Introduction

With the development of percutaneous coronary intervention (PCI), the mortality rates of ischemic heart disease (IHD) and myocardial infarction (MI) have significantly decreased. However, the consequent fibrosis and heart failure have created tremendous economic and social burdens for public healthcare [1, 2]. Therefore, exploring the pathophysiological mechanisms of acute inflammatory injury to chronic fibrotic remodeling after myocardial ischemia reperfusion (MIR) injury is of utmost importance for improving the prognosis of cardiovascular diseases. Acute myocardial ischemia and hypoxia can lead to the death of myocardial cells, the destruction of the structural integrity of the vascular endothelium, and the massive infiltration of inflammatory cells from the damaged endothelium to the injured site [3, 4]. With the progression of inflammation, necrotic cells and debris are cleared, and myofibroblasts proliferate, promoting tissue repair and fibrosis remodeling [5]. Various immune cells, such as macrophages, dendritic cells (DCs), T cells, and myofibroblasts, are involved in the transition from acute inflammation to chronic fibrosis. It is widely believed that myofibroblasts are the main effector cells, and macrophages also play a critical role in fibrosis onset. Therefore, exploring the signal exchange between macrophages and fibroblasts may create a novel target for alleviating the transition from acute MIR injury to fibrosis.

Granulocyte-macrophage colony-stimulating factor (GMCSF) is a monomeric glycoprotein primarily derived from endothelial cells, fibroblasts, and hematopoietic cells. GMCSF is mainly released during inflammatory stimulation and can effectively promote the maturation and activation of bone marrow-derived cells, such as monocytes, macrophages, and DCs. Currently, GMCSF is considered a key regulatory factor and target in the inflammatory response. In fibrotic transformation after kidney injury, GMCSF can effectively recruit macrophages to promote fibrosis progression [6]. In liver fibrosis, inhibiting GMCSF can effectively reduce the accumulation of CD206+ macrophages and the level of liver fibrosis [7]. In the cardiovascular system, anti-GMCSF can effectively reduce the level of MIR injury [8]. During acute MIR injury, GMCSF recruits a large number of immune cells to infiltrate the injured site, promoting the inflammatory response [8]. At this time, macrophages differentiated from migrated monocytes in the blood mainly present the M1 phenotype, releasing various inflammatory mediators, such as tumor necrosis factor-α (TNFα) and interleukin-1β (IL-1β), to promote the inflammatory response [9, 10]. Meanwhile, macrophages can release a large amount of chemokines such as c-c motif chemokine ligand 2 (CCL2), promoting the infiltration of immune cells in the blood to the injured site of the heart. CCL2 is a ligand for CC chemokine receptor 2 (CCR2) and can recruit a large number of CCR2+ immune cells to infiltrate damaged myocardial tissue [11]. CCL2 and CCR2 have long been considered important therapeutic targets for reducing MIR injury, and inhibiting CCL2/CCR2 can effectively reduce infarct size [12, 13]. Furthermore, it has been reported that CCL2/CCR2 plays an important role in the process of renal fibrosis [14, 15]. However, whether intervening with CCL2/CCR2 can alleviate the pathogenesis of myocardial fibrosis is currently unclear.

As the acute injury phase inflammatory response subsides, macrophages mainly transform into a reparative M2 phenotype, and the released transforming growth factor-β (Tgf-β) can promote the transformation of fibroblasts into myofibroblasts [16, 17]. Myofibroblasts can produce collagen and fibronectin, promoting extracellular matrix components, which are beneficial for the repair and remodeling of infarcted heart tissue, while also promoting scar formation and myocardial fibrosis progression [5, 18]. In addition to fibroblasts transforming into myofibroblasts, macrophages also have the potential to transform into myofibroblasts [19,20,21]. Therefore, exploring the mechanism of macrophages in the fibrosis process is crucial for fibrosis treatment. This study establishes an MIR model to observe the fibrotic transformation process after myocardial ischemic injury. Through in vivo and in vitro experiments, we explored the sources of GMCSF and the effects of GMCSF and CCL2 on macrophage phenotypic transformation. At the same time, mass cytometry was used to detect changes in immune cell function during fibrotic transformation when intervening with CCL2/CCR2 signaling, exploring the effects of GMCSF/CCL2/CCR2 signaling cascade intervention on immune cell homing/presence in fibrosis.

Materials and methods

Animal experiments

All animal experiments were conducted at Anhui Medical University and approved by the Anhui Medical University Ethics Committee (Approval No: LLSC20210347). Since estrogen has been proven to have an impact on MIR injury, this study uses male mice to exclude such effects [22]. In this study, we used 8–10-week-old wild-type (WT) and CCR2−/− male mice, all with a C57BL/6 background. WT mice were procured from the Animal Experiment Center of Anhui Medical University and CCR2−/− male mice were purchased from Shanghai Model Organisms. All the mice were housed in a 12-h light-dark cycle environment with ad libitum access to food and water and were acclimated for one week before the experiment began.

Mouse myocardial ischemia reperfusion model

A gas anesthesia machine (cat# R500IP, RWD Life Science, Shenzhen, China) and a small animal ventilator (cat# ALC-V8S, Alcbio, Shanghai, China) were used for continuous gas anesthesia and mechanical ventilation. Mice were fixed on the operating table, and the heart was exposed between the third and fourth ribs on the left side of the sternum. After 30 min of ligation of the left coronary artery, the ligature was untied for reperfusion. In addition to Sham group, the MIR groups were divided according to the reperfusion times of 1 day, 7 days, 14 days, and 30 days.

Drug treatment

RS102895 (cat# R1903, Sigma-Aldrich, MO, USA) is a specific CCR2 inhibitor, which is intraperitoneally injected into mice one day before MIR (5 mg/kg per 12 h) and continuously injected until the endpoint of the experiment [6]. GMCSF neutralizing antibody (NTAB, cat# MAB415, R&D, MN, USA) and isotype control (NTAB, cat# MAB006, R&D, MN, USA) were injected intravenously through the tail one day before MIR (300 μg/kg per 24 h) for two consecutive days [8].

In vivo, 24 h before MIR, mouse were intravenously injected with Clodronate liposomes (cat# CP-005-005, LIPOSOMA, Amsterdam, Netherlands) and repeated every 5 days to clear macrophages from the mouse.

Measurement of myocardial infarct size

After 2 h of reperfusion, the coronary artery was ligated in situ again, and 0.4% Evans Blue (cat# E2129, Sigma-Aldrich, MO, USA) was injected into the mouse’s internal jugular vein. One minute later, the heart was removed and washed with balanced saline solution to remove residual dye and blood from the mouse heart chamber. After fixing the morphology at low temperature, the heart was cut into approximately 2-mm thin slices and incubated with 2% 2,3,5-triphenyltetrazolium chloride (TTC) (cat# T8877, Sigma-Aldrich, MO, USA) at 37 °C for 10 min. An Epson scanner (Epson, Japan) was used to detect images.

Measurement of lactate dehydrogenase (LDH), creatine kinase myocardial band (CKMB), and cardiac troponin I (CTnI)

Mouse LDH assay kit (cat# A020-1-2, Jiancheng, Nanjing, China), mouse CK-MB isoenzyme assay kits (cat# H197-1-2, Jiancheng, Nanjing, China), and mouse cTn-I ELISA kit (cat# E-EL-M1203c, Elabscience, Wuhan, China) were used to test the levels of LDH, CKMB, and cTnI in mouse serum.

Histology, immunohistochemistry, and immunofluorescence of heart

Mouse heart specimens were collected at each time point, fixed in 10% formaldehyde, and embedded in paraffin. Sirius red staining, Masson’s trichrome staining, hematoxylin-eosin (H&E) staining, immunohistochemistry, and immunofluorescence were performed according to standard protocols [23]. Antigen retrieval was performed with citrate buffer at pH 6.0. Primary antibodies against CD68 (cat# 97778, CST, MA, USA), CCR2 (cat# ab273050, Abcam, Cambridge, UK), Ly6g (cat# ab238132, Abcam, Cambridge, UK), CD19 (cat# 90176, CST, MA, USA), and CD3e (cat# 85061, CST, MA, USA) were used to measure the levels of CD68+ cell, CCR2+ cells, neutrophils, B cells, and T cells in mouse heart tissue. Primary antibodies against Arg-1 (cat# 93668, CST, MA, USA) and iNos (cat# 13120, CST, MA, USA) were used to measure the polarization of macrophages in mouse heart tissue. Primary antibodies against GMCSF (cat# 17762-1-AP, Proteintech, Wuhan, China) and CCL2 (cat# 26161-1-AP, Proteintech, Wuhan, China) were used to measure expression levels. Co-staining of CD68, CCL2, and CCR2 was detected using primary antibodies against CD68 (cat# 97778, CST, MA, USA), CCL2 (cat# 26161-1-AP, Proteintech, Wuhan, China), and CCR2 (cat# ab273050, Abcam, Cambridge, UK). Levels of Tgf-β and Col1α1 were detected using primary antibodies against Tgf-β (cat# 21898-1-AP, Proteintech, Wuhan, China) and Col1α1 (cat# 72026, CST, MA, USA). After incubation with secondary goat antibody, a coverslip agent containing DAPI (cat# P0131, Beyotime, Nantong, China) was added. Observations were performed under a fluorescence microscope. The final area of the corresponding antibody-labeled positive cells was quantified using ImageJ (National Institutes of Health [NIH]).

Quantification of cytokines and chemokines

In this study, mouse heart tissue was collected at different time points following MIR, and tissue homogenates were prepared. The concentrations of cytokines and chemokines in the mouse heart homogenates were quantified using the Bio-Plex Pro Mouse Chemokine Panel 31-plex kit (cat# 12009159, Bio-Rad, CA, USA). The plate was read using the Luminex X200 system (cat# X200, Luminex Corporation, TX, USA).

Preparation of cardiac single-cell suspension

Fresh mouse heart tissue was collected, washed with PBS, and then cut into 1 mm3 pieces using surgical scissors. Mouse heart tissue was dissociated into single-cell suspensions using the Neonatal Heart Dissociation Kit (cat# 130-098-373, Miltenyi Biotec, Colonge, Germany). The single-cell suspension was then filtered through a 70 µM cell strainer, centrifuged at 400× g for 10 min, and red blood cells were lysed using Red Blood Cell Lysis Buffer (cat# R1010, Solarbio, Beijing, China). Cells were resuspended in PBS containing 10% Fetal Bovine serum (FBS, cat# FB15015, Clark Bioscience, WV, USA) and counted for dead and live cells.

Flow cytometry

Heart tissue was collected on 7th days and 14th day post reperfusion and prepared as single-cell suspensions. After cell counting, single-cell suspensions were incubated with antibodies (Biolegend, CA, USA) at 4 °C for 30 min for cell surface staining. Single-cell suspensions were washed twice with PBS and then resuspended in 100 µL PBS. DCs were defined as CD45+CD11c+ cells, and macrophages were defined as CD45+CD11b+/CD45+F4/80+ cells. Samples were collected using a flow cytometer (BECKMAN, Germany) and analyzed using Flow JO version 10.

Non-cardiomyocyte isolation and magnetic cell sorting

On 1st day after MIR, single-cell suspensions were isolated from mouse hearts and cardiomyocytes were removed by Percoll (cat# 65455-52-9, Solarbio, Beijing, China) density-gradient centrifugation [24]. Dead cells were removed using the MojoSort™ Mouse Dead Cell Removal Kit (cat# 480157, Biolegend, CA, USA), followed by cell counting. The CD45 positive sorting kit (cat# 480027, Biolegend, CA, USA) was used to completely separate CD45+ cells in non-cardiomyocytes, while cells not bound by magnetic beads were considered non-immune cells of the heart tissue. CD11b (cat# 480109, Biolegend, CA, USA) and CD11c (cat# 480077, Biolegend, CA, USA) positive sorting kits were used to further classify CD45+ cells. DCs were defined as CD45+CD11c+ cells, macrophages were defined as CD45+CD11b+ cells, and the remaining CD45+CD11b-CD11c− cells were primarily granulocytes. All those cells were cultured 24 h without serum to detect the level of GMCSF.

Mass cytometry

Mass cytometry was performed by Zhejiang Puluoting Health Technology Co., Ltd (Hangzhou, China). Cells were stained on ice for 5 min with 0.25 μM Cell-ID™ Cisplatin-194Pt (cat# 201194, Fluidigm, CA, USA) to distinguish live and dead cells in single cell suspensions. Cells were then resuspended in FACS buffer to quench cisplatin staining. Subsequently, cells were blocked with blocking solution on ice for 20 min to reduce non-specific binding. Surface antibody mix (Fluidigm, CA, USA) was added and incubated on ice for 30 min. After washing three times with FACS buffer, cells were incubated at room temperature for 1 h with Maxpar® Fix and Perm Buffer (cat# 201067, Fluidigm, CA, USA). Cells were then resuspended in 100 μL of intracellular antibody cocktail (Fluidigm, CA, USA) and incubated on ice for 30 min. After washing, cells were resuspended in EQ™ Four Element Calibration Beads (cat# 201078, Fluidigm, CA, USA) and data were acquired on Mass cytometry (Fluidigm, CA, USA). CyTOF data were analyzed using R (version 4.3.1, Austria). In this study, non-cardiomyocyte cells from nine mice’s hearts per group were pooled to reduce single sample error.

Primary cardiac fibroblasts (PCFs) extraction

Newborn C57BL/6 mouse aged 1–3 days were collected, soaked in 75% alcohol for 30 s, and their hearts were minced. The tissue was digested with 0.25% trypsin (cat# C0201, Beyotime, Nantong, China) in a 37 °C water bath until the digestion solution became turbid, and the first digestion solution was discarded. The digestion was continued with 0.25% trypsin, and after the supernatant became turbid, DMEM medium (cat# 11965118, Gibco, NY, USA) containing 10% FBS (cat# FB15015, Clark Bioscience, WV, USA) was added to terminate the digestion. The above digestion steps were repeated until the heart tissue was completely digested, and then centrifuged at 1500 rpm for 5 min. Cells were resuspended in DMEM medium containing 10% FBS and cultured in gelatin-coated dishes in a 37 °C incubator for 1 h. The medium was discarded and replaced with fresh DMEM medium containing 10% FBS. PCFs from passages 2–4 were used for subsequent experiments.

Bone marrow-derived macrophages (BMDMs) extraction and culture

Tibiae and femurs were collected from 6-8 week-old mouse to extract BMDMs. The cells were resuspended in 15% L929 cell-conditioned medium and 85% FBS-containing 1640 medium (cat# 11875093, Gibco, NY, USA) and cultured in a 37 °C, 5% CO2 incubator for 7 days. In 6-well plates used for BMDMs culture, recombinant mouse GMCSF (cat# 415-ML, R&D System, MN, USA), and recombinant mouse CCL2 (cat# 250-10, Peprotech, NJ, USA) were added to stimulate BMDMs for 24 h to detect the polarization status of BMDMs and Tgf-β release levels. Mouse cardiac microvascular endothelial cells (MCMECs) were bought from Shanghai Qincheng Biotechnology Co., Ltd. (Shanghai, China). In order to investigate the effects of MCMECs on macrophages, MCMECs were seeded in the upper chamber of 0.4-μm polyester membrane Transwell inserts (cat# 3450, Corning, NY, USA), and BMDMs were seeded in a six-well plate. The upper chamber containing MCMECs were then inserted into the six-well plate containing BMDMs, and co-cultured for 24 h with the medium changed according to different groups containing either 10% serum or serum-free medium. In the serum-free co-culture system of MCMECs and BMDMs, recombinant mouse GMCSF neutralizing antibody (NTAB, cat# MAB415, R&D, MN, USA) or IgG2A isotype control (cat# MAB006, R&D, MN, USA) was added and treated for 24 h. To investigate the effect of BMDMs on PCFs, BMDMs were seeded in the 0.4-μm polyester membrane Transwell inserts suitable for six-well plates, and the Transwell chamber containing BMDMs was inserted into the six-well plate for co-culturing with PCFs for 24 h.

Migration assay

The supernatant of serum-free 1640 medium and 10% FBS-containing 1640 medium cultured with MCMECs for 24 h were collected. 10% FBS was added to the serum-free conditioned medium to exclude the influence of serum on the migration assay. BMDMs were seeded in the 8-μm polyester membrane Transwell inserts (cat# 14342, Labselect, Hefei, China), and the lower chamber was filled with the collected MCMEC medium or 1640 medium containing recombinant mouse GMCSF and/or CCL2 to be co-cultured with BMDMs for 24 h. After removing the upper chamber, cells were fixed with 4% paraformaldehyde (cat# BL539A, Biosharp, Hefei, China) at room temperature for 30 min, washed three times with PBS, stained with 0.1% crystal violet solution (cat# G1064, Solarbio, Beijing, China) for 15 min, washed three times with PBS, and photographed under a microscope.

Concentration of GMCSF, Tgf-β, CCL2 in medium

The cell supernatant was collected, and the levels of GMCSF, Tgf-β, and CCL2 were measured according to the instructions of GMCSF (cat# E-EL-M0032c, Elabscience, Wuhan, China), Tgf-β (cat# EMC107b.96, NeoBioscience Technology Co, Ltd, Shenzhen, China), and CCL2 (cat# EMC113.96, NeoBioscience Technology Co, Ltd, Shenzhen, China) assay kits.

Immunocytochemistry staining

BMDMs, MCMECs, or PCFs were cultured in 12-well plates containing glass slides and treated for 24 h. Cells were then fixed with 4% paraformaldehyde at room temperature for 30 min, permeabilized with 0.1% Triton X-100 (cat# P0096, Beyotime, Nantong, China) at 4 °C for 20 min, and washed with PBS. Non-specific fluorescence was blocked using immunofluorescence-specific blocking solution (cat# P0260, Beyotime, Nantong, China). Primary antibodies against F4/80 (cat# MAB5580, R&D System, MN, USA), CCL2 (cat# 66272-1-lg, Proteintech, Wuhan, USA), Tgf-β (cat# 21898-1-AP, Proteintech, Wuhan, China), iNos (cat# 13120, CST, MA, USA), GMCSF (cat# 17762-1-AP, Proteintech, Wuhan, China), and Col1α1 (cat# 72026, CST, MA, USA) were used for overnight staining at 4 °C. After washing with PBS, cells were co-incubated with fluorophore-conjugated secondary antibodies (Abcam, Cambridge, UK). Finally, cells were mounted with DAPI-containing mounting medium (cat# P0131, Beyotime, Nantong, China) and images were acquired using a fluorescence microscope (cat# DMI8, Leica, Wetzlar, Germany).

Western blot

The samples were subjected to 10% and 12.5% SDS PAGE, and Western blot analysis was performed as standard protocol. The PVDF membranes (cat# IPVH00010, Millipore, MA, USA) were incubated with primary antibodies against GMCSF (cat# 17762-1-AP, Proteintech, Wuhan, China), Arg-1 (cat# 93668, CST, MA, USA), Tgf-β (cat# 21898-1-AP, Proteintech, Wuhan, China), CCL2 (cat# 66272-1-lg, Proteintech, Wuhan, USA), NLRP3 (cat# ab263899, Abcam, Cambridge, UK), Caspase-1 (cat# 22915-1-AP, Proteintech, Wuhan, China), IL-1β (cat# 16806-1-AP, Proteintech, Wuhan, China), β-actin (cat# 66009-1-Ig, Proteintech, Wuhan, China), and GAPDH (cat# 10494-1-AP, 1:5000, Proteintech, Wuhan, China) followed by horseradish peroxidase-conjugated secondary antibody incubation. Bands were visualized by an electro-chemiluminescence (ECL) system (Bio-Rad, USA) and quantified using ImageJ software.

Statistical analysis

Data with normal distribution were presented as mean ± standard deviation (SD), while non-normal distribution data were presented as median and quartiles. Comparisons of numerical parameters between normally distributed data were conducted using one-way ANOVA, with Bonferroni post-hoc test for subgroup comparisons. Comparisons of numerical parameters between two groups were analyzed using t-test. All statistical analyses and graph production were performed using SPSS 16.0 and GraphPad Prism 9.0 (GraphPad Software, USA), respectively.

Results

Chronic fibrosis remodeling after acute MIR injury is related to macrophages

H&E staining confirmed myocardial fiber disruption and significant inflammatory cell infiltration after MIR. As the reperfusion time increased, the level of inflammatory cell infiltration into cardiac tissue gradually rose, peaking at 24 h post reperfusion (Fig. 1a). With increasing reperfusion time, Masson and Sirius red staining showed that there was early myocardial fibrosis on the 7th day after MIR, with a large amount of collagen fiber infiltration in the injured tissue interstitial space and complete formation of fibrosis on the 14th day (Fig. 1b, c). Meanwhile, immunohistochemistry of CD68 indicated that macrophages had the highest level on the 1st day after MIR, and the early infiltration of macrophages into myocardial tissue was characterized by a small number of M2 macrophages with high expression of Arg-1, with the majority exhibiting a proinflammatory M1 phenotype with high expression of iNos (Fig. 1d–i). Interestingly, the number of M1 macrophages was significantly reduced on the 7th day, replaced by the reparative M2 anti-inflammatory phenotype, suggesting that the 7th day is a critical time point for fibrosis repair (Fig. 1e, f, h, i).

Fig. 1: Macrophages infiltration during the transition from acute MIR injury to fibrosis.
figure 1

a Representative images of heart HE staining at each time point before and after MIR. Scale bars = 50 µm, n = 5. b Masson staining detect the fibrosis level of each group. Scale bars = 500 µm, n = 5. c Sirius red staining detects the fibrosis level of each group. Scale bars = 50 µm, n = 5. d–f Hearts were harvested in each group and heart sections immunostained with anti-CD68, iNos, and Arg-1. Scale bars = 50 µm, n = 5. g–i Quantification of CD68, iNos, and Arg-1-positive area as performed in (d–f), n = 5. ns no significance; **P < 0.01 compared to Sham group, one-way ANOVA.

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GMCSF/CCL2/CCR2 signal activation during the transition to fibrosis after ischemic myocardial injury

Luminex chemokine array analysis was used to analyze the changes in chemokines in the heart that underwent acute MIR injury. Most of these chemokines peaked on the 1st day after MIR and dropped to the level of the Sham group on the 7th day after MIR (Fig. 2a). Among them, the changes in GMCSF and CCL2 were the most significant. Luminex chemokine array analysis showed that the levels of GMCSF on the 1st day after MIR were approximately 5 times higher than those in the sham group, and CCL2 levels were 16 times higher than those in the Sham group (Fig. 2a). On the 7th day after MIR, the levels of GMCSF and CCL2 further decreased and subsequently decreased to levels similar to those of the Sham group (Fig. 2a). Immunohistochemistry of GMCSF and CCL2 also confirmed this result, and the aggregation level of CCR2+ cells was consistent with the change in GMCSF and CCL2 levels on the 1st day after reperfusion (Fig. 2b–g). Although CCR2+ cells on the 7th day were lower than those on the 1st day after MIR, they were still significantly higher than those in the Sham group until fibrosis was completely formed on the 14th day after MIR, which then decreased to the level of the Sham group.

Fig. 2: Cytokine levels in mouse heart tissue at different time points after MIR.
figure 2

a Quantification of heart chemokine concentrations at different time points after MIR injury and Sham group, n = 3. ns no significance; *P < 0.05; **P < 0.01 compared to D1, one-way ANOVA. b–g Hearts were harvested at each group and heart sections immunostained with anti-GMCSF, CCL2, and CCR2, and Quantification of GMCSF, CCL2, and CCR2-positive area. Scale bars = 50 µm, n = 5. ns no significance; *P < 0.05; **P < 0.01 compared to Sham, one-way ANOVA. h the concentration of GMCSF and CCL2 in heart at different time points within 24 h after MIR injury and Sham group, n = 5. ns no significance; *P < 0.05; **P < 0.01 compared to Sham, one-way ANOVA. i On the 1st day post MIR, heart tissue was digested into a single-cell suspension and subjected to cell sorting to explore the source of GMCSF. n = 5. j Representative immunofluorescence images of anti-CD68, CCL2, CCR2 co-staining of sham or 2, 6, 12, 24 h post MIR in damaged heart tissues. Yellow arrows indicate the co-localization of CCR2+CD68+ cells and CCL2 in the heart. Scale bars = 50 µm, n = 5.

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We further examined the trends in GMCSF and CCL2 changes within 24 h post MIR. After reperfusion, GMCSF began to increase at 2 h, reaching a peak at 6 h and gradually decreasing thereafter (Fig. 2h). CCL2 started to increase at 2 h, peaking at 24 h (Fig. 2h). This further confirmed GMCSF as the primary chemoattractant factor driving macrophage infiltration. To explore the source of GMCSF, we performed cell sorting on the hearts of mice on the 1st day after MIR and then detected the expression levels of GMCSF in different cell populations. The results showed that higher levels of GMCSF were detected in nonimmune cells (Fig. 2i).

CCR2 is the receptor for CCL2, and CCR2+ macrophages have long been considered infiltrating macrophages. Four-color immunofluorescence confirmed that CCR2+ macrophages infiltrated the heart as early as 2 h post MIR, peaking at 24 h, indicating that CCR2+ macrophages respond early to inflammation (Fig. 2j, Supplementary Fig. S1a).

GMCSF derived from MCMECs promotes the release of CCL2 by BMDMs

We extracted PCFs and MCMECs from neonatal mice in vitro and simulated ischemic injury in vivo by starvation treatment. Compared with MCMECs cultured in 10% serum-containing medium, MCMECs cultured in serum-free medium showed higher levels of GMCSF fluorescence (Fig. 3a, b). PCFs seemed to have no ability to release GMCSF before transitioning to myofibroblasts (Supplementary Fig. S1b, c). Western blot and ELISA detection showed that GMCSF expression in serum-free cultured MCMECs was significantly increased (Fig. 3c–e). Next, we extracted mouse-derived BMDMs, and immunofluorescence of F4/80 was used to detect macrophage purity (Supplementary Fig. S1d). Compared with the M0 group, BMDMs treated with recombinant mouse GMCSF expressed higher levels of CCL2 (Fig. 3f–h, Supplementary Fig. S1e). Starvation-treated macrophages showed no significant changes, while BMDMs co-cultured with MCMECs released higher levels of CCL2 than M0 macrophages (Fig. 3f–h, Supplementary Fig. S1e). Interestingly, the effect of starvation-induced MCMECs on promoting BMDM release of CCL2 could be reversed by exogenous GMCSF NTAB, while the isotype control of GMCSF NTAB had no significant effect on MCMECs-induced BMDMs release of CCL2 (Fig. 3f–h, Supplementary Fig. S1e). Crystal violet staining also confirmed that GMCSF in the lower chamber significantly promoted the migration of macrophages to the lower chamber, and CCL2 combined with GMCSF further enhanced the migration induced by GMCSF in the co-culture system (Fig. 3i, j). The results also showed that GMCSF released by MCMECs can effectively promote the migration of BMDMs, and this effect can be reversed by exogenous GMCSF NTAB (Fig. 3i, j). Next, we examined the expression levels of CCL2 and CCR2 in mice treated with GMCSF NTAB and isotype controls on the 1st day after MIR. In vivo experiments showed that GMCSF NTAB could also inhibit the expression levels of CCL2 and CCR2 (Fig. 3k–m).

Fig. 3: MCMECs releases GMCSF under starvation treatment, inducing macrophage migration and releasing CCL2.
figure 3

a Detection of GMCSF expression level in MCMECs by immunofluorescence and b Quantification of GMCSF-positive area as performed in (a). Scale bars = 50 µm, n = 5. ns no significance; **P < 0.01 compared to control, Student’s t-test. c Western blot of GMCSF in MCMECs and d quantified and normalized to GAPDH. n = 3. ns no significance; **P < 0.01 compared to control, Student’s t-test. e Released GMCSF levels in MCMEC culture supernatants, n = 8. ns no significance; **P < 0.01 compared to control, Student’s t-test. f The CCL2 expression level in BMDMs after treatment for 24 h with exogenous recombinant mouse GMCSF, co-cultured with MCMECs with or without GMCSF NTAB. Scale bars = 50 µm, n = 5. g Quantification of CCL2-positive area as performed in (f). n = 5. ns no significance; **P < 0.01, one-way ANOVA. h Released CCL2 levels in BMDMs culture supernatants, n = 8. ns no significance; **P < 0.01, one-way ANOVA. i, j The numbers of Transwell migrated adherent BMDMs after 24 h of GM‐CSF (50 ng/mL) and/or CCL2 (50 ng/mL) treatments, cultured with MCMEC medium with/without GMCSF NTAB. Scale bars = 100 µm, n = 5. ns no significance; *P < 0.05; **P < 0.01, one-way ANOVA. k–m The expression levels of CCL2 and CCR2 in mouse treated with GMCSF NTAB and isotype controls after MIR were measured. Scale bars = 50 µm, n = 5. ns no significance; *P < 0.05 compared to MIR-D1, one-way ANOVA.

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Knocking out CCR2-gene or pharmacologically inhibiting CCR2 effectively alleviated MIR injury

Compared with the Sham group, the infarct area of WT mice after MIR was significantly increased, and it was accompanied by a significant increase in myocardial injury-related biomarker levels (Fig. 4a–d). We generated CCR2−/− mice and subjected them to the same MIR treatment. Compared with WT mice on the 1st day after MIR, CCR2−/− mice showed a significant reduction in MI size and myocardial injury biomarkers in serum (Fig. 4a–d). Administration of the CCR2 inhibitor also significantly reduced the MI area and myocardial injury levels, while no significance was observed when the CCR2 inhibitor vehicle was applied to the MIR group (Fig. 4a–d). We further detected changes in myocardial inflammation in mice and found that compared with those in WT mice undergoing MIR, GMCSF and CCL2 were significantly reduced in the heart tissue of CCR2−/− mice on the 1st day after MIR (Fig. 4e). At the same time, the levels of inflammatory factors released by macrophages, such as TNF-α, IL-1β, IL-4, IL-6, and IL-10, were consistent with the changes in GMCSF and CCL2 (Fig. 4e). Western blot analysis further confirmed that knocking out CCR2-gene could reduce GMCSF and CCL2 levels (Fig. 4f, g). After MIR, the expression of NLRP3 inflammasome-related signals significantly increased, leading to inflammatory damage (Fig. 4f, g). Knocking out CCR2-gene effectively reduced acute inflammatory damage caused by MIR (Fig. 4f, g). CCR2 inhibitors also effectively reduced inflammatory damage mediated by NLRP3/Caspase-1/IL-1β signaling (Fig. 4h, i).

Fig. 4: GMCSF/CCL2/CCR2 promotes the transition from MIR injury to cardiac fibrosis through the NLRP3 signaling pathway.
figure 4

a, b Evans blue and TTC staining detect the infarct size and area at risk after MIR in WT mice treated with CCR2 inhibitor or not and CCR2−/− mice, n = 5. ns no significance; *P < 0.05; **P < 0.01 compared to WT-MIR, one-way ANOVA. c, d The serum level of cTnI, CKMB, and LDH was detected at 1st day after MIR in WT mice treated with CCR2 inhibitor (RS102895) or not and CCR2−/− mice, n = 8. ns no significance; *P < 0.05; **P < 0.01 compared to WT-MIR, one-way ANOVA. e Inflammatory factor levels in mouse after MIR, n = 3. ns no significance; *P < 0.05; **P < 0.01, Student’s t-test. f, g Western blots of GMCSF, CCL2, NLRP3, Caspase-1, and IL-1β and the quantification of expression on 1st day after MIR and sham group. n = 5. ns no significance; **P < 0.01 compared to WT-MIR, one-way ANOVA. h, i Western blots of NLRP3, Caspase-1, and IL-1β and the quantification of expression on 1st day after MIR. n = 5. ns no significance; *P < 0.05, **P < 0.01 compared to WT-MIR group, one-way ANOVA.

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CCL2 induces macrophage M2 phenotype transition, later promoting the transformation of cardiac fibroblasts into myofibroblasts

After treatment with GMCSF, BMDMs showed a significant increase in iNos expression, and treatment with CCL2 resulted in a significant increase in Arg-1 levels (Fig. 5a–d, Supplementary Fig. S2a). Treating macrophages with CCL2 promoted Arg-1 expression and reversed the GMCSF-induced increase in iNos (Fig. 5a–d, Supplementary Fig. S2a). After the 1st day of MIR, Tgf-β significantly increased, peaking on the 7th day and gradually decreasing thereafter (Fig. 5e). Tgf-β has long been considered the primary driver of fibrosis. To explore whether macrophages are the main source of Tgf-β, we examined Tgf-β expression levels in mice after macrophage depletion (MD) and MIR (Fig. 5f, g). Compared to mice that underwent MIR, mice that underwent MD had significantly lower levels of Tgf-β expression (Fig. 5f, g). Since M2 macrophages can promote fibrotic repair by Tgf-β, we detected the level of Tgf-β. The fluorescence of Tgf-β suggested that CCL2 could reverse M1 macrophages induced by GMCSF to M2 macrophages and promote the release of Tgf-β (Fig. 5h, i, Supplementary Fig. S2b). However, BMDMs from CCR2−/− mice showed a significant decrease in Tgf-β release under combined stimulation with GMCSF and CCL2 (Fig. 5h, i, Supplementary Fig. S2b). Next, we co-cultured PCFs with BMDMs for 24 h. Compared with normal M0 medium and GMCSF and CCL2-containing medium, M0 medium cultured BMDMs co-cultured with cardiac fibroblasts for 24 h promoted the release of Col1α1 by cardiac fibroblasts, and the BMDMs treated with GMCSF and CCL2 combination co-cultured with cardiac fibroblasts further aggravated the fibrotic transformation of cardiac fibroblasts, promoting the release of Col1α1 by myofibroblasts (Fig. 5j–l, Supplementary Fig. S2c). After combined GMCSF and CCL2 treatment for 24 h, the ability of CCR2−/− mice BMDMs to promote the release of Col1α1 by myofibroblasts was significantly reduced compared with that of WT mice-derived BMDMs (Fig. 5j–l, Supplementary Fig. S2c).

Fig. 5: CCL2 reverses M1 macrophage-induced by GMCSF to the M2 macrophage, which releases Tgf-β to promote the transformation of fibroblasts into myofibroblasts.
figure 5

a The expression level of iNos in BMDMs after 24 h of GMCSF (50 ng/mL) and/or CCL2 (50 ng/mL) treatments. Scale bars, 50 µm, n = 5. b Quantification of iNos-positive area as performed in Fig. 5a, n = 5. ns no significance; **P < 0.01, one-way ANOVA. c Western blot of Arg-1 in BMDMs and d quantified and normalized to GAPDH, n = 5. ns no significance; **P < 0.01, one-way ANOVA. e The concentration of Tgf-β in heart at different time points after MIR injury and Sham group, n = 5. ns no significance; **P < 0.01 compared to Sham, one-way ANOVA. f Mouse experiencing macrophage depletion showed changes in Tgf-β post MIR and g quantified and normalized to GAPDH, n = 5. ns no significance; *P < 0.05, **P < 0.01 compared to WT-MIR, one-way ANOVA. h The expression level of Tgf-β in BMDMs after 24 h of recombinant mouse GMCSF (50 ng/mL) and/or CCL2 (50 ng/mL) treatments. Scale bars = 50 µm, n = 5. i Quantification of Tgf-β-positive area as performed in (h), n = 5. ns no significance; *P < 0.05, **P < 0.01, one-way ANOVA. j The expression level of Col1α1 in PCFs co-cultured with BMDMs. Scale bars = 50 µm, n = 5. k, l Quantification of myofibroblast size and Col1α1-positive area as performed in (j), n = 5. ns no significance; **P < 0.01, one-way ANOVA.

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Interference with CCR2 alleviates acute MIR injury and chronic fibrotic transformation

We also examined the expression levels of Tgf-β and Col1α1 and found that heart tissue on the 7th day post MIR had high expression of Tgf-β and Col1α1, and inhibiting CCR2 effectively reduced Tgf-β and Col1α1 levels (Fig. 6a–c, Supplementary Fig. S2d). Consistent with the changes in Tgf-β and Col1α1, there were fibrotic changes. On the 7th day after MIR in WT mice, the fibrosis area was significantly increased, while the fibrosis size in CCR2−/− mice was significantly reduced, and the CCR2 inhibitor also significantly reduced the level of myocardial fibrosis (Fig. 6d, e). On the 14th day after MIR in WT mice, the fibrosis trend was consistent with that on the 7th day (Fig. 6f, g). We also detected the cardiac function of mice on the 30th day after MIR, and the normal mouse left ventricular ejection fraction (LVEF) was as high as 90%, but the LVEF of mice after MIR decreased by 40% (Fig. 6h–k). Knockout of the CCR2 gene or use of a CCR2 inhibitor significantly increased the LVEF after MIR, protecting cardiac function. The changes in left ventricular fractional shortening (FS) were similar to those of LVEF, and the changes in left ventricular end-systolic volume/left ventricular end-diastolic volume (LVESV/LVEDV) were opposite to those of LVEF (Fig. 6h–k). These indicators all represent the quantification of cardiac function, confirming that CCR2 can effectively reverse the decline in cardiac function in mice after MIR. In addition to the transthoracic echocardiogram, the cardiac index at 30th days after MIR can also intuitively reflect the level of compensatory hypertrophy of the heart (Fig. 6l). Our results suggest that the MIR group and the vehicle control group had the highest cardiac index, while the CCR2 inhibitor significantly reduced the cardiac index after MIR (Fig. 6l). The cardiac index of CCR2−/− mice was lower than that of the other three groups of mice that underwent MIR. The above results confirm that inhibiting CCR2+ immune cells can alleviate acute myocardial injury as well as long-term myocardial fibrosis and cardiac remodeling.

Fig. 6: The effect of inhibiting CCR2 on fibrosis and cardiac function after MIR injury.
figure 6

a Heart sections were immunostained with anti-Col1α1 (green), anti-Tgf-β (red), and DAPI (identifies nuclei, blue). Scale bars = 50 µm, n = 5. b, c Quantification of Col1α1 and Tgf-β-positive area as performed in (a), n = 5. ns no significance; **P < 0.01, one-way ANOVA. d, e Masson staining of heart at 7th days after MIR and Quantification of fibrosis size. Scale bars = 500 µm, n = 5. ns no significance; **P < 0.01, one-way ANOVA. f, g Masson staining of heart at 14th day after MIR and Quantification of fibrosis size. Scale bars = 500 µm, n = 5. ns no significance; **P < 0.01, one-way ANOVA. h Representative M-mode echocardiographic images at 30th day after MIR. i–k LVESV/LVED, LVEF, and FS as performed in (h), n = 5. ns no significance; *P < 0.05; **P < 0.01 compared to MIR-D30, one-way ANOVA. l Heart weight index was presented as heart weight/body weight at 30th day after MIR, n = 5. ns no significance; **P < 0.01 compared to MIR-D30, one-way ANOVA.

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Mass cytometry explores cellular microenvironment changes in acute MIR injury to fibrosis transition

We used mass cytometry to detect changes in the cardiac microenvironment of Sham mice, WT mice and CCR2−/− mice on the 7th day after MIR. Based on the marker signatures of various mouse cell types, we classified them into 33 clusters and eight cell types based on their lineage (Fig. 7a). Cell-specific markers were used to construct the gating strategy for the identification of cardiac immune cell types in Cytof (Supplementary Fig. S3). Compared to the Sham group, the level of CD45+ cells in the heart on the 7th day after MIR was significantly increased, mainly manifested as an increase in macrophages and DCs (Fig. 7b). The TSNE map suggests that the increased macrophages at 7th days after MIR highly express markers such as CD64, F4/80, CD11b, CX3CR1, and MHCII (Fig. 7c). Among them, macrophages highly expressed CD206 on the 7th day and expressed lower levels of iNos (Fig. 7d). Flow cytometry detected changes in macrophages, confirming that the levels of macrophages were elevated on the 7th day after MIR and reduced in CCR2−/− mice (Fig. 7e, g). In CCR2−/− mice that underwent MIR, the level of CD206+ macrophages were significantly reduced, while iNos did not show a significant difference compared to the MIR-D7 group (Fig. 7d). We further detected the levels of Arg-1 and iNos by immunohistochemistry and found that Arg-1+ macrophages were highly expressed on the 7th day after MIR, and Arg-1+ macrophages were significantly reduced in CCR2−/− mice that underwent MIR (Fig. 7f, h). There was no significant difference in iNos between the sham group and the MIR-D7 group (Fig. 7i, j).

Fig. 7: Mass cytometry detection of the cardiac immune microenvironment status of mouse on the seventh day after MIR.
figure 7

a Representative TSNE plot showing unsupervised clustering of cardiac immune cells. b Quantification of each cell in Sham and MIR conditions, n = 9. Each dot represents a single cell in the TSNE plot. c, d Representative TSNE plots from CyTOF data showing colored expression in arbitrary units (AU) of CD64, F4/80, CD11b, CX3CR1, MHC-II, CD206, and iNos in cardiac immune cells in Sham and MIR conditions, n = 9. e Representative flow cytometry plots and g quantification of cardiac macrophages in Sham and 7th day after MIR in mice. n = 5. ns no significance; **P < 0.01 compared to MIR-D7, one-way ANOVA. f Hearts were harvested at Sham and MIR group and heart sections immunostained with anti-Arg-1 and h quantification of Arg-1-positive area as performed in (f). Scale bars = 50 µm, n = 5. ns no significance; **P < 0.01 compared to MIR-D7, one-way ANOVA. i Hearts were harvested at Sham and MIR condition and heart sections immunostained with anti-Arg-1. Scale bars = 50 µm, n = 5. j Quantification of Arg-1-positive area as performed in (i), n = 5. ns no significance; *P < 0.05; **P < 0.01 compared to MIR-D7, one-way ANOVA.

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The TSNE map suggests that the level of DCs was elevated on the 7th day after MIR and declined in CCR2−/− mice (Fig. 8a). Flow cytometry detected changes in DCs, verifying that CCR2−/− mice underwent MIR recruitment of lower levels of DCs than WT mice that underwent the same MIR treatment (Fig. 8b, f). However, there were no significant changes in the levels of neutrophils, T cells, and B cells in the MIR-D7 group and CCR2−/−-MIR-D7 group compared to the Sham group (Fig. 8a). Immunohistochemistry confirmed that knockout of the CCR2-gene had no significant effect on these cells in the cardiac microenvironment, and no particular difference was observed with the administration of CCR2 inhibitors (Fig. 8c–e, g–i). To further explore the roles of DCs and macrophages during fibrosis formation and termination, we further detected the levels of macrophages and DCs in the heart tissue of mice on the 14th day after MIR surgery and found that although the overall levels were lower than those on the 7th day, the levels of DCs and macrophages in CCR2−/− mice were lower than those in WT mice. CCR2 inhibitors also significantly inhibited macrophage and DC infiltration (Supplementary Fig. S4a–e). The above results confirm that T cells mainly participate in the acute injury stage of MIR injury and do not participate in the later fibrosis pathological process. In the chronic fibrosis remodeling period, macrophages mainly release Tgf-β to promote fibrosis progression.

Fig. 8: Mass cytometry detection of the cardiac immune microenvironment status of mouse on the 7th day after MIR.
figure 8

a Representative TSNE plots from CyTOF data showing colored expression in arbitrary units (AU) of CD11c, Ly6g, CD3e, and CD19 in cardiac immune cells in Sham and MIR conditions, n = 9. b Representative flow cytometry plots and f quantification of cardiac dendritic cells in Sham and 7th days after MIR in mouse. n = 5. c–e Hearts were harvested at sham and MIR group and heart sections immunostained with anti-Ly6g, CD3e, and CD19. Scale bars = 50 µm, n = 5. g–i Quantification of Ly6g, CD3e, and CD19-positive area as performed in (c–e), n = 5. ns no significance; **P < 0.01 compared to MIR-D7, one-way ANOVA.

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Discussion

Macrophages have been demonstrated to play pivotal roles in acute MIR injury, myocardial inflammation, myocardial fibrosis, and ventricular remodeling [25, 26]. However, due to the phenotypic heterogeneity and functional complexity of macrophages, especially in intricate pathophysiological contexts, the precise role of macrophages remains to be elucidated. In this study, by integrating mass cytometry with traditional foundational research methodologies, we underscored the significant role of infiltrating macrophages in the progression from MIR to fibrosis. Our novel findings include the following: (1) GMCSF serves as the primary chemotactic factor for macrophage migration to the heart post MIR; (2) GMCSF/CCL2-recruited CCR2+ macrophages can activate the NLRP3/Caspase-1/IL-1β signaling pathway, amplifying the inflammatory response; and (3) CCL2-mediated CCR2+ macrophages transform into a reparative M2 phenotype and release Tgf-β, promoting fibrosis progression (Fig. 9).

Fig. 9: After myocardial ischemic injury, cardiac microvascular endothelial cells release a large amount of GMCSF to attract monocytes to migrate to the heart, which differentiate into macrophages and transform into pro-inflammatory M1 phenotype under GMCSF induction, releasing a large amount of inflammatory factors and CCL2.
figure 9

CCL2 recruits a large number of CCR2+ cells to infiltrate the damaged myocardial tissue, releasing inflammatory mediators to promote inflammatory response. CCL2 stimulates CCR2+ macrophages to transform into reparative M2 phenotype, which releases Tgf-β to promote the transformation of fibroblasts into myofibroblasts and release a large amount of extracellular matrix, leading to fibrosis (by Figdraw).

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Macrophages in the heart can be categorized into cardiac resident macrophages (CRMs) and infiltrating monocyte-derived macrophages (IMs). Previous studies have shown that CRMs can effectively mitigate fibrosis and cardiac function decline induced by excessive stress overload following transverse aortic constriction (TAC) surgery [27]. However, the role of IMs in the transition from acute myocardial inflammation to fibrosis remains unclear. Acute ischemic hypoxic injury to the myocardium results in extensive cardiomyocyte necrosis, with a significant release of chemokines and inflammatory mediators, playing a crucial role in recruiting bone marrow-derived monocytes in response to cardiac injury signals [28]. In the early stages of acute injury, these infiltrating macrophages predominantly differentiate into the M1 proinflammatory phenotype, releasing inflammatory mediators such as TNF-α and IL-1β, exacerbating inflammatory damage and facilitating necrotic tissue clearance [29]. They also release a plethora of chemokines, such as CCL2 and CX3CL1, which are ligands for CCR2 and CX3CR1 receptors, respectively, promoting the infiltration of CCR2+ cells (including macrophages, DCs, T cells, and polymorphonuclear leukocytes) and CX3CR1+ cells to the myocardial injury site [30, 31].

The tissue microenvironment and temporal dynamics post MIR are critical determinants influencing macrophage infiltration and differentiation. Through a Luminex chemokine array and ELISA, we confirmed that GMCSF is expressed at higher levels in intrinsic cardiac tissue cells than in cardiomyocytes, promoting monocyte migration to the heart and differentiation into infiltrating macrophages. Starvation-induced cellular experiments further elucidated that MCMECs are the primary source of GMCSF, with T cells also releasing minor amounts. GMCSF, with its multifaceted biological functions, not only promotes monocyte infiltration into damaged cardiac tissue but also induces macrophage differentiation into the M1 inflammatory phenotype, exerting inflammatory effects. After GMCSF-recruited macrophages infiltrate the heart, CCL2/CCR2 signaling induces CCR2+ cell infiltration. As early as 2 h post MIR, CCR2+ cells begin to infiltrate the heart, further suggesting that CCR2+ macrophages are the primary inflammatory effector cells. Using GMCSF NTAB to inhibit GMCSF, we effectively suppressed CCL2/CCR2 signaling both in vitro and in vivo. This further confirmed that GMCSF is upstream of the CCL2/CCR2 signal. Interestingly, we observed reduced levels of GMCSF and CCL2 in CCR2−/− mice. This might be attributed to the expression of CCR2 on both T cells and macrophages, with CCR2 gene knockout reducing acute-phase T-cell and macrophage levels, subsequently leading to decreased GMCSF and CCL2 levels. The levels of IL-4 and IL-1β in the heart also confirmed that CCR2+ immune cells, such as T cells and macrophages, are involved in acute MIR injury. Compared to WT mice post MIR, knocking out the CCR2-gene effectively reduced infarction size and injury levels and diminished the reperfusion-induced NLRP3/Caspase-1/IL-1β inflammatory cascade.

Upon entering the chronic fibrosis phase, macrophages predominantly exhibit the M2 phenotype, which suppresses excessive inflammatory responses and promotes fibrotic repair [32, 33]. Myofibroblasts serve as effector cells in fibrotic alterations, releasing abundant extracellular matrix components, such as collagen fibers and fibronectin, and facilitating myocardial tissue repair and ventricular remodeling [34]. Under normal conditions, cardiac fibroblasts present in heart tissue do not possess fibrogenic capabilities. These fibroblasts only transform into myofibroblasts under Tgf-β stimulation, playing a role in fibrosis [17]. Our study revealed that M2 macrophages, compared to other macrophage phenotypes, release a significant amount of Tgf-β, underscoring their irreplaceable role in fibrotic repair and tissue remodeling [35]. However, GMCSF appears to promote the transformation of macrophages into the M1 proinflammatory phenotype, seemingly contradicting the progression of myocardial fibrosis facilitated by M2 macrophages [36]. Intriguingly, CCL2 effectively reverses the M1 phenotype of macrophages induced by GMCSF, promoting their transition to the M2 phenotype. This finding aligns with previous studies on the transformation of macrophages into the M2 phenotype by CCL2 [37, 38].

As a chemokine ligand for CCR2, CCL2 not only promotes the phenotypic transformation of macrophages to aid in fibrotic repair but also recruits CCR2+ immune cells. Our findings indicate that during the M2 transition induced by CCL2, a substantial amount of Tgf-β is released, and the ability of macrophages with knocked-out CCR2 gene to release Tgf-β is diminished. This further suggests that the CCL2/CCR2 axis is pivotal in macrophage phenotypic transformation. The mass cytometry results indicated that on the 7th day post reperfusion, a significant infiltration of CCR2+ cells, including elevated levels of macrophages and DCs, was observed in mouse cardiac tissue. Knocking out the CCR2-gene did not significantly affect the proportions of T cells, B cells, or neutrophils on the 7th day post reperfusion. On the 7th day after MIR, the levels of macrophages and DCs in the hearts of CCR2−/− mice were significantly reduced, with the decreased macrophage phenotype being predominantly M2. This indirectly confirms that CCR2 is a dependency molecule for the M2 phenotype of macrophages and that inhibiting the M2 phenotype can reduce the level of myocardial fibrosis. CCR2 is primarily expressed on the surface of immune cells, and it has been previously established that CCR2 is not expressed on the surface of myofibroblasts. Thus, we ruled out the possibility that CCL2 released by macrophages directly induces the proliferation of CCR2-expressing myofibroblasts to promote fibrotic progression. This finding implies that the infiltration of CCR2+ macrophages is a crucial mechanism influencing fibrotic progression. In the past, in renal fibrosis, it has been demonstrated that inhibiting the infiltration of CCR2+ cells can effectively reduce the level of renal fibrosis [6]. Compared to WT mice undergoing MIR, CCR2−/− mice experienced significantly reduced fibrosis levels and improved cardiac function after MIR. Consistent with changes in cardiac function and fibrosis were the levels of Tgf-β. To further validate in vivo that Tgf-β primarily originates from macrophages, we established an MIR model in MD mice and observed a significant reduction in Tgf-β, further confirming that Tgf-β mainly originates from macrophages. These results all affirm that the GMCSF/CCL2/CCR2 axis can serve as a key therapeutic target to inhibit MIR injury and alleviate myocardial fibrotic repair.

To further explore the pharmacological inhibitory effects of CCR2 on fibrotic transformation after myocardial injury, we treated MIR mice with a CCR2 inhibitor and a vehicle control. The study found that there was no significant difference between the MIR groups with or without the addition of the vehicle control. In contrast, mice treated with the CCR2 inhibitor exhibited significantly reduced myocardial injury and infarct size, along with decreased levels of the NLRP3/Caspase-1/IL-1β signaling pathway. Additionally, the CCR2 inhibitor significantly reduced fibrosis levels after MIR, preserving cardiac function. This may offer a novel therapeutic approach for chronic fibrotic remodeling following myocardial injury.

However, this study has limitations, as it was primarily conducted in animal models without further integration with clinical research. Moreover, our research found that infiltrating macrophages promote fibrosis in the later stages by releasing Tgf-β, but whether infiltrating DCs affect fibrosis in the later stages was not further explored in this study. Therefore, further clinical research is needed to explore GMCSF/CCL2/CCR2 as the therapeutic target in improving fibrosis and the impact of DCs on cardiac fibrosis.

Conclusion

After MIR, GMCSF/CCL2 recruits CCR2+ macrophages to activate NLRP3/Caspase-1/IL-1β-mediated and amplified acute-phase inflammation. Numerous CCL2 mediate CCR2+ macrophage transformation into the M2 phenotype and release Tgf-β to promote myocardial fibrosis.