P2X4 receptor limits adverse cardiac remodeling following ischemia–reperfusion through regulation of autophagy in cardiac fibroblasts

Abstract

Background

Pathological ventricular remodeling is an adverse tissue response following acute myocardial infarction (AMI), characterized by an exaggerated fibrosis leading to tissue disorganization and chronic heart failure. Cardiac fibroblasts have a prominent role in the healing process, but also in the fibrosis development, through their differentiation into myofibroblasts, which produce the extracellular matrix, in response to ischemia–reperfusion. Recently, the ATP-gated P2X4 ionotropic receptor has been identified as a key regulator of macro-autophagy, activated under metabolic challenges.

Methods

In this study, we investigated the participation of the P2X4 receptor in the metabolic response of cardiac fibroblasts to ischemia–reperfusion and the myofibroblast switch. To this end, we submitted a primary culture of cardiac fibroblasts to an ischemia–reperfusion simulation and assessed autophagic flux, fibrotic differentiation markers and the correlation to P2X4 expression and activity. Then, using pharmacological inhibition and siRNA-mediated knockdown receptor under-expression, we investigated its role in the response of cardiac fibroblasts to this metabolic challenge. Finally, we validated our findings in an in vivo murine model of coronary ligation in wild-type and P2rx4^−/− mice.

Results

Our results reveal that human cardiac fibroblasts subjected to simulated ischemia present an increased autophagic activity associated with a concomitant overexpression of P2X4 and its activity at the plasma membrane of cells. Pharmacological antagonism of P2X4, or its silencing, impaired autophagy, as evidenced by the accumulation of autophagy markers, and induced the expression of fibrotic markers. In an in vivo model of AMI through transient coronary ligation, P2rx4^−/− mice presented the accumulation of autophagic markers both in healthy and ischemic zones, an extended tissue inflammation and fibrosis compared to wild type mice.

Conclusions

This study suggests, for the first time, that the P2X4 receptor may contribute to steering cardiac fibroblasts toward a healing-associated profile under ischemia–reperfusion conditions, possibly by modulating autophagic activity and limiting extensive pro-fibrotic differentiation.

Supplementary Information

The online version contains supplementary material available at https://doi.org/10.1186/s12964-026-03050-4.

Introduction

Acute myocardial infarction (AMI), caused by the sudden occlusion of a coronary artery, is responsible for ischemic damages leading to cardiomyocytes necrosis and tissue inflammation [1]. Its clinical management with urgent and stable coronary reperfusion, though necessary, may cause additional damages called reperfusion injuries [2]. The size of the initial necrotic area and the subsequent remodeling induced by the healing process are major prognosis factors, correlated with the successful restoration of myocardial contractile function and patient survival [3].

Cardiac fibroblasts, as resident cells, have a prominent role in the healing process through their differentiation into myofibroblasts, producing the extracellular matrix [4]. However, this capacity must be tightly controlled to avoid any pathological fibrotic process. This phenotypical and functional plasticity of cardiac fibroblasts relies partially on their capacity to sense and adapt to constraints in the environment, such as metabolic pressure induced by ischemia and activate surviving pathways, including macro-autophagy.

Macro-autophagy, hereafter called autophagy, is a catabolic process whereby cells degrade misfolded materials that are present in the cytosol [5]. Degradation of long-lived or misfolded proteins as well as damaged organelles allows the recycling of fundamental nutrients such as amino acids, necessary for cellular homeostasis. In cases of energetic substrate deprivation, e.g. during an ischemic event, cells can also degrade their own metabolic reserves, such as lipid droplets, and compartments to maintain a relative level of energy production and support cell survival for a short period of time [6]. Levels of Autophagy related protein 5 (ATG5) were previously reported to be higher in the serum of patients with coronary artery disease or AMI compared to healthy controls [7]. In addition, it has been shown that a defect in autophagy, combined with a weakening of mitochondrial activity, may have detrimental effects on the survival of cardiomyocytes during hypoxic or ischemic episodes [8, 9]. Similar conclusions were drawn in an in vivo murine model of AMI in which the administration of bafilomycin A1, an inhibitor of the autophagic process, significantly increased the infarct size [10]. Furthermore, genetic ablation of the AMP-activated protein kinase (AMPK) signaling pathway, leading to a defective autophagy, resulted in the increase in infarct size in murine models of AMI [11, 12]. Autophagy may also have a crucial role in the response to reperfusion, which is known to induce additional lesions [2]. Indeed, the induction of autophagy by rapamycin, administered at the time of reperfusion in a murine model of AMI, inhibited mTORC1 and p38 signaling and improved post-infarct survival and cardiac function [13]. It also decreased NF-kB activation and attenuated the sterile inflammatory response correlated with adverse ventricular remodeling [14, 15]. In addition, the inhibition of NF-kB in vitro has been demonstrated to restore the autophagy flux [16]. Taken together, these results identified autophagy as an innate and potent process supporting cardioprotective effects against ischemia–reperfusion injuries.

Mechanisms inducing autophagy during ischemia appear to be different than those involved after reperfusion. The energy restriction, which massively stimulates autophagy in the heart during the ischemic period, is at least partially resolved at the time of reperfusion. Instead, there is a large production of reactive oxidative species (ROS) which might promote autophagy during reperfusion [17]. However, the effect of ROS on autophagy process is still unclear. Indeed, ROS produced during reperfusion could also impair the clearance of autophagosomes and be responsible for cardiomyocyte death [18].

Surprisingly, the literature on autophagy in myocardial ischemia–reperfusion essentially focused on its role on cardiomyocyte survival. However, it is now well established that patients outcomes and risk of chronic heart failure, due to persistent myocardial dysfunction, not only depend on the extent and severity of the initial myocardial necrosis, but also on the effectiveness of the post-infarct tissue healing [19, 20]. We then questioned the role of autophagy in cardiac fibroblasts phenotypic changes under ischemia–reperfusion, and the potential consequence of its modulation on the development of post-infarction fibrosis. During the ischemia and reperfusion phases, cardiac fibroblasts are subjected to a harmful environment containing damage-associated molecular patterns (DAMPs) released by necrotic and suffering cells, e.g. extracellular Adenosine 5’-triphosphate (eATP). Extracellular ATP can activate P2 purinergic receptors expressed at the plasma membrane of cells. These receptors are ubiquitously distributed, and we hypothesized that they could modulate the phenotype and activity of cardiac fibroblasts. Furthermore, many members of the P2 receptor family have recently been demonstrated to play a key role in the post-myocardial infarction inflammatory response [21]. Among them, our present work focuses on the role of the ATP-gated P2X4 ionotropic receptor. Interestingly, the P2X4 receptor, which is mainly expressed in endosomal and lysosomal related-organelles, has been recently demonstrated to promote autophagy under metabolic stress in another cell model [22, 23].

In this study, we show that the P2X4 receptor is upregulated under ischemic conditions and that autophagy is similarly induced during ischemia–reperfusion in human cardiac fibroblasts. Pharmacological inhibition or silencing of P2X4 further enhanced ischemia–reperfusion-induced differentiation of cardiac fibroblasts into myofibroblasts and impaired autophagic activity. In an in vivo model of acute myocardial infarction (AMI) in C57BL/6 wild type mice, we observed a modest increase in fibrotic markers within ischemic regions compared with non-ischemic myocardial tissue at seven days post-infarction. By contrast, P2rx4^−/− mice subjected to the same procedure exhibited exacerbated inflammation and more extensive fibrosis. Collectively, our results suggest that P2X4 may exert a protective role by limiting the phenotypic transition of cardiac fibroblasts and attenuating post-ischemic cardiac remodeling, potentially through the regulation of autophagy.

Results

Human cardiac fibroblasts survive ischemia–reperfusion and present an active autophagy flux

We first studied the viability of human primary cardiac fibroblasts under normoxia or a 5-h simulated ischemia protocol (Fig. 1a) using Annexin V – Pi labelling in flow cytometry. There was no difference in cell survival between these two conditions (Fig. 1b, normoxia {3.5% Annexin V- PI +; 0.96% Annexin V + Pi +; 88% Annexin V- PI-; 2.36% Annexin V + PI}, ischemia 5 h {5.8% Annexin V- PI +; 0.73% Annexin V + Pi +; 92.67% Annexin V- PI-; 0.46% Annexin V + PI-}, ns). These initial results demonstrated that fibroblasts could survive ischemia for 5 h, probably through that activation of survival pathways, giving them the ability to resist these cellular stresses.

Fig. 1.

Cardiac fibroblasts survive under restrictive conditions and present an activated autophagic flux. (a) Schematic representation of the in vitro experimental procedure using a primary culture of human cardiac ventricular fibroblasts. (b) Annexin V-Pi staining by flow cytometry showing the viability of cardiac fibroblasts according to the duration of ischemia (n = 5 independent experiments). (c) Electron microscopy micrographs of cells in (1, i) normoxia, (2, ii) ischemia 5 h and (3, iii) ischemia 5 h treated with Bafilomycin A1 (100 nM). Autophagic vacuoles are indicated with yellow arrows (scale bar, 5 or 2 µm). (d) Representative epifluorescence images of cells transfected with RFP-GFP-LC3 (scale bar, 50 µm). (e) Quantification of the number of LC3-positive dots RFP⁺ and RFP⁺GFP⁺(n = 30 per condition from 3 independent experiments) (scale bar: 50 μm). (f) Representative western blot of LC3-II and p62 in human cardiac fibroblasts after CQ (100 μM) or Bafilomycin A1 (100 nM) treatment in normoxia or after 5-h ischemia. (g) LC3-II and p62 protein expression assessed as in (d) in human cardiac fibroblasts cultivated in normoxia or 5-h ischemia with or without Bafilomycin A1 (100 nM), expressed relatively to Hsc70 (n = 4–6 independent experiments). (h) Autophagic flux was calculated as the ratio of LC3-II expression with and without bafilomycin A1 (n = 4–6 independent experiments). Graphs show box plots or median with interquartile range, and results were compared using a Mann–Whitney test, *p < 0.05, **p < 0.01. ns stands for no statistical difference

Transmission electron microscopy images of cardiac fibroblasts revealed the presence of autophagic vacuoles under normoxic conditions while, under simulated ischemia, the same structures were observed but with less content (Fig. 1c, yellow arrows), suggesting an increased turn-over. The use of Bafilomycin A1, an inhibitor of the V-ATPase lysosomal proton pump, led to the accumulation of autophagic vacuoles undergoing degradation, demonstrating the presence of an active autophagy in ischemic conditions. Similarly, the measure of autophagosome clearance using RFP-GFP tandem fluorescent-tagged LC3 (Fig. 1d, e) is consistent with an enhanced autophagic activity under ischemia relative to normoxia (median: 3{2: 4} vs median: 7.5 {4: 9.5} respectively, p = 0.001). This clearance was inhibited by the use Bafilomycin A1 (median: 9.5 {4: 16} respectively, p = 0.001).

To confirm these observations, we assessed levels of LC3-II and p62 by Western Blotting (Fig. 1f, g). LC3-II relative expression significantly decreased during ischemia comparatively to normoxia (median: 0.06686 {0.01384: 0.1304} vs median: 1.033 {0.8940: 1.791} respectively, p = 0.0095). LC3-II protein expression was rescued with the use of Bafilomycin A1 in ischemic conditions (normoxia + Baf A1 median: 1.622 {1.112: 3.497}, ns; ischemia + Baf A1 median: 1.120 {1.008: 1.389}, p = 0.0022). This allowed us to calculate an autophagic flux (Fig. 1h) that was significantly increased in ischemia compared to normoxia (median: 25.41 {9.983: 63.70} vs 1.802 {1.049: 2.022}, p = 0.028). The expression of p62 was also significantly decreased during ischemia compared to normoxia (median: 0.3481 {0.2007: 0.4922} vs 1.101 {0.08819: 1.732}, p = 0.0095) and rescued by the addition of Bafilomycin A1 (normoxia + Baf A1 median: 1.878{1.134: 4.445}, ns; ischemia + Baf A1 median: 1.643{0.7190: 2.033}, p = 0.022).

P2X4 expression increases following simulated ischemia

We recently demonstrated that the P2X4 purinergic receptor was a key regulator of autophagy during nutrients and oxygen deprivation [23]. Yu et al. performed bulk RNA Sequencing (GSE236374) from total murine hearts at different times after a temporary coronary artery ligature [24]. A closer study of P2X receptor expression showed that P2rx4, P2rx5, and P2rx7 genes were the major to be expressed in sham models and that only P2rx4 and P2rx7, encoding for P2X4 and P2X7 respectively, were overexpressed seven days after AMI (Fig. 2a). These increases persisted at 28 days post-infarction. However, in this experiment, the analysis was performed on total heart extractions containing different cell types. We then analyzed another database of RNA sequencing experiments (GSE132146) performed by Ruiz-Villalba et al. from murine cardiac fibroblasts, specifically isolated, at different times post-AMI, i.e. at 7, 14 and 30 days [25]. Our analyses demonstrated that P2rx4 was mostly expressed in cardiac fibroblasts, and its level of expression was significantly increased at seven days post-AMI and was still high at 14 days. At 30 days post-AMI, P2rx4 mRNA level was back to control level (Fig. 2b).

Fig. 2.

P2X4 expression increases under simulated ischemia. (a, b) Expression of P2X receptors assessed by RNA-seq [24, 25] after acute myocardial infarction in (a) total mice heart or (b) in isolated mouse cardiac fibroblasts. (c) P2X4 mRNA expression in human cardiac fibroblasts after ischemia 5 h or ischemia 5 h-reperfusion (1, 3, 5 24 and 48 h) normalized to normoxia (n = 7–11 independent experiments). (d) Immunochemistry representative images of P2X4, Lysotracker and Lamp-1 staining in human cardiac fibroblasts cultivated in normoxia. (e) Representative western blot for P2X4 expression in normoxia and 5-h ischemia (2 independent experiments). (f) Quantification of P2X4 protein expression after 5-h ischemia or ischemia–reperfusion (24 and 48 h) normalized to normoxia (n = 11 independent experiments). (g) Representative variations in Fluo4 AM signal after stimulation with 10 μM ATP in presence or not of BX 430 (25 μM) (n = 3 independent experiments). (h) Quantification of fluorescence variation after 10 μM ATP stimulation alone or with BX 430 (25 μM), as in (g), normalized to cells basal fluorescence for each condition. (i) P2X4 expression in siCTL or siP2X4 treated cells assessed by western blot. (j) Representative variation in Fluo4 AM signal after stimulation with 10 μM ATP in control and silencing P2X4 cells. (k) Quantification of fluorescence variation after 10 μM ATP stimulation in control and silencing P2X4 cells normalized to cells basal fluorescence for each condition (n = 3 independent experiments). Graphs show in box plots and results were compared using Wilcoxon-ranked or Mann–Whitney test, *p < 0.05, **p < 0.01, ***p < 0.001

Following these analyses, we wondered about P2X4 expression in our in vitro model of primary culture of human cardiac fibroblasts submitted to simulated ischemia–reperfusion.

We observed that P2X4 mRNA was significantly overexpressed at the end of the 5 h-ischemia (median: 3.095{2.125: 4.57}, p = 0.0026), comparatively to normoxia. After one to five hours of reperfusion, P2RX4 mRNA levels were comparable to normoxia and even reduced at both 24 and 48 h of reperfusion (Median: 0.5150 {0.4182: 0.9158}, p = 0.0107; and 0.64 {0.24:0.83}, p = 0.0414, respectively, Fig. 2c). Co-staining with P2X4/Lysotracker and P2X4/Lamp-1suggested lysosomal enrichment of P2X4 (Fig. 2d). P2X4 protein level, from total cell extract, was significantly increased at the end of ischemia (Fig. 2e and f) (median:1.370 {1.111: 2.205}, p = 0.0137) and its expression was maintained at 24 h of reperfusion (median:1.501 {0.9280: 2.59}, p = 0.0234) compared to normoxia. Interestingly, at 48 h of reperfusion, P2X4 protein expression decreased compared to normoxia (median: 0.587 {0.4236: 1.123}, p = 0.0303, Fig. 2e, f), suggesting a rapid recycling and relatively short half-life of the protein.

After evaluating the protein expression of P2X4, we questioned its activity at the plasma membrane of cardiac fibroblasts. We measured the Ca²⁺ influx in control or ischemia conditions and after the addition of 10 µM ATP, using the Fluo-4 probe. In normoxia, ATP elicited an increase of intracellular Ca²⁺ content that was maintained over 60 s (normoxia median = 21.03 {18.06: 30.81}). This increase was not prevented using selective human P2X4 antagonist BX 430, suggesting the participation of other P2 receptors, most probably G-coupled P2Y receptors. Interestingly, cells subjected to 5 h-ischemia demonstrated a higher ATP-induced Ca²⁺ response (5 h-ischemia median = 52.28 {42.54: 58.71}, p < 0.0001), which was prevented by the use of BX 430 (5 h-ischemia + BX 430 (median = 34.23 {27.93: 37.95}, p = 0.003) (Fig. 2g and h), suggesting an increased level of plasmalemmal P2X4 receptors. Similar results were obtained after reperfusion, when the addition of BX 430 decreased the fluorescence (median BX 430: 37.07 {34.2: 40.77}, median untreated cells: 48.09 {45.8: 69.2}, p < 0.001) (Fig. 2g and h). To confirm these results, we aimed at specifically knocking-down the expression of P2X4 in cardiac fibroblasts. The use of specific siRNA targeting P2X4 expression (siP2X4) effectively reduced the P2X4 protein level by 56% (median: 0.44, p = 0.04) comparatively to the transfection with a null-target siRNA (siCTL) (Fig. 2i).

Transfection with siP2X4 did not affect the ATP-induced increase in intracellular Ca²⁺ under normoxic conditions (normoxia, siCTL: 31 [29.38–33.97] vs. siP2X4: 30.98 [24.43–33.87], ns). However, following 5 h of ischemia, an increase in ATP-induced cytosolic Ca²⁺ responses was observed in siCTL cells, whereas this effect was abolished in siP2X4-transfected cells (median: 32.93 [29.3–37.9], p = 0.04) (Fig. 2j, k). Moreover, P2X4 knockdown in cardiac fibroblasts resulted in a lower fluorescence signal following reperfusion compared with control cells (siP2X4: median 33.68 [21.51–43.13] vs. siCTL: median 58.45 [55.8–74.18], p = 0.0022).

P2X4 regulates ischemia-induced autophagy in human cardiac fibroblasts

We next assessed autophagic activity in cardiac fibroblasts subjected to 5 h of ischemia following pharmacological inhibition of P2X4 with BX430 or gene silencing using siP2X4. Transmission electron micrographs showed that, after ischemia, the inhibition or silencing of P2X4 was responsible for an increased proportion of autophagic vacuoles with incompletely digested content (yellow arrows, Fig. 3a). Impaired autophagic flux was further evidenced by defective autophagosome clearance measured with the RFP–GFP tandem fluorescent-tagged LC3 in P2X4-inhibited versus untreated cells (I 5 h median: 3 {2: 4} vs I 5 h + BX 430 median: 9.5 {4: 16.5} respectively, p = 0.001; I 5 h siCTL median: 3 {1: 5,2} vs I 5 h siP2X4 median: 6.5 {5: 8.25} respectively, p = 0.001) as shown in Fig. 3b and c.

Fig. 3.

P2X4 regulates ischemia-induced autophagy in human cardiac fibroblasts. (a) Electron microscopy micrographs of cells after (1, i) 5 h ischemia (2, ii) 5 h ischemia treated with BX 430 (5 μM) and (3, iii) 5 h ischemia in cells transfected with siP2X4 (scale bar, 2 µm). Yellow arrows indicate autophagic vacuoles. (b) Representative epifluorescence images of cells transfected with RFP-GFP-LC3 (scale bar, 50 µm). (c) Quantification of the number of LC3-positive dots RFP⁺ and RFP⁺GFP.⁺ (n = 30 per condition from 3 independent experiments) (scale bar: 50 μm). (d) Representative western blots of LC3-II and p62 expression in ischemia conditions in human cardiac fibroblasts either treated with BX 430 (5 μM) or under-expressing P2X4. (e, f) Quantification of LC3-II and p62 expression by western blot in human cardiac fibroblasts treated with BX 430 (5 μM) (e) or knocked-down for the expression of P2X4 (f). Data are normalized to untreated ischemia condition. (n = 6–10 independent experiments) Graphs show median with interquartile range and results were compared using Wilcoxon-ranked test, *p < 0.05

Similarly, defective autophagosome clearance was demonstrated by the accumulation of LC3-II and p62 following ischemia in P2X4-inhibited versus untreated cells (Fig. 3d). For cells treated with BX 430, we quantified a significant accumulation of LC3-II protein (I 5 h median: 2.297 {1.504: 2.731}, p = 0.0312}, IR {median: 1.938 {1.619: 4.032}, p = 0.0312}) (Fig. 3e). Similar results were observed for p62 expression (I 5 h median:1.38 {1.127:1.943, p = 0.0312}, IR median: 1.559{1.1: 10.35}, p = 0.0312}).

Compared to ischemia alone and ischemia–reperfusion siCTL condition, fibroblasts treated with siP2X4 presented a significant accumulation of LC3-II protein in both ischemia and ischemia–reperfusion conditions (5 h-ischemia siP2X4, I 5 h siP2X4, median: 1.977 {1.097: 3.63}, p = 0.0469}, 5 h-ischemia-24 h-reperfusion siP2X4, IR siP2X4, median: 1.336 {1.09: 2.941}, p = 0.0312}) (Fig. 3f). For p62 expression, there was also accumulation following ischemia and ischemia–reperfusion in siP2X4 cells compared to siCTL (I5h median: 1.160 {1.025: 1.99}, p = 0.0322}, IR median: 1.312 {1.029: 1.874}, p = 0.0137). The same results were observed with the use of another P2X4 antagonist, 5-BDBD (Fig. S2b). Interestingly, this regulation of autophagy mediated by P2X4 appeared AMPK-independent (Fig. S3).

P2X4 inhibition or silencing enhances ischemia-induced differentiation of fibroblasts into myofibroblasts

Cardiac fibroblasts are very sensitive to their microenvironment conditions and adapt rapidly to changes through phenotypic and functional transitions (Fig. 4a). Following simulated ischemia–reperfusion, we observed a differentiation of fibroblasts into myofibroblasts, as shown by the increase of α-Smooth Muscle Actin (α-SMA) expression (Fig. S1). This functional switch is identified as a critical step in the development of fibrosis observed in pathological left ventricular remodeling [4]. We previously identified a role for P2X4 in ischemia-induced autophagy in cardiac fibroblasts and asked whether this could also interfere with their differentiation in myofibroblasts. Following a 5 h-simulated ischemia, cardiac fibroblasts significantly overexpressed pro-fibrotic markers α-SMA but not collagen-I either in BX 430 (median: 1.289, p < 0.01; 0.88, ns, respectively) or siP2X4 (median: 1.423, p = 0.016; 1.336, ns, respectively) conditions compared with the appropriate control condition (siCTL or vehicle, respectively, Fig. 4c and d). This overexpression was maintained at 24 h of simulated reperfusion for BX 430 treatment (median: 1.389, p = 0.03; 1.420, p = 0.125 respectively) and siP2X4 cells for both markers (median: 1.616, p = 0.015; 1.434, p = 0.012 respectively) or (Fig. 4e and f). These data were confirmed with the use of another P2X4 antagonist, 5-BDBD (Fig. S2c, d.). Similar results were obtained using Bafilomycin A1 treatment, to inhibit autophagy, both in normoxia and simulated ischemia conditions, leading to α-SMA and collagen-I accumulation (Fig. S4).

Fig. 4.

P2X4 antagonism, or silencing, potentiates fibroblast differentiation into myofibroblast under ischemia. (a) Schematic representation of fibroblast phenotypic change and associated features [26]. (b) Representative western blot for α-SMA and Collagen-I expression in ischemia conditions in fibroblasts either treated with BX 430 (5 μM) or under-expressing P2X4. (c-f) Quantification of α-SMA and Collagen-I expression in cardiac fibroblasts under-expressing P2X4 and cardiac fibroblasts treated or not with BX 430 (5 μM) after ischemia alone (c, d) or ischemia–reperfusion (24 h) (e, f) (n = 7–9 independent experiments). (g, i) Representative images of Scratch Assay during time for after each protocol. (h, j) Quantification of coverage rate as a function of initial scratch surface (n = 3–4 independent experiments). Graphs show median with interquartile range or box plots and results were compared using Wilcoxon-ranked test, Mann–Whitney-test or two-Way ANOVA with Bonferroni-correction, *p < 0.05, **p < 0.01, ***p < 0.001

To further investigate the role of P2X4 in fibroblast properties, we performed a wound-healing assay during the reperfusion phase following simulated ischemia or normoxic conditions to assess migrative capacity, as determined by the progressive closure of the cell-free area over time (Fig. 4g and i). Of note, EDU labeling experiments showed only ~ 40% positivity, reflecting a low baseline proliferation rate. Compared to normoxia alone (median: 2.243–2.795), the addition of the profibrotic agent TGF-β 1 (5 ng/mL), which leads to myodifferentiation, did not affect migratory capacities of cardiac fibroblasts (median: 1.681–2.01, ns). However, fibroblasts previously exposed to ischemia demonstrated higher migration capacities after 12 h and 18 h of reperfusion (median: 5.281–4.505, respectively, p < 0.001) as compared to normoxia. Interestingly, the inhibition of P2X4 using BX 430 or its silencing with siP2X4 reduced fibroblast migration rate induced by ischemia (BX 430 treated-median: 2.812–3.447, p < 0.01, and siP2X4-median: 2.451, p = 0.02; 2.307, p < 0.01, respectively) (Fig. 4h, j).

P2X4 deficiency impairs autophagy and exacerbates fibrosis in murine model of acute myocardial infarction

To assess the role of P2X4 in post-infarct left ventricular remodeling, we developed an in vivo model of AMI using wild-type C57BL/6 J (WT) and P2rx4^−/− mice from the same strain (Fig. 5a). Interestingly, we observed an increased expression of the P2X4 protein in the infarcted zone of WT, compared to the healthy zone (Fig. S5). This result is in line with previous RNA sequencing analyses (Fig. 2a).

Fig. 5.

P2X4 knock-out impairs autophagy and increases fibrosis in an in vivo murine model of acute myocardial infarction. (a) Schematic representation of the experimental procedure of acute myocardial infarction in vivo. (b) Representative western blot of LC3-II and p62 in healthy and infarcted zones for Wild-Type and P2rx4.^−/− mice. (c) Quantification of LC3-II and p62 expression by western blot in healthy and infarcted zone collected from mice hearts at 7 days post-infarction (n = 7–8 independent experiments). (d) Images the left ventricle sections after Evans blue and TTC staining observed with a binocular magnifying glass. (e) Percentage of necrosis area normalized to at-risk area determined with TTC coloration (n = 3–7 independent experiments). (f) Left-ventricle weight (mg) normalized to body weight (g). (g) Representative western blot of α-SMA and Collagen-I in healthy and infarcted zones of the two models. (h) Quantification of α-SMA and Collagen-I expression by western blot in healthy and infarcted zone collected from mice hearts at 7 days post-infarction (n = 7–9 independent experiments). (i, k) Representative images of Hematoxylin and Eosin (i) or Red Sirus staining (k) (grey arrows indicate pathological area). (j,l) Percentage of inflammatory and fibrosis areas normalized to total ventricle area (n = 5–15 independent experiments). Graphs show median and results were compared using Mann–Whitney test, *p < 0.05, **p < 0.01

We then investigated whether the expression of P2X4 could modulate the ischemia–reperfusion – related responses in vivo. At seven days after AMI, P2rx4^−/− mice presented a significantly higher expression of LC3-II (median: 1.735 {0.778: 2.805} vs 0.707 {0.317: 0.991}, p = 0.036) and p62 (median: 1.335 {0.873: 1.545} vs 0.515 {0.307:1.018}, p = 0.012) in infarcted zones, compared to WT mice. This suggested an altered autophagic process, as evidenced by the accumulation of autophagy markers in the left ventricular tissue (Fig. 5b, c). However, this potential defect had no impact on the survival of animals, over the duration of the experiment (7 days) or on the extent of left ventricular infarct size, which was comparable in the two groups (median WT: 5.125{4.644;5.325}, vs P2rx4^−/−: 5.254 {4.195;6.695} respectively, ns) (Fig. 5d, e). Of note, left ventricles coming from P2rx4^−/− mice had a tendency, yet not significant with our size sample, to be heavier than those of the wild-type (median: 6.747 g {5.288; 10.577}, vs 4,885 g {3.934;5.358} respectively, p = 0.083), suggesting the onset of ventricular hypertrophy and the development of adverse ventricular remodeling (Fig. 5f). To confirm this hypothesis, we quantified the levels of fibrotic markers in either healthy or infarcted zones by Western Blotting. There was again a tendency, yet not significant, for the increase of α-SMA (median: 1.519{0.652;2.412} vs 0.962{0.709;1.219} p = 0.072) and a significant increase of Collagen-I (median: 1.162{0.142;1.162} vs 0.027{0.02;0.187}, p = 0.007) in infarcted zones in P2rx4^−/− compared to WT mice (Fig. 5g, h). This confirmed the tissue remodeling, that was further highlighted at the macroscopic level with histological staining: P2rx4^−/− mice exhibited a larger fibrotic area identified by Sirus Red staining compared to WT (median = 6% {4.708; 7.639}, median = 0% {0;2.351}, p = 0.031) as well as a higher inflammatory cell infiltration, assessed by Hematoxylin and Eosin staining (median = 4.924% {3.802; 9.647}, median = 0% {0; 2.361}, p = 0.0192) (Fig. 5j, l). Furthermore, P2rx4^−/− mice demonstrated an increase in levels of circulating IL-1β, IL-23 and IL-17 in the serum (Fig. S6), suggesting a Th17 lymphocyte orientation [27].

Discussion

While the capacity of cardiac fibroblasts to survive ischemic stress and contribute to tissue repair following myocardial injury is well established, the role of autophagy in regulating their response to ischemia–reperfusion injury remains incompletely understood. The present study provides evidence for the first time supporting the involvement of the P2X4 receptor in shaping the phenotype of cardiac fibroblasts following AMI, potentially through the regulation of autophagic processes. This role may be particularly relevant, as fibroblast fate after ischemic injury critically influences the balance between adaptive scar formation and maladaptive left ventricular remodeling.

Our findings demonstrate the activation of autophagy in a primary culture of human cardiac fibroblasts subjected to an in vitro ischemic model combining hypoxia and nutrient deprivation. This response was associated with enhanced differentiation of fibroblasts into myofibroblasts under ischemia. In parallel, we observed a dynamic regulation of the ionotropic receptor P2X4 throughout the ischemia–reperfusion process, characterized by marked upregulation during ischemia followed by downregulation during reperfusion. These expression dynamics suggest that P2X4 may contribute to the adaptation of cardiac fibroblasts to metabolic stress.

P2X4 displays a dual subcellular localization and function depending on organelle trafficking dynamics. Following membrane trafficking, a fraction of the receptor can localize to the plasma membrane, where its stimulation with ATP promotes Ca²⁺ influx and downstream Ca²⁺-dependent signaling pathways [28]. However, the present study does not distinguish between extracellular Ca²⁺ influx and Ca²⁺ release from intracellular stores, thereby limiting mechanistic conclusions regarding the role of plasmalemmal P2X4. Importantly, P2X4 has been demonstrated to be predominantly localized to lysosomal membranes, where it facilitates lysosome–membrane fusion through a Ca²⁺-dependent activation of the SNARE complex [29]. Through this lysosomal localization, P2X4 may regulate lysosomal function and lysosome-dependent pathways, including autophagy, as previously reported in breast cancer cells under hypoxic stress [30]. Nevertheless, its contribution to autophagic regulation under ischemic conditions in cardiac cells remains largely unexplored.

In our experimental setting, both pharmacological antagonism and siRNA-mediated silencing of P2X4 impaired autophagic flux in human cardiac fibroblasts, consistent with observations reported in other contexts of metabolic stress [23]. Based on these findings, we hypothesize that P2X4 may contribute to maintain the balance between pro- and anti-fibrotic signaling in cardiac fibroblasts through the regulation of autophagic activity, thereby influencing post-ischemic remodeling. Supporting this hypothesis, P2X4 inhibition was associated with reduced autophagic flux, accumulation of pro-fibrotic markers, and impaired cardiac fibroblasts migratory capacity, collectively suggesting a shift toward a more differentiated myofibroblast phenotype. Given the central role of fibroblasts in extracellular matrix deposition and scar maturation, such alterations may substantially influence cardiac repair outcomes after ischemic injury.

Consistent with our in vitro observations, P2rx4^−/− mice subjected to AMI exhibited features associated with adverse cardiac remodeling, including enhanced inflammation and increased fibrosis at seven days post-infarction. Interestingly, autophagic markers accumulated not only within the infarcted regions but also in adjacent non-ischemic myocardial tissue in P2rx4^−/− mice, a pattern not observed in wildtype animals. Collectively, these findings suggest that dysregulated autophagy, in the absence of P2X4, may contribute to maladaptive fibroblast activation and pathological tissue remodeling. Notably, these effects appeared to occur independently of AMPK signaling, supporting a potentially distinct lysosome-associated regulatory function of P2X4, where the receptor appears to be predominantly enriched.

Previous work from Nagaraju et al. demonstrated that cardiac fibroblasts isolated from failing hearts exhibit a myofibroblast phenotype, highlighting the clinical relevance of regulating fibroblast differentiation to limit progression toward maladaptive remodeling [31]. In this context, modulation of autophagy may represent a promising therapeutic opportunity for preventing or attenuating post-infarction ventricular remodeling. However, the relationship between autophagy and cardiac repair is complex, as excessive autophagic activation may also promote cell death and aggravate tissue injury [29]. Therefore, preserving an appropriate balance in autophagic activity appears critical.

In this context, P2X4 emerges as potentially relevant therapeutic target for the modulation of autophagic flux under ischemic conditions. Supporting this concept, Yang et al. demonstrated that P2X4 deficiency induces systolic ventricular dysfunction in experiment models of heart failure and highlighted a cardioprotective role of P2X4 in cardiomyocytes [30]. Furthermore, cardiac-specific overexpression of P2X4 has been associated with improved contractile function following AMI induced by left anterior descending coronary artery ligation, with beneficial effects observed at seven days, one month, and two months post-infarction compared with wildtype animals [32]. In agreement with these studies, P2X4 silencing in our model induced deleterious effects comparable to those observed following pharmacological inhibition of autophagy during ischemia–reperfusion, further supporting a protective role of P2X4 signaling in cardiac injury.

In addition to our study supporting a role of P2X4 in autophagy and fibroblast phenotypic regulation, other findings also suggest a potential contribution of P2X4 to post-ischemic immune modulation. Specifically, loss of P2X4 was associated with a shift toward a Th17-type inflammatory profile, characterized by increased secretion of IL-1β, IL-23 and IL-17. Chang et al. demonstrated that the Th17 response promoted cardiac ventricular remodeling in ischemic heart failure [33]. Together, these observations suggest that P2X4 may influence pathological remodeling through both fibroblast-dependent and immune-mediated mechanisms.

Despite the novelty of these findings, several limitations should be acknowledged. Although pharmacological inhibition and siRNA-mediated silencing consistently support a role for P2X4, rescue experiments restoring receptor expression would further strengthen causal specificity. In addition, in vivo analyses could be expanded through echocardiographic assessment and longer-term functional follow-up to better characterize cardiac performance and define the contribution of P2X4 to chronic remodeling. Finally, sex-stratified analyses would improve the robustness and translational relevance of the in vivo findings.

In conclusion, our results suggest a previously unrecognized protective role of P2X4 in limiting pathological left ventricular remodeling following myocardial infarction. P2X4 expression is dynamically regulated in cardiac fibroblasts exposed to ischemic stress and appears to be associated with autophagic activation. Through the modulation of autophagic flux, P2X4 may contribute to maintaining the balance between pro-fibrotic signaling and reparative responses, thereby regulating fibroblast-to-myofibroblast differentiation and limiting excessive fibrotic remodeling after ischemic injury.

Material and methods

Reagents, antagonists

Adenosine Triphosphate, bafilomycin A1 and TGF-β1 were purchased from Sigma-Aldrich, France (#A2383; #SML1661; #T7039). Annexin V-Pi kit, Fluo-4 AM were purchased from Invitrogen (#V13242; #F14201). BX 430 (#5545), 5-BDBD (#3579) and Suramin (#1472) were purchased from Tocris Bioscience, Bristol, United Kingdom. Lysotracker Red DND-99 (#L7528) and Click-iT Plus EdU Flow Cytometry Assay Kit (#C10632) were purchased from Invitrogen (France).

Cell culture

In vitro studies were performed with a primary culture of human cardiac fibroblast of ventricular origin (ScienceCell, Carlsbad, CA, USA, #6310) cultured according to manufacturer’s recommendations. All the experiments were conducted between passages 2-5 to limit spontaneous differentiation into myofibroblast. This parameter was verified with a-SMA and collagen expression levels in control condition.

Ischemia-reperfusion simulation

To reproduce O₂ and nutrients deprivation, which are key features of ischemia, cells were placed in a hypoxic chamber INVIVO (Ruskinn Technology) set with 1% O₂, 5% CO₂ and 94% N²at 37°C. To do so, the day before, the cells were acclimatized to an environment with a reliable glucose content closer to physiological conditions. On the day of experiment, cells were placed in the hypoxic chamber with Phosphate Buffered Saline containing Ca²and Mg²⁺(Corning,#21-040-CV) previously balanced for ischemic condition, or with Dulbecco’s Modified Eagle Medium (DMEM, Thermofisher Scientific, #11885084) with Low glucose in an incubator for sham condition. After 5 hours of simulated ischemia, cells were collected for further experiments. To simulate reperfusion following the 5 hours of simulated ischemia, the PBS was replaced for all condition with fresh DMEM with Low Glucose during 24-48 hours before cells collection. The 5-hour duration of ischemia was based on the maximum ischemia time of acute myocardial infarction patients included in the Hibiscus cohort and the protocol used with this primary cell culture.

Transfection

A model of human cardiac fibroblast that under-expresses P2X4 was generated with P2X4R-targeted small interfering RNA (siRNA). Transfection was performed using TransIT-X2 (Mirus, Madison, USA, #Mir6004) and P2X4 siRNA (Santa Cruz Biotechnology, USA, #sc-42569) for 6 hours. Transfection media were replaced before subsequent experiments. For autophagic flux quantification, GFP-RFP-LC3 vector (addgene #21074) was used using TransIT-X2 (Mirus, Madison).

Cell viability

Annexin V / Propidium Iodide labelling was performed to assess cell viability according to manufacturer’s recommendations and cells were sorted by flow cytometry (BD FACS Melody Cell Sorter).

Electron microscopy

Electron microscopy was performed as previously described [23]. After experiments, cells were harvested with 0.025% trypsin and fixed for at least 24 h in 4% paraformaldehyde and 1% glutaraldehyde (Sigma, St-Louis, MO) in 0.1 M phosphate buffer (pH 7.2). Cells were washed in PBS and refixed with 2% osmium tetroxide (Agar Scientific, Stansted, UK) for 1 hour. They were then fully dehydrated in a series of ethanol and propylene oxide solutions. Finally, they underwent an impregnation step with a mixture (1∶1) of propylene oxide and Epon resin (Sigma), then left overnight in pure resin. Samples were then embedded in Epon (Sigma) resin, which polymerized for 48 h at 60°C. Ultra-fine sections (90 nm) of these blocks were obtained using a Leica EM UC7 ultramicrotome (Wetzlar, Germany). Sections were stained with 2% uranyl acetate (Agar Scientific), 5% lead citrate (Sigma). Images were obtained with a JEOL 1400 plus transmission electron microscope (JEOL, Tokyo, Japan).

Immunocytochemistry experiments

Cells were seeded on 18-mm sterile coverslips pre-coated with rat-tail collagen I in 12 well-plates for 24 hours. Then cells were placed in hypoxic chamber or not according to the protocol. Lysotracker was added according to the manufacturer protocol’s in normoxia conditions. Cells were then washed with PBS containing Ca²⁺ and Mg²and then fixed 4% paraformaldehyde for 15 minutes. After fixation, cells were washed two times with PBS containing Ca²⁺ and Mg²beforepermeabilization with 0.1% triton X-100. For avoiding unspecific antibody binding, cells were saturated with 3% BSA for 30 minutes. Primary antibodies for P2X4 (Alomone, APR-002, 1:200), Lamp-1 (Abcam, #ab25245, 1:800) were incubated overnight at 4°C. Cells were then washed with PBS containing Ca²⁺ and Mg²and incubated with Alexa Fluor 488- and Alexa Fluor 647-conjugated secondary antibodies (1:10 000) for 1h at room temperature. Coverslips were mounted on slides using ProLong® Gold Antifade Mountant with DAPI (Invitrogen, France). Images from immunocytochemistry and for autophagic flux with GFP-RFP-LC3 plasmid were obtained using epifluorescence microscope (EVOS M7000, Thermofisher, X60 objective). LC3 positive dots were quantified with ImageJ Software.

RNA extraction and RT-qPCR

Total RNA from fibroblasts was extracted with TRIzol (ThermoFischer Scientific #15596026) according to the manufacturer’s instructions (ThermoFischer Scientific, Waltham, MA, USA). Reverse transcription was performed using PrimeScript® RT Reagent kit (Takara, Shiga, Japan) and Flexlid thermocycler (Eppendorf, Hamburg, Germany) following Takara’s instructions. Then a semi-quantitative qPCR was done using TB green® Premix ExTaq (Takara) in a CFX Connect Real Time System (Bio-Rad) following Takara’s recommendations. P2X4 primers: F- 5′-TTACGACCAAGGTCAAGGGC-3’; R: 5′-CCTGTTGAGACTCCGTTGCT-3′. HPRT1: F- 5′-TTGCTGACCTGCTGGATTAC-3’; R: 5′-TATGTCCCCTGTTGACTGGT-3′.

Proteins extraction and western blot

Total protein extracts were obtained with chemical cell lysis using RIPA buffer including protease inhibitor cocktail (Sigma-Aldrich—P8340, Sigma-Aldrich -Merck, Saint Quentin Fallavier, France) and phosphatase inhibitors (Halt Phosphatase Inhibitor Cocktail—Thermo Fisher Scientific, Courtaboeuf, France). After the determination of protein concentration by BCA assay (Pierce™ BCA Protein Assay Kit, Thermo Fisher Scientific, Courtaboeuf, France), 35 μg of total protein were loaded on SDS-PAGE. Proteins were transferred onto nitrocellulose membrane and incubated with the following primary antibodies: LC3B (Abcam, #ab51520, 1:2,000), p62 (Abcam, #ab109012,1:1,000), P2X4 (Cell Signaling Technology, mAb#70659 , 1:1000), α-SMA (Cell signaling Technology, mAb #19245 #, 1:1000), Collagen1α (mAb #91144 , 1:1000) and Hsc70 (Santa Cruz Biotechnology, #sc-7298, 1:5,000). Densitometry was evaluated using ImageJ software.

Measurement of calcium influx with the Fluo-4-AM dye

To assess P2X4 channel activity, Fluo-4-AM was loaded at 1 μM for 45 minutes at 37°C in the different conditions. Then the loading media were removed, and the cells were put in PSS (140 mM NaCl, 4mM KCl, 2mM CaCl2, 1mM MgCl2, 10mM Hepes, 11,1mM D-glucose) for the experiment. After recording the basal fluorescence, 10 μM of ATP were added alone or with a selective noncompetitive allosteric antagonist BX 430. Fluorescence was acquired with an Evos M7000 microscope (Thermofisher, France). For the analysis, a difference of maximal signal and the basal signal was assessed per cell. The signal was quantified for 10 cells per field for a total of 3 different fields per duplicate. At total, we performed 3 different experiments. Controls were performed in the presence of suramin to determine the weight of P2X4 over the other ATP-sensitive P2Y receptors on calcium influx.

Wound-healing assay for the assessment of fibroblast migration

Cells were placed at confluence the day before the experiment. Then, a scratch was made with a tip, and the cells were placed in the different conditions: normoxia or ischemia for 5 hours. At the end, an image was captured with an Evos Microscope 7000 and considered as t=0h. Subsequent photos were taken at the 12h, 18h and 24 hours of reperfusion. All conditions were performed in duplicate for n=3-4 independent experiments. Analysis was performed with Fiji^TM software, and a coverage rate was calculated with the area at each point rationalized to the first area.

In vivo model of acute myocardial infarction:

Animal models

Female and male wild-type and P2rx4^-/-C57BL/6J mice were used in this study. P2rx4^-/-mice were provided from Centre Broca Nouvelle Aquitaine. All animal experiments were validated by the local ethics committee (CEEA VdL) and have been performed in accordance with the European Ethics rules (Ref: 2022111517354243).

Coronary artery ligation

General anesthesia was conducted by intraperitoneal injection of a mixture of ketamine-xyzaline (80mg/kg-8mg/kg) following institution recommendations. After thoracotomy in the 5^th intercostal space, a temporary ligation of the left anterior descending (LAD) coronary artery was performed using a 7.0 Prolene suture thread for 30 minutes. At the end of the procedure, a loose knot was left over the artery, and the ends were buried in the thorax. After closure of the chest, positive pressure was maintained by the ventilator to expand the lung and expel air from the thorax, to limit the risk of postoperative pneumothorax. Bupivacaine was injected subcutaneously over the scar, and a rehydration injection of 0.5 ml saline was given subcutaneously. At 7^th day, the ligation knot was tied for terminal ligation of the LAD, and 4% Evans Blue was injected to stain the perfused, i.e. healthy tissue, leaving only the at-risk tissue unstained. The heart was then removed and placed in PBS to wash off the excess of Evans blue. According to the following protocol, left ventricles were used for histology or Western blot experiments. To assess protein level expression the left ventricle was separated into two zones: the infarcted zone located downstream the ligature point and the healthy zone located upstream.

Necrosis and at-risk area determination

After removing the atria and the right ventricle, the left ventricle was sliced with mouse heart matrices (AgnThos 5mm). Slices were incubated with 2,3,5-Triphenyltetrazolium Chloride (TTC) (Sigma Aldrich, France, T8877) for 5 minutes at 37°C and then fixed with 4% Paraformaldehyde for at least 24 hours. Healthy area was delimitated with Evans Blue, at-risk non-necrotic area reacted with TTC, and necrosis area appeared white. All areas were quantified with FIJI^TM software.

Histology

Left ventricles fixed with 4% Paraformaldehyde were embedded in paraffin and 4 µm sections were obtained. Slides were stained using hematoxylin phloxin and saffron or Sirius Red. Glass slides were digitized using Nanozoomer s360 Digital slide scanner (Hamamatsu Photonics, Shizuoka, Japan) to obtain whole slide images. The analysis was performed using NDP view2 Image viewing software (Hamamatsu Photonics, Shizuoka, Japan). For each ventricle, the pathological section was chosen to quantify the inflammatory and fibrosis area and was expressed as the percentage of pathological area of the total ventricular surface.

Statistics and reproducibility

Data were displayed as median ± range as indicated in the figure legends. Mann-Whitney rank-sum test was used for comparisons between two groups. Wilcoxon matched-pairs signed-rank test was used for comparisons between two groups. Two-way ANOVA, followed by post-hoc pairwise comparisons with Bonferroni correction to adjust for multiple testing was used for comparison >3 groups. Confidence intervals of 95% were used and significance was considered when the p-value was less than 0.05, using GraphPad Prism v.9.0.0 (GraphPad Software).. The number of times an experiment was repeated is indicated in the figure legends.

Abbreviations

α-SMA

α-Smooth Muscle Actin

AMI

Acute Myocardial infarction

AMPK

AMP-activated protein kinase

ATG5

Autophagy related protein 5

Baf A1

Bafilomycin A1

ColI-α

Collagen-Iα

DAMPs

Damage-associated molecules

eATP

Extracellular Adenosine 5’-TriPhosphate

mTORC1

Mammalian target of rapamycin complex 1

NF-kB

Nuclear factor kappa B

P2

Purinergic Receptors 2

p38

P38 mitogen-activated protein kinases

Pi

Propidium Iodide

ROS

Reactive oxygen species

Authors’ contributions

All authors contributed extensively to the work presented in this study. A.V.V.d.S performed cell culture, molecular and cellular biology experiments and assessed calcium measurement and scratch assay. J.S., A.A., T.G. and L.C. performed in vivo mouse experiments. M.O. performed bioinformatic analysis. A.H. performed transmission electron microscopy. E.M-S. performed IHC analysis. F.I. directed the research. F.I., S.Cha., S.R. designed the study. A.V.V.d.S., F.I., S.Cha., S.R. analyzed the data. A.V.V.d.S. wrote a first draft of the paper. F.I., S.Cha., S.R., T.G. and E.B-G. corrected the paper.

Data availability

The data that support the findings of this study are available from the corresponding author, F.I., upon reasonable request.

Declarations

Competing interests

The authors declare no competing interests.