoQH2 ferroptosis defense system that can inhibit mitochondrial lipid
peroxidation and ferroptosis by promoting the reduction of CoQ to
CoQH2 (independent of GPX4 or FSP1) [32]. As a derivative of coen­
zyme Q10, MitoQ exerts antioxidant effects by selectively accumulating
in mitochondria. MitoQ supplementation provided substrates for the
DHODH-CoQH2 ferroptosis defense system, enhancing mitochondrial
resistance to lipid peroxidation [39]. Our study confirmed for the first
time that MitoQ effectively attenuated PM2.5-induced cardiac ferroptosis
(Fig. 4).
Based on the abnormal accumulation and distribution of labile iron
observed in vivo and in vitro, the regulatory role of PM2.5-induced iron
homeostasis imbalance in ferroptosis was further explored. As shown in
Fig. 5, the abnormal expression of iron regulatory molecules (NCOA4,
FTH1) suggested iron homeostasis imbalance and activation of ferriti­
nophagy after exposure to PM2.5; meanwhile, it is worth noting that
mitophagy was activated, accompanied by mitochondrial dysfunction.
Currently, researches have shown that both ferritinophagy and
mitophagy are important inducements of ferroptosis [16,40,41]. How­
ever, there was still a lack of research on how the two selective auto­
phagy were coordinated. Subsequently, it was observed that the
colocalization of ferrous iron with lysosomes occurred before mito­
chondria, indicating that the activation of ferritinophagy preceded
mitophagy through time series exposure of PM2.5 (Fig. 6). It reminded us
that abnormal iron metabolism drove the crosstalk of ferritinophagy and
mitophagy. In addition, with the prolonged time of PM2.5 exposure,
FTH1 and FTMT in AC16 cells were significantly downregulated after a
brief compensatory increase with the upregulation of NCOA4. There­
fore, we speculated that NCOA4 played a central role in the crosstalk of
4. Discussion
Mounting epidemiological studies have reported the adverse effects
of exposure to ambient particulate matter on CVD [21,22]. It has been
clearly stated that PM2.5 is one of the major modifiable risk factors
relevant to cardiovascular events in a Joint Opinion from the World
Heart Federation [23]. Therefore, it is crucial for the prevention and
management of CVD to elucidate the potential mechanisms of cardio­
vascular toxicity of PM2.5. This study demonstrated that iron homeo­
stasis imbalance caused by PM2.5 evoked cardiac hypertrophy through
ferroptosis for the first time. In addition, it was found that the mecha­
nism by which PM2.5 triggered ferroptosis was cardiac iron homeostasis
imbalance mediated the crosstalk of ferritinophagy and mitophagy.
A recent population evidence has shown that PM2.5 is correlated with
impairments of cardiac structure and function [8]. In this study, cardiac
hypertrophy was observed in mice exposed to PM2.5 (Fig. 1). Through
echocardiography, changes in left ventricular wall thickness, and vol­
ume brought on by PM2.5 were detected, which triggered systolic and
diastolic dysfunction. The results were consistent with our previous
findings [24]. Left ventricular mass was adopted clinically to determine
cardiac hypertrophy. Cardiac hypertrophy is characterized by individual
cardiomyocyte growth in both length and width [25]. Fecht et al. found
that the elevation of PM2.5 concentration by one interquartile range was
positively correlated with a 2.9 g/m2 increase in left ventricular mass,
which provided support for our findings [26]. A recent study observed
that PM2.5 could lead to histopathological features of cardiac hyper­
trophy in mice [27]. In addition, Yue et al. pointed out that short-term
exposure to PM2.5 exacerbated myocardial cell hypertrophy in heart
failure (HF) mice compared with HF mice treated with the filtered air
[28]. The exposure chamber used in Yue’s study and the intratracheal
instillation used in our study are both commonly used exposure methods
in toxicology mechanism research. The exposure chamber can better
simulate real-world exposure, which is a limitation of our exposure
model [28]. In comparison, the intratracheal instillation was deter­
mined based on the recommended annual AQG level, which has more
stable exposure dose, and can better evaluate the dose-response rela­
tionship. Our results confirmed the findings of the above studies. Mice
exposed to PM2.5 also exhibited pathological injury such as increased
cardiomyocyte cell volume and disordered arrangement (Fig. 1C).
Meanwhile, expression of fetal genes such as BNP, ANP, skeletal muscle
α-actin and MYH7 are ordinarily observed in pathological hypertrophy
[29]. In line with other studies, it was observed that ANP and BNP were
reactivated in PM2.5-induced cardiac hypertrophy [30].
It has been observed that feeding mice high-iron diet induced hy­
pertrophic cardiomyopathy, with typical features of ferroptosis [31]. In
addition, Shi et al. identified that ferroptosis can promote pathological
cardiac remodeling and dysfunction, ultimately leading to cardiac hy­
pertrophy [10]. The prerequisites of ferroptosis consisted of disrupted
iron metabolism, impaired lipid peroxidation repair capacity and
abnormal mitochondrial metabolism [32]. A recent study reported that
PM2.5 triggered ferroptosis in respiratory systems [17]. Similarly, it was
found that the elevated iron content, lipid peroxidation and
10
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Redox Biology 72 (2024) 103158
Fig. 8. The schematic of PM2.5 promoting cardiac hypertrophy via triggering cardiomyocyte ferroptosis.
ferritinophagy and mitophagy. The subsequent results confirmed that
NCOA4 knockdown significantly inhibited the consumption of FTH1,
while alleviating the activation of mitophagy caused by PM2.5. In
addition, the characteristic ferroptosis events in response to PM2.5 were
attenuated in NCOA4 knockdown AC16 cells. The above results collec­
tively demonstrated that iron homeostasis imbalance mediated the
crosstalk of ferritinophagy and mitophagy was essential for PM2.5-in­
duced cardiac ferroptosis, in which NCOA4 played an important role
(Fig. 8). A recent study observed that PM2.5 up-regulated the level of
oxidative stress-related genes and ROS production [42]. ROS was
observed to be significantly elevated in AC16 cells of PM2.5 group in this
study. It has been reported that ROS plays a crucial role in triggering
NCOA4-mediated ferritinophagy [12,43]. In addition, it has been re­
ported that Fe accumulation can also regulate the activation of ferriti­
nophagy by affecting the expression of NCOA4 [44]. Although Fe cannot
be isolated from PM2.5 through existing technology, our previous study
has shown that the content of Fe in PM2.5 is 11.28 ± 2.61 mg/g [19].
Therefore, the accumulation of ROS and Fe may both mediate the
increased level of NCOA4 induced by PM2.5, followed by activation of
subsequent cascades. In addition, some study has observed the changes
in toxicity of PM2.5 through photochemical reactions [45], which is a
limitation of PM2.5 disinfection via UV irradiation. Therefore, more
studies are encouraged to investigate on this issue in future.
reducing the risk and burden of CVD.
5. Conclusion
Acknowledgements
In summary, it was found that exposure to PM2.5 resulted in cardiac
hypertrophy through DHODH-mediated ferroptosis. The crosstalk of
NCOA4-mediated ferritinophagy and PINK1-mediated mitophagy
driven by PM2.5-triggered iron homeostasis imbalance was elucidated
for the first time. In addition, there was a protective effect of MitoQ on
PM2.5-induced cardiac toxicity. Therefore, our data will provide new
target and treatment strategy for the prevention and management of
CVD related to air pollution, which has important implications for
This work was supported by the National Key R&D Program of China
(2022YFA0806900), National Natural Science Foundation of China
(82273659, 82330101).
CRediT authorship contribution statement
Tianyu Li: Writing – original draft, Visualization, Software, Meth­
odology, Data curation, Conceptualization. Mengqi Sun: Writing –
original draft, Software. Qinglin Sun: Software, Methodology. Xiaoke
Ren: Software, Methodology. Qing Xu: Software, Methodology, Data
curation. Zhiwei Sun: Writing – review & editing, Supervision, Project
administration, Funding acquisition, Conceptualization. Junchao
Duan: Writing – review & editing, Supervision, Methodology, Funding
acquisition, Conceptualization.
Declaration of competing interest
The authors declare that they have no known competing financial
interests or personal relationships that could have appeared to influence
the work reported in this paper.
Data availability
Data will be made available on request.
Appendix A. Supplementary data
Supplementary data to this article can be found online at https://doi.
org/10.1016/j.redox.2024.103158.
11
T. Li et al.
Redox Biology 72 (2024) 103158
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