Abstract

   Atherosclerosis is a chronic inflammatory disease and a major
   pathological basis of numerous cardiovascular conditions, with a high
   global mortality rate. Macrophages play a pivotal role in its
   pathogenesis through phenotypic switching and foam cell formation.
   Prostaglandin E2 receptor subtype 4 (EP4) highly expressed on the
   macrophage surface, is involved in various pathophysiological
   processes, such as inflammation and lipid metabolism. However, the role
   of macrophage EP4 in the progression of atherosclerosis remains
   unclear. To determine whether macrophage EP4 affects the progression of
   atherosclerosis by regulating foam cell formation and macrophage
   polarization. Myeloid-specific EP4 knockout mice with an ApoE-deficient
   background were fed a Western diet for 16 weeks. Our results showed
   that EP4 expression was significantly downregulated during
   atherosclerosis. EP4 deficiency was found to exacerbate atherosclerotic
   plaque formation and destabilizes plaques. In vitro studies further
   demonstrated that loss of EP4 in myeloid cells promoted foam cell
   formation and M1 macrophage polarization. Both transcriptomic and
   proteomic analysis showed that EP4 may regulate these processes by
   regulating CD36 expression in macrophage, which was further confirmed
   by Western blot and qPCR. In summary, deficiency of EP4 receptor in
   macrophages enhance foam cell formation and M1 polarization by
   upregulating CD36 expression, thereby accelerating the progression of
   atherosclerosis.

   Keywords: EP4, macrophage polarization, foam cell, CD36,
   atherosclerosis

1. Introduction

   Atherosclerosis (AS) is a chronic and progressive vascular disease
   characterized by the accumulation of plaques within the arterial wall.
   The plaques, composed of lipids, cholesterol, calcium, cellular debris,
   and fibrous tissue, gradually narrow and stiffen the arteries, thereby
   reducing blood flow to vital organs [[40]1]. With a complex
   pathogenesis, AS serves as the underlying cause of various
   cardiovascular diseases, including coronary artery disease (e.g.,
   myocardial infarction, cerebrovascular disease (e.g., ischemic stroke),
   and peripheral artery disease, each contributing substantially to
   global morbidity, disability, and healthcare burdens
   [[41]2,[42]3,[43]4]. As a result, AS remains the leading cause of death
   worldwide, accounting for approximately 17.9 million deaths annually.

   Macrophages have been shown to play an important role in the
   pathogenesis of AS [[44]5]. They engulf oxidized low-density
   lipoproteins (oxLDL) via scavenger receptors, resulting in
   intracellular lipid accumulation and foam cell formation hallmark of
   early atherosclerotic lesions. The process disrupts lipid homeostasis
   and contributes to AS progression [[45]6,[46]7]. In addition, the
   inflammatory cytokines secreted by macrophages and the associated
   immune response also influence the development of plaque. The ratio of
   M1 and M2 macrophages within the plaques is the key determinant of
   plaque fate [[47]8,[48]9]. Therefore, elucidating the mechanisms
   underlying macrophage foam cell formation and polarization is essential
   for the development of targeted therapies against AS.

   Prostaglandin E2 (PGE2) is a biologically active endogenous lipid
   molecule widely distributed in human tissues, involved in diverse
   physiological and pathological processes, such as inflammation
   [[49]10], smooth muscle contraction and relaxation [[50]11], and lipid
   metabolism [[51]12]. PGE2 exerts its effects through four G
   protein-coupled receptors termed Prostaglandin E receptor EP1, EP2, EP3
   and EP4. Among these, prostaglandin E2 receptor subtype 4 (EP4) is
   highly expressed in macrophages with high affinity, suggesting that
   many of the biological effects of PGE2 are mediated through EP4
   signaling [[52]13]. EP4 has been found to play a critical role in
   cardiovascular pathophysiology. EP4 deficiency impairs cardiac
   function, aggregates cardiomyocyte injury and necrosis [[53]14];
   exacerbates inflammatory response, promotes smooth muscle cell
   apoptosis, and contributes to the development of abdominal aortic
   aneurysms [[54]15]. In addition, EP4 deficiency has been associated
   with hypercholesterolemia [[55]16] and impaired cardiac fatty acid
   metabolism [[56]17], whereas EP4 agonists have been shown to prevent
   diet-induced hypercholesterolemia [[57]16] and improve cardiomyocyte
   function [[58]17]. Collectively, these findings suggest that EP4 helps
   maintain cardiovascular homeostasis by regulating inflammation and
   lipid metabolism. However, its precise mechanistic role in
   atherosclerosis needs to be further explored.

   In this study, we investigated the role of macrophage EP4 in the
   development of AS. EP4 expression was found to be downregulated in both
   atherosclerotic plaques and oxLDL-stimulated macrophages.
   Macrophage-specific EP4 knockdown promoted atherosclerotic lesion
   formation. EP4 deficiency enhanced macrophage foam cell formation and
   M1 polarization at least in part by upregulating CD36 expression.

2. Methods

2.1. Animals

   EP4^Flox mice were generated using the CRISPR-Cas9 system by inserting
   loxP site flanking exons 2 at Shanghai Model Organisms Center, Inc.
   (Shanghai, China). Lyz2-Cre mice were purchased from Jackson
   Laboratory. Myeloid-specific EP4 knockout mice (EP4^MKO) were generated
   by crossing EP4^Flox mice with Lyz2-Cre mice. ApoE^−/−EP4^Flox mice
   were generated by crossing ApoE^−/^− mice with EP4^Flox mice, and
   ApoE^−/^−EP4^MKO mice were subsequently generated by crossing
   ApoE^−/^−EP4^Flox mice with Lyz2-Cre mice. All genotypes were confirmed
   by qRT-PCR. All mice were group-housed 3–5/cage in the SPF animal
   facility. Four-week-old ApoE^−/^−EP4^Flox mice and ApoE^−/^−EP4^MKO
   mice were randomly selected for the experiment and assigned to groups,
   each group consisting of 6–8 mice, then fed either a Western diet (40%
   kcal% Fat, 1.25% Cholesterol, 0.5% Cholic, D12109C, Research Diets, New
   Brunswick, NJ, USA) or Chow diet for 16 weeks. For rescue experiments,
   all mice were fed Western diet for 12 weeks then mice were
   intraperitoneally injected with Sulfosuccinimidyl oleate sodium (SSO,
   25 mg/kg/week, HY-112847A, MCE, Monmouth Junction, NJ, USA), an
   irreversible inhibitor of CD36 that covalently binds to extracellular
   lysine residues of the receptor, thereby blocking its ligand-binding
   capacity, for 4 weeks while continuing the Western diet. The sample
   size of 6–8 mice per group was determined based on established
   standards in the field and compliance with the 3Rs principles to ensure
   both statistical robustness and ethical animal use. The order of
   removing treatment-group mice from each cage was systematically
   balanced across all cages to prevent any treatment group from
   consistently being handled first or last during the extraction process.
   Investigators were unblinded to treatment groups during compound
   administration but maintained blinding throughout later experimental
   stages and statistical evaluation. No survival procedures were
   performed; euthanasia was conducted under anesthesia without recovery.
   All animal experiments were approved by the Animal Care and Use
   Committee of Guangzhou Women and Children’s Medical Center, Guangzhou
   Medical University, China (RSDW-2023-01397).

2.2. Genotyping

   Genomic DNA was extracted from 2 mm tail biopsies of 4-week-old mice
   using the Direct Mouse Genotyping Kit Plus (K1027, APExBio, Houston,
   TX, USA). PCR was performed to amplify target region, and specific
   primers were designed to detect mouse alleles. The primer sequences are
   listed in [59]Table S1.

2.3. Macrophage Culture and Treatments

   Bone marrow-derived macrophages (BMDMs) were isolated from wild-type
   (WT), EP4^Flox, and EP4^MKO mice. Briefly, fresh bone marrow cells were
   cultured in DMEM (C11995500BT, Gibco, Grand Island, NY, USA) with 10%
   FBS (164210-50, Procell, Wuhan, China), 1% penicillin/streptomycin
   (15140-122, Gibco, Grand Island, NY, USA), and murine M-CSF (10 ng/mL,
   315-02, Peprotech, Cranbury, NJ, USA) for 7 days. Human
   monocyte-derived macrophages (HMDMs) were isolated from healthy donor
   PBMCs using magnetic-activated cell sorting (MACS), then differentiated
   for 7 days in M-CSF-containing medium (20 ng/mL). To ascertain EP4
   function, BMDMs were treated with CAY10580 (100 nM, HY-135259, MCE,
   Monmouth Junction, NJ, USA) for 24 h.

2.4. RNA Sequencing (RNA-Seq)

   WT BMDMs were either treated with (oxidized low-density lipoprotein)
   oxLDL or left untreated for 24 h. Total RNA was extracted using Trizol
   reagent, and RNA libraries were constructed using the NEBNext^®
   Ultra^TM RNA Library Prep Kit (E7770, NEB, Ipswich, MA, USA) and
   NEBNext^® Ultra^TM Directional RNA Library Prep Kit (E7760s, NEB,
   Ipswich, MA, USA), according to the manufacturer’s instructions. RNA
   sequencing was performed on an Illumina platform (NEB, Ipswich, MA,
   USA). Differential expression analysis was performed using the DESeq2 R
   package (1.16.1). Genes with an adjusted p-value (Padj) < 0.05 were
   considered differentially expressed. Functional enrichment analysis of
   the differentially expressed genes in KEGG pathways was performed using
   the cluster Profiler R package (v4.8.0). Padj < 0.05 was considered
   significantly enriched.

2.5. Proteomic Sequencing

   Proteomic analysis was performed on BMDMs isolated from EP4^Flox and
   EP4^MKO mice following treatment with oxLDL. Briefly, samples were
   lysed in SDT buffer (4%SDS, 100 Mm Tris-HCl, Ph 7.6), incubated in
   boiling water for 15 min, and centrifuged at 14,000× g for 15 min.
   Protein concentrations were determined using the BCA protein assay kit
   (P0012, Beyotime, Shanghai, China), and samples were stored at −80 °C
   until further analysis. To assess protein quality, 20 μg of protein
   from each sample was separated on a 12% SDS-PAGE gel and visualized
   using Coomassie Blue R-250 staining (BB-3720, Bestbio, Shanghai,
   China). For proteomic analysis, 100 μg of protein per sample was
   processed using the filter-aided sample preparation (FASP) method and
   subsequently lyophilized. The digested peptides were analyzed on a
   nanoElute UHPLC system (Bruker, Bremen, Germany) coupled to a timsTOF
   Pro mass spectrometer (Bruker, Bremen, Germany) equipped with a
   CaptiveSpray source. Raw mass spectrometry data were processed using
   MaxQuant software (v1.6.17.0, Max Planck Institute of Biochemistry,
   Martinsried, Germany). MS data were searched against the UniProt
   database. Label-free quantification (LFQ) was applied to determine
   based on normalized spectral protein intensities. Proteins with a Fold
   change >2 or <0.5 and p value (Student’s t test) < 0.05 were considered
   deferentially expressed proteins (DEPs). Pathway enrichment analysis
   was performed using KEGG Orthology And Links Annotation (KOALA)
   software (v V2.2, Kanehisa Laboratories, Kyoto University, Kyoto,
   Japan). The KEGG GENES database (version: KO_INFO_END.txt (2022.11.05)
   was used for aligning target protein sequences, which were subsequently
   annotated with KO label, and automatically classified into associated
   pathways.

2.6. Foam Cell Induction and Uptake Assay

   For foam cell induction, BMDMs and HMDMs were incubated with oxLDL (50
   μg/mL, YB-002, Yiyuan biotechnologies, Guangzhou, China) for 24 h.
   Cells were then fixed and stained with Oil Red O (G1015, Servicebio,
   Wuhan, China), and lipid accumulation was observed by light microscopy.
   For the ox-LDL uptake assay, cells were serum-starved in DMEM for 12 h,
   followed by incubation with Dil-labeled oxLDL (50 μg/mL, YB-001, Yiyuan
   biotechnologies, Guangzhou, China) for 4 h. Cells were then fixed with
   4% paraformaldehyde and stained with hoechst (H3570, Thermo Scientific,
   Waltham, MA, USA) for nuclei visualization. Fluorescence images were
   observed using confocal microscopy.

2.7. Macrophage Polarization

   To induce macrophage polarization, cells were cultured in complete DMEM
   (10% FBS). BMDMs were stimulated for 24 h under the following
   conditions: for M1 polarization, lipopolysaccharide (LPS, 100 ng/mL,
   Sigma, Burlington, MA, USA) and IFN-γ (20 ng/mL, 315-05, Peprotech,
   Cranbury, NJ, USA); for M2 polarization, IL4 (20 ng/mL, 214-14,
   Peprotech, Cranbury, NJ, USA) and IL13 (20 ng/mL, 210-13, Peprotech,
   Cranbury, NJ, USA). Expression of the phenotypic markers was analyzed
   by qRCR for TNF-α, Arg1, etc., and by flow cytometry using surface
   markers CD86 (M1) and CD206 (M2).

2.8. Histological Analyses

   For en face staining of atherosclerotic plaques, aortas were isolated
   from ApoE^−/−EP4^Flox mice and ApoE^−/−EP4^MKO mice after 16 weeks of
   Western diet feeding or Chow diet feeding. Then aortas were fixed in 4%
   paraformaldehyde for 24 h and stained with Oil Red O. En face images
   were acquired by light microscopy and analyzed using Image J software
   (v 1.53, NIH, Bethesda, MD, USA). For cross-sectional analysis, mouse
   aortic roots were fixed in 4% paraformaldehyde (G1101, Servicebio,
   Wuhan, China), embedded in Tissue-Tek O.C.T. compound (4583, SAKURA,
   Tokyo, Japan), and sliced into 6 μm sections. The sections were stained
   with hematoxylin and eosin (H&E), Masson’s trichrome, and Oil Red O.
   Histological images were acquired using a light microscope (Leica DM4,
   Wetzlar, Germany) and analyzed by Image J software (v 1.53, NIH,
   Bethesda, MD, USA).

2.9. Immunofluorescence

   For immunofluorescence staining of cultured macrophages, BMDMs and
   HMDMs were seeded into glass-bottomed culture dishes at a density of 2
   × 10^5 per well and incubated with oxLDL for 24 h. Cells were then
   fixed with 4% paraformaldehyde and incubated overnight at 4 °C with the
   following primary antibodies: Anti-CD68 (1:100, 25747-1-AP,
   Proteintech, Wuhan, China) and Anti-EP4 (1:100, 24895-1-AP,
   Proteintech, Wuhan, China). The next day, cells were incubated with
   Alexa Fluor 594-conjugated IgG antibody (1:200, ab150116, Abcam,
   Cambridge, UK) and Fluor 488-conjugated IgG antibody (1:200, ab150077,
   Abcam, Cambridge, UK) for 1 h at room temperature. Nuclei were stained
   with Hoechst (1:2000, H3570, Thermo Scientific, Waltham, MA, USA) and
   the images were captured using a Leica SP8 confocal microscope (Leica,
   Wetzlar, Germany), and analyzed with Image J software (v 1.53, NIH,
   Bethesda, MD, USA).

   For immunostaining of aortic tissue sections, frozen sections were
   blocked with PBS (SH30256.01B, Cytiva, Marlborough, MA, USA) containing
   5% normal goat serum (16210-64, Gibco, Grand Island, NY, USA) and 5%
   bovine serum albumin (B2064-100G, Sigma, Burlington, MA, USA) for 1 h
   at room temperature. Sections were then incubated overnight at 4 °C
   with the following primary antibodies: anti-EP4, anti-CD36 (18836-1-AP,
   Proteintech, Wuhan, China), anti-Arg1 (16001-1-AP, Proteintech, Wuhan,
   China), anti-F4/80 (GB113373-100, Servicebio, Wuhan, China), anti-α-SMA
   (ab7818, Abcam, Cambridge, UK). The next day, the slides were incubated
   with Alexa Fluor 488-conjugated IgG secondary antibody (1:200,
   ab150077, Abcam, Cambridge, UK) or Alexa Fluor 594-conjugated IgG
   secondary antibody (1:200, ab150160, Abcam, Cambridge, UK) at 37 °C for
   2 h. Nuclei were visualized with DAPI (H-1200, Vector Laboratories,
   Burlingame, CA, USA). Immunofluorescence images were captured by Leica
   SP8 confocal microscopy (Leica, Wetzlar, Germany). Fluorescence
   intensity quantification was performed using ImageJ software (v 1.53,
   NIH, Bethesda, MD, USA). The mean fluorescence intensity (MFI) for each
   region of interest (ROI) was calculated, with group-level MFI
   representing the average of all matched sections.

2.10. Flow Cytometry Analysis

   BMDMs were resuspended in PBS with 2% FBS. To assess macrophage
   polarization following stimulation, cells were first incubated with
   anti-Mouse CD16/CD32 monoclonal antibody (1:200, 14-0161-85,
   Invitrogen, Carlsbad, CA, USA) for 15 min to block Fc receptors, then
   were stained with anti-CD11b-PE-Cy7 (1:200, 101216, Biolegend, San
   Diego, CA, USA), anti-F4/80-FITC (1:200, 123108, Biolegend, San Diego,
   CA, USA), anti-CD86-PE (1:200, 105007, Biolegend, San Diego, CA, USA),
   and anti-CD206-APC (1:200, 321110, Biolegend, San Diego, CA, USA) for
   30 min on ice. Then, cell suspensions were filtered through a cell
   strainer-cap FACS tube and vortex-mixed at medium speed for 10 s
   immediately before flow cytometry analysis to ensure uniform cell
   distribution. Data were analyzed by Flowjo software (v 10.6.2, BD,
   Franklin Lakes, NJ, USA).

2.11. RNA Extraction and RT-qPCR

   Total RNA was extracted from cells using a RNeasy kit (74104, Qiagen,
   Hilden, Germany), according to the manufacturer’s instructions. Then,
   the extracted RNAs were reverse-transcribed using a PrimeScriptTM RT
   kit (RR037A, Takara, Kusatsu City, Japan). Quantitative real-time PCR
   was performed using the SYBR-Green (RR820A, Takara, Kusatsu City,
   Japan) on Applied Biosystems Q6 FastReal-Time PCR System sequencer
   detector (Thermo Scientific, Waltham, MA, USA). The primers are listed
   in [60]Table S2. Gene expression was normalized to mouse β-actin or
   human GAPDH.

2.12. Western Blot Analysis

   Total protein was extracted using radioimmunoprecipitation assay lysis
   buffer (P0013, Beyotime, Shanghai, China) supplemented with a protease
   inhibitor cocktail (539131, Merck, Darmstadt, Germany). Protein
   concentrations were determined using a BCA Assay Kit (P0009, Beyotime,
   Shanghai, China). Equal amounts of protein lysates were separated on
   10% SDS-PAGE gels and transferred to PVDF membranes. Following blocking
   with 5% skimmed milk in TBST for 1 h at room temperature, the membranes
   were incubated overnight at 4 °C with the following primary antibodies:
   anti-EP4 (1:1000, 24895-1-AP, Proteintech, Wuhan, China), anti-CD36
   (1:2000, ab133625, Abcam, Cambridge, UK), anti-Arg1 (1:1000,
   16001-1-AP, Proteintech, Wuhan, China), anti-iNOS (1:1000, ab178945,
   Abcam, Cambridge, UK), anti-Hsp90 (1:10,000, ab203126, Abcam,
   Cambridge, UK), anti-GAPDH (1:10,000, ab8245, Abcam, Cambridge, UK).
   After washing, membranes were incubated with the secondary antibodies
   for 1 h at room temperature. Protein bands were visualized using an
   enhanced chemiluminescence (ECL) detection kit (WBKLS0500, Merck,
   Darmstadt, Germany) and quantified using ImageLab software (v 4.0,
   Bio-rad, Hercules, CA, USA).

2.13. siRNA Transfection

   Mouse CD36 siRNA and human EP4 siRNA were transiently transfected into
   cells using Lipofectamine^TM RNAiMAX Transfection Reagent (13778150,
   Thermo Fisher, Waltham, MA, USA), according to the manufacturer’s
   instructions. A non-targeting siRNA was used as a negative control
   ([61]Table S3). The knockdown efficiency was assessed by quantitative
   PCR and Western blot analysis.

2.14. Statistical Analysis

   Data were presented as mean ± SEM, and the statistical analysis was
   performed using GraphPad Prism software (v8.3.0, GraphPad Software, San
   Diego, CA, USA). Exclusion criteria included spontaneous death or
   failure to meet experimental endpoints. The normality of data was
   checked using the Shapiro–Wilk normality test. For comparison with two
   groups, an unpaired two-tailed Student’s t-test was used if the data
   passed the normality test. For comparisons involving more than two
   groups, One-way ANOVA followed by Tukey’s multiple comparisons was
   used. Two-way ANOVA with Tukey’s multiple comparisons test was employed
   for multifactorial comparisons involving three or more groups. The
   difference with p < 0.05 was considered statistically significant.

3. Results

3.1. Macrophage EP4 Expression Was Reduced During Atherosclerosis

   To investigate genes potentially involved in the pathological
   progression of atherosclerosis, we established an in vitro cell model
   using bone marrow-derived macrophages (BMDM) stimulated with oxidized
   low-density lipoprotein (oxLDL). Following 24 h of oxLDL stimulation,
   BMDMs were collected for RNA-sequencing. Compared to the control group,
   a total of 520 differentially expressed genes (DEGs) were identified,
   including 96 upregulated genes and 424 downregulated genes ([62]Figure
   1A). Subsequent gene set enrichment analysis (GSEA) and KEGG pathway
   enrichment analysis revealed that these DEGs were significantly
   enriched in inflammatory response pathways, such as human
   cytomegalovirus infection, human papillomavirus infection, and
   inflammatory mediator regulation of TRP channels ([63]Figure 1B). By
   intersecting the gene sets enriched in the three pathways, two
   candidate genes, Ptger4 (EP4) and plcb1, were identified ([64]Figure
   1C,D), among which EP4 exhibited the highest fold change in RNA
   sequencing. The downregulation of the EP4 protein ([65]Figure 1E,G) and
   mRNA ([66]Figure 1F,H) was further confirmed in both BMDMs and HMDMs
   following oxLDL stimulation in vitro. To confirm the findings in vivo,
   atherosclerosis was induced in ApoE^−/− mice by feeding a Western diet
   for 16 weeks. Immunofluorescence staining revealed significantly
   reduced EP4 expression in macrophages from Western diet-fed mice,
   compared to those fed on Chow diet ([67]Figure 1I). Collectively, these
   results indicate that EP4 expression is downregulated in macrophages
   during atherogenesis.

Figure 1.

   [68]Figure 1
   [69]Open in a new tab

   Macrophage EP4 expression was reduced in oxLDL-treated macrophages and
   atherosclerotic lesions. BMDMs were stimulated with oxLDL for 24 h and
   collected for RNA sequencing. (A) The histogram depicting DEGs in
   wild-type BMDMs treated with or without oxLDL. (B) KEGG pathway
   enrichment analysis of DEGs. (C) The gene sets enriched in each KEGG
   inflammation-related pathway. (D) Schematic diagram illustrating the
   overlap of DEGs among KEGG inflammation-related signaling pathways. Bar
   graph showing downregulation of ptger4 (EP4) expression following
   treatment with oxLDL. Immunofluorescence staining of EP4 (red) and CD68
   (green) in BMDMs (E) and HMDMs (G) cultured with oxLDL (50 μg/mL) for
   24 h was performed. Nuclei were stained with DAPI (blue). The
   fluorescence intensity of EP4 was quantified by ImageJ (n = 6). Scale
   bar: 20 μm. Quantitative RT-PCR analysis of EP4 mRNA expression in
   BMDMs (F) and HMDMs (H) cultured with oxLDL (50 μg/mL) for 24 h (n =
   6). (I) Immunofluorescence staining of EP4 (red) and CD68 (green)
   within the plaque lesions from ApoE^−/− mice after 16 weeks of Western
   diet or Chow diet (n = 5). EP4 fluorescence intensity was quantified by
   ImageJ. Nuclei were stained with DAPI (blue). Scale bar: 20 μm. Data
   are presented as mean ± SEM, ** p < 0.01, *** p < 0.001 versus control
   group, Unpaired t-test. DEGs: differentially expressed genes; BMDM:
   bone marrow-derived macrophage; HMDM: human monocyte-derived
   macrophage; oxLDL: oxidized low-density lipoprotein; WD: Western diet.

3.2. Macrophage EP4 Deficiency Exacerbated Atherosclerosis and Destabilizes
Plaques

   To further investigate the role of macrophage EP4 in atherosclerosis,
   we generated myeloid-specific EP4 knockout mice on an ApoE^−/^−
   background (ApoE^−/^− EP4^MKO mice) ([70]Figure S1A–C). Efficient
   Cre-mediated recombination at the locus of Ptger4 was confirmed in
   BMDMs isolated from EP4^MKO mice ([71]Figure S1D,E). Notably, compared
   to ApoE^−/^−EP4^Flox mice, ApoE^−/^− EP4^MKO mice exhibited
   significantly larger atherosclerotic plaque areas ([72]Figure 2A,C,E)
   and increased lipid deposition ([73]Figure 2B,D) within the aortic
   root, suggesting that EP4 deficiency exacerbates plaque formation and
   intraplaque lipid accumulation.

Figure 2.

   [74]Figure 2
   [75]Open in a new tab

   Macrophage EP4 deficiency promoted plaque formation and instability in
   atherosclerosis. ApoE^−/−EP4^Flox and ApoE^−/−EP4^MKO mice were fed
   either with Western diet or Chow diet for 16 weeks. Total aorta and
   aortic roots were collected. (A) Representative En face images and
   quantification of Oil Red O-stained aorta from ApoE^−/−EP4^Flox and
   ApoE^−/−EP4^MKO mice (Chow: n = 6; WD: n = 6). Representative images
   and quantification of Oil Red O staining ((B,D), scale bar: 200 μm),
   H&E staining ((C,E), scale bar: 200 μm), Masson’s trichrome staining
   ((F,G), scale bar: 200 μm) in aortic root sections from
   ApoE^−/−EP4^Flox and ApoE^−/−EP4^MKO mice after 16 weeks of Western
   diet. Immunofluorescence staining and quantitation of α-SMA (the smooth
   muscle cell marker) (H,J) and CD68 (the macrophage marker) (I,K) in the
   aortic roots from ApoE^−/−EP4^Flox and ApoE^−/−EP4^MKO mice, the
   fluorescence intensity was quantified by ImageJ (n = 6), α-SMA stained
   as red, CD68 as green, nuclei as blue. Scale bar: 20 μm. Data presented
   as mean ± SEM, ns: not significant, * p < 0.05, ** p < 0.01, *** p <
   0.001, **** p < 0.0001, versus the ApoE^−/−EP4^Flox mice, two-way ANOVA
   in (A), Unpaired t-test in (D,E,G,J,K). WD: Western diet.

   Plaque stability is a critical determinant of clinical outcomes during
   atherosclerosis, as unstable plaques are prone to rupture, leading to
   thrombosis and acute cardiovascular events [[76]18]. To investigate
   whether macrophage EP4 deficiency affects plaque stability, collagen
   content within plaques was examined using Masson’s trichrome staining
   (red, muscle fibers; blue, collagen). As shown in [77]Figure 2F,G,
   ApoE^−/−EP4^MKO mice showed a significant reduction in collagen
   content, suggesting compromised plaque stability. In addition, plaques
   from ApoE^−/^−EP4^MKO mice displayed decreased expression of α-SMA
   ([78]Figure 2H,J) and increased infiltration of CD68-positive
   macrophages ([79]Figure 2I,K). Collectively, these data demonstrate
   that macrophage EP4 deficiency promotes atherosclerotic plaque
   progression and destabilization, highlighting its critical role in both
   lesion development and plaque stability.

3.3. EP4 Deficiency Promoted Macrophage M1 Polarization Both In Vitro and In
Vivo

   Macrophage phenotypic switching plays a critical role in the
   progression of atherosclerosis. Accumulating evidence indicates that
   unstable plaques are enriched with pro-inflammatory M1 macrophages,
   whereas stable plaques predominantly contain anti-inflammatory M2
   macrophages [[80]19]. To determine whether macrophage EP4 deficiency
   affects phenotypic switching, EP4^MKO BMDMs were subjected to M1-
   (LPS/IFN-γ) or M2- (IL4/IL13) polarizing conditions. Upon LPS/IFN-γ
   stimulation, EP4^MKO BMDMs exhibited increased CD86 expression, a
   surface marker of M1 macrophages ([81]Figure 3A), whereas CD206 (M2
   surface marker) remained unchanged upon IL4/IL13 stimulation
   ([82]Figure 3B). RT-qPCR analysis further confirmed upregulation of
   M1-associated genes (iNOS, TNFα, IL6, and CCL2) in EP4^MKO BMDMs upon
   LPS/IFN-γ stimulation ([83]Figure 3C), and downregulation of M2 markers
   (CD206, Arg-1) upon IL4/IL13 stimulation ([84]Figure 3D). Western blot
   analysis corroborated these findings, showing increased iNOS and
   decreased Arg-1 protein levels in EP4^MKO BMDMs ([85]Figure 3I).

Figure 3.

   [86]Figure 3
   [87]Open in a new tab

   EP4 deficiency promoted macrophage M1 polarization both in vitro and in
   vivo. (A,B) Flow cytometry analysis of CD86 and CD206 expression in
   BMDMs from EP4^Flox and EP4^MKO mice following stimulation with
   LPS/IFNγ or IL4/IL13 (n = 6). (C,D) The mRNA level of inflammation
   cytokines and macrophage polarization markers in BMDMs from EP4^Flox
   and EP4^MKO mice after stimulation with LPS/IFNγ or IL4/IL13 was
   determined by qRT-PCR (n = 6). Immunofluorescence staining and
   quantification of iNOS (E,G) and Arg1 (F,H) in the aortic roots from
   ApoE^−/−EP4^Flox and ApoE^−/−EP4^MKO mice after 16 weeks of Western
   diet; the fluorescence intensity was quantified by ImageJ (n = 4).
   Nuclei were stained with DAPI (blue); iNOS or Arg1 was stained red;
   F4/80 as green. Scale bar: 20 μm. (I) Western blot analysis and
   quantification of Arg1 and iNOS expression in BMDMs from EP4^Flox and
   EP4^MKO mice (n = 6). Data presented as mean ± SEM, ns: not
   significant, * p < 0.05, ** p < 0.01, **** p < 0.0001 versus the
   ApoE^−/−EP4^Flox mice or EP4^Flox mice, Unpaired t-test. BMDM: bone
   marrow-derived macrophage.

   In vivo, aortic root sections from ApoE^−/^−EP4^MKO mice and
   ApoE^−/−EP4^Flox mice fed a Western diet for 16 weeks were analyzed.
   Immunofluorescence staining revealed increased expression of iNOS (M1
   marker) ([88]Figure 3E,G) and reduced Arg1 (M2 marker) ([89]Figure
   3F,H) in plaques from ApoE^−/^−EP4^MKO mice, indicating a shift toward
   M1 polarization. Taken together, these data demonstrate that
   macrophage-specific EP4 deficiency promotes M1-polarization both in
   vitro and in vivo, contributing to a pro-inflammatory phenotype in
   atherosclerotic lesions.

3.4. Macrophage EP4 Deficiency Enhanced Foam Cell Formation

   To investigate the potential role of EP4 in foam cell formation, BMDMs
   derived from EP4^MKO and EP4^Flox mice were treated with oxLDL. Oil Red
   O staining showed that EP4 deficiency significantly enhanced foam cell
   formation and intracellular lipid accumulation compared to EP4^Flox
   BMDMs ([90]Figure 4A–C). To determine whether this phenotype was driven
   by increased uptake of modified lipoproteins, a Dil-labeled oxLDL
   uptake assay was performed. Immunofluorescence analysis showed that
   increased internalization of Dil-oxLDL in EP4^MKO BMDMs compared to
   EP4^Flox controls ([91]Figure 4D). To validate these findings in human
   cells, HMDMs were transfected with EP4-siRNA ([92]Figure S2). EP4
   knockdown significantly increased uptake of Dil-oxLDL in HMDMs compared
   to those transfected with negative control (NC)-siRNA ([93]Figure 4E).
   Collectively, these data suggest that EP4 deficiency enhances ox-LDL
   uptake and promotes foam cell formation in both murine and human
   macrophages.

Figure 4.

   [94]Figure 4
   [95]Open in a new tab

   Macrophage EP4 deficiency promotes foam cell formation. (A)
   Representative images and quantitation of Oil Red O-stained foam cells
   in EP4^Flox and EP4^MKO BMDMs after culture with or without oxLDL (50
   μg/mL) (n = 6), scale bar: 50 μm. (B,C) Quantification of total
   cholesterol and free cholesterol accumulation in EP4^Flox and EP4^MKO
   BMDMs treated with or without oxLDL for 24 h (n = 6). Representative
   images and quantitation of Dil-labeled oxLDL uptake in BMDMs (D) (n =
   7) and HMDMs (E) (n = 4). Nuclei were stained with DAPI (blue); EP4 was
   stained green. The fluorescence intensity was quantified by ImageJ,
   scale bar: 20 μm. Data presented as mean ± SEM, ns: not significant, *
   p < 0.05, ** p < 0.01, *** p < 0.001, versus EP4^Flox BMDMs or NC-siRNA
   transfected HMDMs, Unpaired t-test. BMDM: bone marrow-derived
   macrophage; HMDM: human monocyte-derived macrophage.

3.5. Activation of EP4 Suppresses Macrophage M1 Polarization and Foam Cell
Formation

   To further explore the functional role of macrophage EP4 in
   atherosclerosis, WT BMDMs were pre-treated with CAY10580, followed by
   stimulation with LPS + IFN-γ and IL4 + IL13. Flow cytometry analysis
   showed a decreased proportion of CD86^+ macrophages after CAY10580
   treatment ([96]Figure 5A), whereas the proportion of CD206^+
   macrophages remained unchanged ([97]Figure 5B). Consistent with these
   findings, qRT-PCR analysis showed that CAY10580 treatment attenuated
   expression of M1-associated pro-inflammatory factors upon LPS and IFN-γ
   stimulation ([98]Figure 5C) while enhancing the expression of
   M2-associated anti-inflammatory factors following IL4 and IL13
   stimulation ([99]Figure 5D).

Figure 5.

   [100]Figure 5
   [101]Open in a new tab

   Activation of EP4 suppressed macrophage M1 polarization and foam cell
   formation. (A,B) Flow cytometry analysis of CD86 and CD206 expression
   in BMDMs treated with Vehicle (0.00068% ethanol) or CAY10580 (100 nM)
   (n = 6). (C,D) qRT-PCR was used to determine the mRNA levels of
   pro-inflammatory cytokines and macrophage polarization markers in BMDMs
   pretreated with Vehicle (0.00068% ethanol) or CAY10580 (100 nM),
   followed by stimulation with LPS/IFN-γ or IL4/IL13 (n = 6). (E) Oil Red
   O staining and quantification of lipid accumulation in WT BMDMs
   pre-treated with Vehicle (0.00068% ethanol) or CAY10580 (100 nM)
   followed by stimulation with or without oxLDL (50 μg/mL) for 24 h (n =
   6), scale bar: 50 μm; (F) representative images and quantitation of
   Dil-labeled oxLDL uptake in BMDMs pretreated with Vehicle (0.00068%
   ethanol) or CAY10580 (n = 6), scale bar: 20 μm; data presented as mean
   ± SEM, ns: not significant, * p < 0.05, ** p < 0.01, **** p < 0.0001,
   Unpaired t-test, BMDM: bone marrow-derived macrophage.

   Then, WT BMDMs were pre-treated with CAY10580, followed by stimulation
   with oxLDL. Compared to cells stimulated with oxLDL alone,
   pre-activation of EP4 with CAY10580 significantly reduced foam cell
   formation ([102]Figure 5E) and decreased Dil-oxLDL uptake ([103]Figure
   5F). These results indicate that pharmacological activation of EP4
   suppresses foam cell formation and M1 polarization, thereby promoting
   an anti-inflammatory macrophage phenotype, which may potentially
   ameliorate atherosclerosis.

3.6. EP4 Upregulates the Expression of CD36

   To elucidate the mechanism by which macrophage EP4 deficiency promotes
   M1 polarization and foam cell formation, RNA-sequencing ([104]Figure
   S3) and proteomic profiling were performed on EP4^MKO and EP4^Flox
   BMDMs following 24 h of oxLDL stimulation. Compared to EP4^Flox BMDMs,
   EP4^MKO BMDMs exhibited significant alterations in 87 proteins,
   including 26 upregulated and 61 downregulated ([105]Figure 6A).
   Subsequent KEGG pathway enrichment analysis identified three pathways
   associated with inflammatory response and lipid metabolism ([106]Figure
   6B,C). Intersection analysis of the enriched proteins revealed CD36 as
   the sole candidate protein ([107]Figure 6D). The upregulation of CD36
   in EP4^MKO BMDMs upon oxLDL stimulation was further validated by
   Western blotting ([108]Figure 6E) and qRT-PCR ([109]Figure 6F).
   Consistently, immunofluorescence staining confirmed elevated CD36
   expression in aortic plaques from ApoE^−/^−EP4^MKO mice compared to
   ApoE^−/^−EP4^Flox mice ([110]Figure 6G), suggesting EP4 deficiency
   exacerbates atherosclerosis through CD36-mediated inflammatory and
   metabolic dysregulation.

Figure 6.

   [111]Figure 6
   [112]Open in a new tab

   EP4 upregulated the expression of CD36. (A) The histogram depicts the
   number of DEGs between BMDMs from EP4^Flox and EP4^MKO mice after
   treatment with oxLDL. (B) The KEGG pathway enrichment analysis of DEGs.
   (C) The gene sets enriched in KEGG pathways related to inflammation and
   lipid metabolism. (D) Schematic diagram illustrating the overlap of
   DEGs across KEGG inflammation and lipid metabolism-related signaling
   pathways. (E) Western blot analysis and quantification of CD36
   expression in BMDMs from EP4^Flox and EP4^MKO mice after treatment with
   oxLDL (50 μg/mL) for 24 h (n = 6). (F) qRT-PCR analysis of CD36 mRNA
   expression in oxLDL-stimulated BMDMs from the EP4^Flox and EP4^MKO mice
   (n = 3). (G) Double-immunofluorescence staining for F4/80 and CD36 in
   the aortic roots from ApoE^−/−EP4^Flox or ApoE^−/−EP4^MKO mice after 16
   weeks of Western diet (n = 5). Nuclei were stained with DAPI (blue);
   F4/80 stained as green; CD36 as red, scale bar: 20 μm. (H) Western blot
   analysis and quantification of CD36 expression in BMDMs from WT mice
   pre-treated with or without CAY10580 (100 nM), followed by stimulation
   with or without oxLDL (50 μg/mL) for 24 h (n = 6). Data presented as
   mean ± SEM, ns: not significant, * p < 0.05, Unpaired t-test in (E,G),
   two-way ANOVA in (G,H). DEGs: differentially expressed genes; BMDM:
   bone marrow-derived macrophage; oxLDL: oxidized low-density
   lipoprotein.

   To further explore the regulatory relationship between CD36 and EP4,
   BMDMs were pretreated with CAY10580, followed by stimulation with
   oxLDL. Western blotting analysis revealed that oxLDL significantly
   increased CD36 expression; such alteration was attenuated by treatment
   with CAY10580 ([113]Figure 6H), demonstrating EP4-dependent regulation
   of CD36.

3.7. EP4 Deficiency Enhanced Macrophage Foam Cell Formation and M1
Polarization at Least in Part by Upregulating CD36 Expression

   To confirm the role of CD36 in EP4-mediated macrophage polarization and
   foam cell formation, EP4^MKO BMDMs were transfected with CD36-siRNA or
   NC-siRNA, followed by oxLDL stimulation ([114]Figure S4). In EP4^MKO
   BMDMs, oxLDL-induced foam cell formation was significantly enhanced;
   however, such a change was attenuated by CD36 knockdown ([115]Figure
   7A). To further assess the interplay between EP4 and CD36 in macrophage
   polarization, CD36-siRNA-transfected BMDMs were subjected to M1- or
   M2-polarizing conditions. Upon LPS/IFN-γ stimulation, CD36 knockdown
   reduced mRNA levels of M1-associated pro-inflammatory factors compared
   to EP4^MKO BMDMs treated with NC-siRNA ([116]Figure 7B); conversely,
   IL-4/IL-13 stimulation of CD36 knockdown BMDMs resulted in increased
   expression of anti-inflammatory genes ([117]Figure 7C).

Figure 7.

   [118]Figure 7
   [119]Open in a new tab

   EP4 deficiency enhanced macrophage foam cell formation and M1
   polarization at least in part by upregulating CD36 expression. EP4^Flox
   and EP4^MKO BMDMs were pre-transfected with siRNA-CD36. (A) Oil Red O
   staining and quantification of foam cells in BMDMs after stimulation
   with or without oxLDL (50 μg/mL) for 24 h (n = 6), scale bar: 50 µm.
   (B,C) qRT-PCR was used to determine the mRNA levels of pro-inflammatory
   cytokines and macrophage polarization markers in BMDMs after
   stimulation with LPS/IFNγ or IL4/IL13 (n = 4). ApoE^−/−EP4^Flox and
   ApoE^−/−EP4^MKO mice were fed a Western diet for 12 weeks, followed by
   intraperitoneally injected with SSO for 4 weeks. The total aorta and
   aortic roots were collected. (D) Representative En face images and
   quantitation of Oil Red O-stained aorta from ApoE^−/−EP4^Flox and
   ApoE^−/−EP4^MKO mice (n = 4). Representative images and quantification
   of H&E staining ((E), n = 6, scale bar: 200 μm), Oil Red O staining
   ((F), n = 5, scale bar: 200 μm), Masson’s trichrome staining ((G), n =
   6, scale bar: 200 μm) in aortic root sections from ApoE^−/−EP4^Flox and
   ApoE^−/−EP4^MKO mice. Data presented as mean ± SEM, ns: not
   significant, * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001.
   One-way ANOVA in (B,C); two-way ANOVA in (A,D–G). BMDM: bone
   marrow-derived macrophage; SSO: Sulfosuccinimidyl oleate sodium.

   To assess the functional relevance of CD36 in EP4^MKO-induced
   atherosclerosis, ApoE^−/^−EP4^Flox and ApoE^−/^−EP4^MKO mice were fed a
   Western diet for 12 weeks in vivo, followed by their being
   intraperitoneally injected with Sulfosuccinimidyl oleate sodium (SSO,
   CD36 inhibitor) or saline for an additional 4 weeks. Compared to
   saline-treated controls, SSO administration significantly reduced
   plaque area in both the whole aorta ([120]Figure 7D) and the aortic
   root ([121]Figure 7E), decreased intraplaque lipid deposition
   ([122]Figure 7F), and increased collagen content ([123]Figure 7G).
   Collectively, these findings demonstrate that EP4 regulates macrophage
   polarization and foam cell formation through CD36 regulation, thereby
   contributing to the progression of atherosclerosis.

4. Discussion

   Atherosclerosis is a chronic vascular inflammatory disease
   characterized by thickening and hardening of the arterial wall due to
   lipid deposition and inflammatory response [[124]6,[125]20].
   Macrophages play an important role in the progression of AS,
   particularly through their roles in foam cell formation and phenotypic
   switching [[126]21,[127]22]. In the present study, we demonstrate that
   EP4 deficiency enhances macrophage foam cell formation and M1
   polarization at least in part by upregulating CD36 expression, thereby
   exacerbating the progression of atherosclerosis.

   EP4 is a PGE2 receptor and is involved in numerous pathophysiological
   processes [[128]23,[129]24]. While in vitro evidence indicates that EP4
   signaling suppresses PGE2-triggered inflammatory responses in
   macrophages [[130]25], the role of EP4 in vivo remains controversial.
   While EP4 deficiency has been reported to induce macrophage apoptosis
   and attenuate early development of atherosclerosis [[131]26], some
   studies reported that EP4 deficiency has no effect on plaque size or
   morphology during early stages of AS, but exacerbates local
   inflammation, and increases the plaque instability during advance
   stages [[132]27]. In type I diabetic mice, EP4 deficiency in myeloid
   cells did not significantly affect plaque formation [[133]28]. We
   herein demonstrate that macrophage-specific EP4 deficiency enhances
   vascular inflammation, accelerates atherosclerotic progression, and
   destabilizes plaques during advanced stages of AS. Notably, EP4 has
   been reported to be upregulated in human atherosclerotic plaques, where
   it exacerbates inflammation through upregulation of matrix
   metalloproteinase [[134]25]. In contrast, we found that EP4 expression
   was significantly downregulated in macrophages of the atherosclerotic
   plaques. Such discrepancy may be attributed to the differential
   expression of EP4 in different cell types in atherosclerotic plaques,
   or EP4 may exert stage-dependent dual roles.

   EP4-associated proteins have been found to interact with NF-KB to
   dampen macrophage activation, acting as a suppressor of inflammation
   [[135]29]. In the context of type II diabetes, EP4 has also been shown
   to influence islet inflammation by regulating macrophage polarization
   [[136]30]. Consistently, our study shows that EP4 deficiency skews
   macrophage polarization toward the M1 phenotype and enhances
   pro-inflammatory cytokine production. With respect to lipid metabolism,
   previous studies have shown that EP4-knockout mice spontaneously
   develop hypercholesterolemia by regulating the synthesis and
   elimination of bile acids [[137]16], suggesting the important role of
   EP4 in lipid homeostasis. In our study, macrophage-specific EP4
   deficiency promotes foam cell formation but does not alter plasma
   cholesterol levels, suggesting that the primary role of macrophage EP4
   lies in regulating lipid uptake rather than systemic lipid metabolism.

   CD36 is a membrane scavenger receptor that facilitates foam cell
   formation by mediating cholesterol uptake [[138]31]. In addition, CD36
   also promotes macrophage-driven chronic inflammation by mitochondrial
   metabolic pathways [[139]32]. During atherosclerosis, activated
   eosinophils can regulate macrophage switching from M2 to M1, which is
   partly mediated by the CD36 signaling pathway [[140]33]. Consistent
   with these studies, our findings reveal that CD36 contributes to both
   lipid accumulation and inflammatory signaling in macrophages, and these
   processes are regulated by EP4. Interestingly, CD36 has been reported
   to exert its function through complex formation with other membrane
   receptors, such as Na/K-ATPase-α1, which exerts a pro-inflammatory
   effect [[141]34]. In our study, we observed an increased expression of
   CD36 in EP4-deficient macrophages; however, the precise mechanism by
   which EP4 regulates CD36 expression remains to be elucidated.

   The study has several limitations. First, although we demonstrated that
   EP4 deficiency promotes macrophage M1 polarization and foam cell
   formation by upregulating CD36 expression, the precise mechanism by
   which EP4 regulates CD36 remains unclear and warrants further
   investigation. Second, since our study was primarily based on a murine
   model, further studies on clinical samples are needed to determine the
   significance of EP4-CD36 signaling axis in human atherosclerosis.
   Third, although the M1/M2 classification has been widely adopted to
   describe macrophage polarization states, it is increasingly recognized
   that this binary framework oversimplifies the spectrum of macrophage
   functional diversity in vivo. Macrophages often display mixed or
   transitional phenotypes rather than M1/M2. Future studies may further
   refine macrophage subpopulation characterization through
   high-dimensional profiling [[142]35,[143]36].

5. Conclusions

   In summary, our study demonstrates that EP4 attenuates the development
   of atherosclerosis by regulating CD36 expression, thereby suppressing
   foam cell formation and M1 macrophage polarization. Targeting the
   signaling components of the EP4-CD36 signaling axis may provide
   potential therapeutic targets for AS.

Supplementary Materials

   The following supporting information can be downloaded at:
   [144]https://www.mdpi.com/article/10.3390/cells14131021/s1, Table S1.
   The primer sequences used in genotyping. Table S2. The primer sequences
   used in qRT-PCR. Table S3. The sequences for siRNA. Figure S1. EP4
   expression is significantly decreased in EP4MKO mice. Figure S2. The
   EP4 level in HMDMs after siRNA transfection. Figure S3. RNA sequencing
   analysis of DEGs in BMDMs from EP4Flox and EP4MKO mice following oxLDL
   stimulation. Figure S4. The CD36 level in BMDMs after siRNA
   transfection.
   [145]cells-14-01021-s001.zip^ (766.6KB, zip)

Author Contributions

   Conceptualization, X.T., Y.Z. and W.T.; methodology, X.T., M.G., Y.W.,
   C.J., Y.B., Y.Z. and W.T.; software, X.T., M.G. and Y.W.; validation,
   Q.C., C.J., Y.B. and T.W.; formal analysis, X.T.; investigation, X.T.,
   Q.C., M.G., Y.W. and C.J.; resources, Y.Z. and W.T.; data curation,
   X.T.; writing—original draft preparation, X.T. and Y.Z.; writing—review
   and editing, X.T., Y.Z. and W.T.; visualization, X.T. and Y.W.;
   supervision, Y.Z. and W.T.; project administration, Y.Z. and W.T.;
   funding acquisition, Y.W., Y.Z. and W.T. All authors have read and
   agreed to the published version of the manuscript.

Institutional Review Board Statement

   Animal studies were reviewed and approved by the Animal Care and Use
   Committee of Guangzhou Women and Children’s Medical Center, Guangzhou
   Medical University, China (RSDW-2023-01397; approval date: 21 December
   2023). The study was conducted in accordance with the Declaration of
   Helsinki and approved by the Medical Ethics Committee of the Guangzhou
   Women and Children’s Medical Center ([2023]167B00; approval date: 7
   March 2023).

Informed Consent Statement

   Informed consent was obtained from all subjects involved in the study.

Data Availability Statement

   The original contributions presented in this study are included in the
   article/[146]Supplementary Material. Further inquiries can be directed
   to the corresponding author.

Conflicts of Interest

   The authors declare no conflicts of interest.

Funding Statement

   This work was supported by the National Science Foundation of Guangdong
   province (Grant No. 2024A1515030045, Y.Z.), the Guangzhou School
   (Institute) Enterprise Joint Funding Project-Outstanding talent project
   (Grant No. 2024A03J1089, W.H.T.), the Guangzhou Basic and applied basic
   research (Grant No. 2024A04J6579, W.H.T., Grant No. 202201011138,
   Y.W.), the National Natural Science Foundation of China (Grant No.
   32100956, Y.W.), plan on enhancing scientific research in GMU (Y.Z.,
   W.H.T.).

Footnotes

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References