Introduction

Aging is a predominant risk factor for cardiovascular disease (CVD) (Moturi et al., 2022). Cardiac aging is accompanied by low-grade chronic inflammation termed “inflammaging” (Cevenini et al., 2013), which is linked to gradual development of cardiac fibrosis and heart failure, whereby macrophages, cardiac fibroblasts, and endothelial cells play critical roles in age-associated cardiac remodeling and dysfunction (Abdellatif et al., 2023; Meschiari et al., 2017). Studies in experimental animal models and in humans provide evidence demonstrating that aging heart is more vulnerable to stressors such as ischemia/reperfusion injury and myocardial infarction and exhibits less efficient reparative capability after injury as compared to the heart of young individuals (Bujak et al., 2008; Mariani et al., 2000). Even in the heart of apparently healthy individuals of old age, chronic inflammation, cardiomyocyte senescence, cell apoptosis, interstitial/perivascular tissue fibrosis, endothelial dysfunction and endothelial-mesenchymal transition (EndMT), and cardiac dysfunction either with preserved or reduced ejection fraction rate are observed (Abdellatif et al., 2023; Ruiz-Meana et al., 2020). Despite the intensive investigation in the past, the underlying causative cellular and molecular mechanisms responsible for the cardiac aging phenotype and the susceptibility of aging heart to injurious stressors are not fully elucidated, yet. Moreover, most preclinical studies preferentially use relatively juvenile animal models which hardly recapitulate clinical scenario. It is therefore critical to investigate mechanisms of cardiac aging phenotype with a naturally advanced aging animal model.

Research in the past years suggests a wide range of functions of the enzyme arginase, including the cytosolic isoenzyme arginase-I (Arg-I) and the mitochondrial isoenzyme arginase-II (Arg-II) in organismal aging as well as age-related organ structural remodeling and functional changes (Caldwell et al., 2015; Li et al., 2022). These functions of arginase have been implicated in cardiac and vascular aging (Xiong, Yepuri, Montani, et al., 2017; Yepuri et al., 2012) and cardiac ischemia/reperfusion injury in rodents (Jung et al., 2010). Arg-I, originally found in hepatocytes, plays a crucial role in urea cycle in the liver to remove ammonia from the blood (Sin et al., 2015), whereas Arg-II is inducible in many cell types under pathological conditions, contributing to chronic tissue inflammation and remodeling (Moretto et al., 2019). Both enzymes are able to catalyze the divalent cation-dependent hydrolysis of L-arginine (Ming et al., 2013). Studies in the literature including our own, demonstrate that Arg-II is involved in aging process, and promotes organ inflammation and fibrosis and inhibition of arginase or genetic ablation of arg-ii slows down aging process and protects against age-associated organ degeneration such as lung (Zhu et al., 2023), pancreas (Xiong, Yepuri, Necetin, et al., 2017) and heart (Xiong, Yepuri, Montani, et al., 2017). However, controversial results on expression of arginase and its isoforms at the cellular levels in the heart are reported. While some studies linked Arg-I to the cardiac ischemia/reperfusion injury (Jung et al., 2010), others suggest a role of Arg-II in this context (Heusch et al., 2010). Even protective functions of Arg-II are suggested in some studies, based on correlation of Arg-II levels with cardiac injury or systemic administration of Arg-II in rat models (Huang et al., 2018; Lu et al., 2012). Importantly, cellular localization of arginase and its isoenzyme in the heart, i.e., expression of arginase in cardiomyocytes and non-cardiomyocytes have not been systematically analyzed and confirmed. Finally, whether and how Arg-II participates in cardiac aging and whether it is through cell-autonomous and/or paracrine effects are not known.

Therefore, our current study is aimed to investigate which cell types in the heart express arginase and what enzymatic isoforms. How arginase and this isoenzyme in these cells influence the cardiac aging process and enhance susceptibility of the aging heart to ischemia/reperfusion injury.

Results

1. Increased arg-ii expression in aging heart

In heart from young (3-4 months in age) and old (22-24 months in age) mice, an age-associated increase in arg-ii expression in both wt male and female mice are observed (Fig. 1A). This age-associated increase in arg-ii expression is more pronounced in females than in males (Fig. 1A). In contrast to arg-ii, arg-i mRNA expression is very low in the heart (with very high Ct values, data not shown), and no age-dependent changes of arg-i gene expression are observed in wt and in arg-ii^−/− mice of either gender (Suppl. Fig. 1A and Suppl. Fig 1B), suggesting a specific enhancement of Arg-II in aging heart. Further experiments are thus focused on Arg-II and female mice. We made effort to analyze Arg-II protein levels by immunoblotting in the heart tissues. However, immunoblotting is not sensitive enough to detect Arg-II protein in the heart tissue of either young or old animals. Therefore, further experiments are performed to detect Arg-II proteins at the cellular level with confocal microscopic immunofluorescence staining as described in later sections.

Fig. 1.

2. Arg-ii ablation reduces cardiac aging phenotype

Next we investigated the impact of arg-ii ablation on cardiac phenotype. Masson’s staining reveals an enhanced peri-vascular and interstitial fibrosis in wt old mice, which is mitigated in age-matched arg-ii^−/− animals (Fig. 1B, 1C). The age-associated increase in cardiac fibrosis and its inhibition by arg-ii^−/− are further confirmed by quantitative measurements of collagen content as assayed by determination of hydroxyproline levels in the heart tissues (Fig. 1D). The increased cardiac fibrosis in aging is however, associated with decreased mRNA levels of collagen-Iα (col-Iα) and collagen-IIIα (col-IIIα), the major isoforms of pre-collagen in the heart (Suppl. Fig. 2A and 2B), which is a well-known phenomenon in cardiac fibrotic remodeling (Besse et al., 1994; Horn et al., 2016). The results demonstrate that age-associated cardiac fibrosis and prevention in arg-ii^−/− mice is due to alterations of translational and/or post-translational regulations including collagen synthesis and/or degradation. Furthermore, an increased density of fibroblasts as characterized by positive staining of PDGF-Rα is observed in wt old mice, but not in age-matched arg-ii^−/− animals (Fig. 1E and Fig. 1F). Interestingly, an age-associated increase in p16^ink4 positive senescent cells in wt mouse heart is prevented in the arg-ii^−/− animals (Fig. 1G and 1H).

Fig. 2.

Moreover, a significant increase in macrophage marker f4/80 (Fig. 2A) and elevated gene expression of numerous pro-inflammatory cytokines such as mcp1, il-1β and tnf-α are observed in aging heart of wt mice (Fig. 2B to Fig. 2D). This age-associated increase in the inflammatory markers and cytokine gene expression (except tnf-α) are overall significantly reduced or prevented in arg-ii^−/− mice (Fig. 2A to Fig. 2D). In line with the gene expression of f4/80, immunofluorescence staining reveals an age-associated increase in the numbers of F4/80⁺ cells in the wt mouse heart, which is reduced in the age-matched arg-ii^−/− animals (Fig. 2E to 2G), demonstrating that arg-ii gene ablation reduces macrophage accumulation in the aging heart. Interestingly, resident macrophages as characterized by LYVE1⁺/F4-80⁺ cells (Fig. 2E and 2H) are predominant in the aging heart as compared to the infiltrated CCR2⁺/F4-80⁺ cells (Fig. 2F and 2I). The increase in both LYVE1⁺/F4-80⁺ and CCR2⁺/F4-80⁺ macrophages in aging heart is reduced in arg-ii^−/− mice (Fig. 2E, 2F , 2H, and 2I).

We also observed an age-associated increase in total numbers of apoptotic cells as demonstrated by TUNEL staining in the heart of wt mice, which is reduced in age-matched arg-ii^−/−animals (Fig. 3A and Fig. 3B). Further experiments with wheat germ agglutinin (WGA)-Alexa Fluor 488 (used to define cell boarders) (Fig. 3C) show a higher percentage of apoptotic cardiomyocytes (CM) than non-cardiomyocytes (NCM) in wt mice (Fig. 3D). This increased percentage of cardiomyocyte apoptosis is also prevented in age-matched arg-ii^−/−animals (Fig. 3D). The percentage of apoptotic non-cardiomyocytes in aged arg-ii^−/− mice tended to be reduced as compared to wt mice, but it does not reach statistical significance (Fig. 3D).

Fig. 3.

In addition, heart from old wt mice reveals elevated levels of SNAIL (the master regulator of End-MT process) and vimentin (mesenchymal marker), which is prevented in arg-ii^−/−mice (Fig. 4A to Fig. 4C). No changes in CD31 (endothelial marker) levels are observed among the four groups (Fig. 4A and Fig. 4D). Immunofluorescence staining reveals that the age-related up-regulation of vimentin is mostly due to expansion of fibroblast population in left ventricle as shown in Fig. 1E and Fig. 1F). Confocal co-immunofluorescence staining shows that vimentin could be found in CD31⁺ endothelial cells of blood vessels in the heart of old wt but rarely in age-matched arg-ii^−/− mice (Fig. 4E). The results suggest at least a partial End-MT occurring in cardiac aging that is prevented by arg-ii ablation. With aging, the heart weight to body weight ratio (HW/BW) increases to a similar extent between wt and arg-ii^−/− mice (Suppl. Fig. 3), suggesting no difference in age-related cardiac hypertrophy between wt and arg-ii^−/−mice.

Fig. 4.

3. Cellular localization of Arg-II in aging heart

Next, the cellular localization of Arg-II in aging heart are investigated. Since arg-ii mRNA levels are upregulated mostly in aged females, heart tissues from old female mice are used mainly for this purpose. To our surprise, confocal co-immunofluorescence staining does not show Arg-II in cardiomyocytes as evidenced by lack of Arg-II staining in troponin T⁺-cells, a specific cardiomyocyte marker (Fig. 5A). Arg-II is only found in non-cardiomyocytes (Fig. 5A). In line with this notion, Arg-II is not detectable by immunoblotting in isolated primary cardiomyocytes from old wt mice at baseline or after exposure to hypoxia (1 % O₂, 24 hours, Fig. 5B), a well know strong stimulus for Arg-II expression in many cell types (Liang et al., 2019; Liang et al., 2021; Ren et al., 2022; Zhu et al., 2023). In contrast to cardiomyocytes, cardiac fibroblasts isolated from old wt mice express Arg-II which is up-regulated by hypoxia (Fig. 5B). The absence of arg-ii mRNA in cardiomyocytes in contrast to fibroblasts isolated from wt old mouse hearts is also confirmed with qRT-PCR (Fig. 5C). The absence of Arg-II in cardiomyocytes are also demonstrated in the heart of other species such as rats in which no staining could be found in the cardiomyocytes (with or without infarction) but in non-cardiomyocytes i.e., cells between myocytes and endocardial cells (Suppl. Fig. 4B). The specificity of the antibody against Arg-II was confirmed in the old wt and arg-ii^−/− mouse kidneys in which Arg-II is constitutively expressed in the S3 proximal tubular cells (Suppl. Fig. 4A) as demonstrated by our previous study (Huang et al., 2016). Again, Arg-II staining could be only found in cells between cardiomyocytes in the old wt mouse but not in arg-ii^−/− animals (Suppl. Fig. 4C). The absence of Arg-II in cardiomyocytes are also confirmed in human heart tissue biopsies (please see the results in Fig. 5H). Moreover, Arg-II is detected in Mac-2⁺ macrophages (Fig. 5D), CD31⁺ endothelial cells (Fig. 5E), and PDGF-Rα⁺-fibroblasts (Fig. 5F), but not in α-SMA⁺ vascular smooth muscle cells (Fig. 5G). These results demonstrate that Arg-II in aging mouse heart is mostly expressed in non-cardiomyocytes such as macrophages, endothelial cells and fibroblasts. Similar to mouse heart, in human heart (obtained from a 66-year-old woman, and a 76-year-old man with no significant cardiac pathology), Arg-II is also absent in cardiomyocytes as proved by lack of co-localization with troponin-T⁺-cells (Fig. 5H), and is expressed in CD31⁺-endothelial cells (Fig. 5I), CD68⁺-macrophages (Fig. 5J), and vimentin⁺-(Fig. 5K) or α-SMA⁺-(Fig. 5L) fibroblasts/myofibroblasts. In contrast to mouse heart, Arg-II could be found in very few vascular smooth muscle cells in human heart (Fig. 5M).

Fig. 5.

4. Role of Arg-II in crosstalk between macrophages and cardiomyocytes through IL-1β

Given that age-associated cardiomyocyte apoptosis is prevented in arg-ii^−/− mice despite absence of Arg-II in the cardiomyocytes, the non-cell-autonomous effects of Arg-II on cardiomyocytes are then investigated. As reported above, macrophage numbers and il-1β mRNA expression are increased in aging heart and reduced in arg-ii^−/− mice (Fig. 2A, 2C, and 2G). Both immunoblotting and immunofluorescence staining confirm that IL-1β protein levels are higher in old wt heart tissues when compared to age-matched arg-ii^−/−(Fig. 6A to 6D). Furthermore, co-immunofluorescence staining reveals that IL-1β is localized in Mac2⁺-macrophages (Fig. 6E). Taking into account that our previous studies demonstrated a relationship of Arg-II and IL-1β in vascular disease and obesity (Ming et al., 2012) and in age-associated organ fibrosis such as renal and pulmonary fibrosis (Huang et al., 2021; Zhu et al., 2023), and IL-1β has been shown to play a causal role in patients with coronary atherosclerotic heart disease as shown by CANTOS trials (Ridker et al., 2017), we therefore focused on the role of IL-1β in crosstalk between macrophages and cardiac cells such as cardiomyocytes, fibroblasts and endothelial cells. Considering that ablation of arg-ii gene specifically reduces the number of TUNEL-positive cardiomyocytes (Fig. 3C and 3D), we hypothesized that Arg-II in macrophages may prime cardiomyocyte apoptosis. To test this hypothesis, splenic macrophages are isolated from young and old wt and age-matched arg-ii^−/− mice, and conditioned medium from the cells are collected. Elevated Arg-II and IL-1β protein levels are observed in old wt macrophages as compared to the cells from young animals (Fig. 7A to 7C). In line with this finding, IL-1β levels in conditioned medium from old wt mouse macrophages are higher as compared to the conditioned medium from young macrophages (Fig. 7D). Interestingly, arg-ii ablation substantially reduces IL-1β levels in the macrophages (Fig. 7A, 7C, and 7D). Importantly, cardiomyocytes isolated from adult (five-months) female wt mice as bioassay cells, when incubated with conditioned medium from old (but not from young) wt macrophages for 24 hours (Fig. 7E), display more apoptosis as monitored by TUNEL assay (Fig. 7F and 7G). This age-associated cardiac damaging effect of macrophages is not observed with arg-ii^−/− macrophage conditioned medium (Fig. 7F and 7G), suggesting a role of Arg-II expressing macrophages in mediating cardiomyocyte apoptosis in aging. To further examine whether IL-1β from macrophages is the paracrine mediator for cardiomyocyte apoptosis, the above described experiment is performed in presence of the IL-1 receptor antagonist IL-Ra. Indeed, cardiomyocyte apoptosis induced by the conditioned medium from old wt macrophages is abolished by IL-Ra (Fig. 7F and 7G).

Fig. 6.

Fig. 7.

Similar results are obtained with Raw264.7 cells, a widely used mouse macrophage model. Raw264.7 cells stimulated with lipopolysaccharide (LPS; 100 ng/mL) for 24 hours, show a significant increase in both Arg-II and IL-1β, which is inhibited by silencing arg-ii (Suppl. Fig. 5A to Suppl. Fig. 5C), confirming the role of Arg-II in the production of IL-1β in macrophages (Suppl. Fig. 5A and Suppl. Fig. 5C). In agreement with the results shown in Fig. 7, conditioned medium from LPS-treated Raw264.7 cells significantly enhances cardiomyocyte apoptosis as assessed by TUNEL staining, which is reduced by either arg-ii gene silencing or by IL-1 receptor blocker ILRa (Suppl. Fig. 5D and Suppl. Fig. 5E). It is of note that that LPS alone does not induce cardiomyocyte apoptosis at the concentration tested (data not shown), confirming the role of Arg-II-mediated IL-1β release from macrophages in cardiomyocyte apoptosis.

Next, we analyzed relationship between Arg-II and iNOS in regulation of IL-1β production in macrophages. In the human THP1 monocytes in which Arg-II but not iNOS is induced by LPS (100 ng/mL for 24 hours) (Suppl. Fig. 6A), mRNA and protein levels of IL-1β are markedly reduced in arg-ii knockout THP1^arg-ii-/- as compared to the THP1^wt cells (Suppl. Fig. 6B and 6C), further confirming that Arg-II promotes IL-1β production as already shown in RAW264.7 macrophages (Suppl. Fig. 5A and 5C). Moreover, in the mouse bone-marrow-derived macrophages, LPS-induced IL-1β production is inhibited by inos deficiency (BMDM^inos-/- vs BMDM^wt) (Suppl. Fig. 6D and 6E), while Arg-II levels are slightly enhanced in the BMDM^inos-/- cells (Suppl. Fig. 6D and 6F). All together, these results suggest that Arg-II and iNOS are upregulated by LPS independently and iNOS slightly reduces Arg-II expression. Both Arg-II and iNOS are required for maximal IL-1β production upon LPS stimulation as illustrated in Suppl. Fig. 6G.

5. Role of Arg-II in crosstalk between macrophages and cardiac fibroblasts through IL-1β

Since cardiac fibrosis is accompanied with macrophage accumulation in our aging heart model, a crosstalk between macrophages and fibroblasts is then investigated. Cardiac fibroblasts isolated from adult (five-months) female wt mice as bioassay cells, produce more hydroxyproline when they are incubated with conditioned medium from old as compared to young wt macrophages (Suppl. Fig. 7A). This effect of old macrophages on fibroblasts is reduced with conditioned medium from arg-ii^−/− macrophages (Fig. 8A). In parallel, expression levels of genes involved in extracellular matrix remodeling such as fibronectin, collagen IIIα, tgf-β1 (Fig. 8B to Fig. 8D) as well as mmp2 and mmp9 (Suppl. Fig. 7B and Suppl. Fig. 7C) are reduced in fibroblasts stimulated with conditioned medium from arg-ii^−/−macrophages as compared to the old wt macrophages, while expression of collagen Iα is not altered under this condition (Suppl. Fig. 7D). Importantly, the increase in hydroxyproline levels in cardiac fibroblasts incubated with conditioned medium from old wt macrophages is inhibited by ILRa (Fig. 8E).

Fig. 8.

6. Cell-autonomous effects of Arg-II in cardiac fibroblasts: role of mtROS

Since Arg-II is expressed in fibroblasts, a cell-autonomous effect of Arg-II in these cells is also investigated in cultured human cardiac fibroblasts (HCF). Overexpression of arg-ii in the cells (Fig. 8F and 8G) increases vimentin and collagen IIIα expression levels (Fig. 8H and 8I), demonstrating a cell-autonomous effect of Arg-II in cardiac fibroblasts. Interestingly, upregulation of collagen IIIα expression by arg-ii is accompanied with increased mitochondrial ROS (mtROS) generation, and inhibition of mtROS by mito-TEMPO (10 μmol/L) prevents col-IIIa upregulation (Fig. 8J , 8K and 8L), suggesting a role of Arg-II-induced mtROS in activation of fibroblasts. The effect of Arg-II in promoting mtROS is also confirmed under hypoxic conditions in which endogenous Arg-II is upregulated accompanied with elevated HIF-1α levels (Fig. 8M and 8O). The enhanced mtROS generation under hypoxic conditions are inhibited either by mito-Tempo (Fig. 8N), the specific inhibitor of mtROS or by arg-ii silencing (Fig. 8O and 8P).

7. Role of Arg-II in crosstalk between macrophages and endothelial cells through IL-1β

Results in Fig. 4 suggests EndMT in aging heart, which is regulated by Arg-II. The role of Arg-II-expressing macrophages in End-MT of endothelial cells is then investigated in cultured cells. Human endothelial cells exposed to macrophage-derived conditioned medium from old (not young) wt mice for 96 hours has no effect on VE-cadherin but enhances N-cadherin and vimentin (mesenchymal markers), and also Arg-II protein levels (Fig. 9A to Fig. 9E), suggesting a partial End-MT induced by aged macrophages. Interestingly, these effects of macrophages are prevented in old arg-ii^−/− mice (Fig. 9A to Fig. 9E) or abolished in the presence of IL-1 receptor antagonist IL-Ra (Fig. 9F to Fig. 9J). The results suggest a role of IL-1β from old wt macrophages in EndMT process.

Fig. 9.

8. Arg-II gene ablation reduces ischemic damage in aging heart

To study the specific role of Arg-II in heart ischemia/reperfusion (I/R) injury during aging, ex vivo experiments with Langendorff-perfused heart from old wt and arg-ii^−/− mice were performed followed by cardiac functional analysis. The experimental protocol of global ischemia/reperfusion is shown in Fig. 10A. Under normoxic baseline condition, the cardiac functions, i.e., dP/dt_max. (maximal rate of contraction), dP/dt_min. (maximal rate of relaxation) and left ventricular developed pressure (LVDP) are comparable between wt and arg-ii^−/− old mice (Suppl. table 1). Arg-ii ablation improves cardiac function following I/R injury, i.e., the inotropic contractile function as measured by LVDP and dP/dt_max, lusitropic dP/dt_min (Fig. 10B). The post I/R recovery is significantly improved in arg-ii^−/− mice when compared to age-matched wt animals (Fig. 10B). In addition to functional recovery, infarct size after I/R-injury was analyzed by triphenyl tetrazolium chloride (TTC) staining in old mice. Under the ischemic/reperfusion condition (20 minutes ischemia and 30 minutes reperfusion), no myocardial infarct was detected (Suppl. Fig. 8), while longer ischemia and reperfusion time (30 minutes and 45 minutes respectively) is required to induce myocardial infarct, i.e., 10.5 ± 2.9% in wt old mice, which is significantly reduced in the age-matched arg-ii^−/−mice (3.2 ± 0.9%, Fig. 10C and Fig. 10D). It is to note that following ischemia/reperfusion injury, male mice display cardiac dysfunction, i.e., reduced left ventricular developed pressure (LVDP), as well as the inotropic and lusitropic states (expressed as dP/dt max and dP/dt min, respectively) (Suppl. Fig. 9). As previously reported (Murphy et al., 2007), we also found that old male mice are more prone to I/R injury than age-matched female animals. Specifically, 15 minutes of ischemic time is enough to significantly affect the left ventricle contractile function in the male mice (Suppl. Fig. 9). As opposite, age-matched old female mice are relatively resistant to I/R injury, and at least 20 min of ischemia are necessary to induce a significant impairment of the contractile function (Fig. 10). Similar to females, the post I/R recovery of cardiac function is also significantly improved in the male arg-ii^−/− mice as compared to age-matched wt animals. In addition to functional recovery, triphenyl tetrazolium chloride (TTC) staining (myocardial infarction) upon I/R-injury in males is significantly reduced in the age-matched male arg-ii^−/− animals (Suppl. Fig. 9C and 9D). All together, these results reveal a role for Arg-II in heart function impairment during aging in both genders with a higher vulnerability in males (Suppl. Fig. 9).

Fig. 10.

Discussion

Previous studies have reported the presence of both Arg-I and Arg-II isoenzymes in the cardiac tissues of various animal models including feline, pig, rats, rabbit, and mice (Gonon et al., 2012; Heusch et al., 2010; Jung et al., 2006; Jung et al., 2010; Steppan et al., 2006). These isoenzymes have been implicated in cardiac ischemic injury and cardiac aging (Jung et al., 2010; Xiong, Yepuri, Montani, et al., 2017). It has been widely assumed that Arg-I and Arg-II are expressed in cardiomyocytes, where they play a role in cardiac injury via a cell-autonomous effect on the cardiomyocytes. However, the cellular localization of arginase isoforms has not been convincingly demonstrated due to the following two reasons: species-specific expression of arginase isoforms and lack of including negative controls with gene knockout samples.

Our current study systematically investigated the expression and cellular localization of the mitochondrial Arg-II in the heart of a natural aging mouse model and in human biopsies. We found an age-associated increase in arg-ii but not arg-i in heart of male and female mice. Data extracted from a high throughput sequencing database of male mice (no data on females are available, unfortunately) confirms our findings showing the age-dependent enhancement of arg-ii but not arg-i expression in the heart (GSE201207) (Wolff et al., 2023) (Suppl. Fig. 1C and 1D). Surprisingly, we found that neither arg-ii gene expression nor its protein levels were detectable in cardiomyocytes from either young or old mice. Despite increased arg-ii mRNA (but not arg-i mRNA) expression in the aging heart, mitochondrial Arg-II was absent from cardiomyocytes. This observation aligns with previous findings in vascular endothelial cells from mice and humans, where Arg-II (not Arg-I) is the predominant isoenzyme (Ming et al., 2004). It is to note that a previous study reported that Arg-II is exclusively expressed in isolated cardiomyocytes from in rats (Steppan et al., 2006). However, we could not confirm this finding. The reason of this discrepancy is not clear. However, we made great effort to confirm the cellular localization of Arg-II in the heart from different species including mice, rats and humans. Importantly negative controls. i.e., arg-ii^−/− samples are always included in our present study avoid any possible background signals. Our experiments confirm that Arg-II could be found only in non-cardiomyocytes but not in cardiomyocytes from different species. The experiments with freshly isolated primary cells also confirm that even under hypoxia, a well know strong stimulus for Arg-II protein levels in different cell types (Liang et al., 2019; Zhu et al., 2023), no Arg-II protein expression could be detected in the cardiomyocytes but elevated in fibroblasts from old female mice. Furthermore, RT-qPCR could not detect arg-ii mRNA in cardiomyocytes but in non-cardiomyocytes. All together, these results demonstrate that Arg-II are not expressed or at negligible levels in cardiomyocytes but expressed in non-cardiomyocytes. Further experiments identified that Arg-II is expressed in macrophages, fibroblasts, and endothelial cells in aged mouse heart and human cardiac biopsies. These results suggest that Arg-II exerts non-cell-autonomous effects on aging cardiomyocytes.

It is well known that the aging heart is associated with accumulation of senescent cells, increased cardiomyocyte apoptosis, chronic inflammation, oxidative stress, and cardiac fibrosis (Vakka et al., 2023). In the current study, we observe that arg-ii^−/− mice are protected from cardiac aging phenotype and reveal a decrease in apoptotic cells, particularly cardiomyocytes but also non-cardiomyocytes. Since cardiomyocytes do not express Arg-II in aging and Arg-II could not even be induced in these cells under hypoxic condition, a non-cell-autonomous effect of Arg-II on cardiomyocyte apoptosis must be involved, meaning that the increased cardiomyocyte apoptosis in aging must be due to paracrine effects of other cell types such as macrophages in which Arg-II levels are elevated in aging. It is evident that crosstalk between cardiomyocytes and non-cardiomyocytes are critical in cardiac functional changes and remodeling (Hulsmans et al., 2016). Substantial evidence demonstrates that immune cells, particularly, monocytes and/or macrophages, are important contributors to cardiac remodeling in aging and disease conditions (Hulsmans et al., 2016; Jimenez et al., 2022; Wang et al., 2020). Under acute and chronic ischemic heart disease conditions, blood monocytes can be mobilized from bone marrow and spleen, and recruited into the injury site of the heart where they play an important role in cardiac remodeling (Honold et al., 2018; Swirski et al., 2009). An increased accumulation of myeloid cells including in aging heart has been reported and contributes to cardiac inflammaging (Esfahani et al., 2021). Besides infiltrated macrophages, tissue resident macrophages also contribute to cardiac inflammation and remodeling in heart disease (Suku et al., 2022; Weinberger et al., 2024). Interestingly, we demonstrate that total numbers of macrophages in the heart are significantly increased in aging heart, which is due to both enhanced accumulation of resident tissue macrophages and infiltrated macrophages and both macrophage populations are reduced in arg-ii^−/− mice. The enhanced infiltrated macrophages must be due to sustained cardiac injury occurring with aging in the heart accompanied with repairing process which is manifested by resident macrophage proliferation and fibroblast activation (Weinberger et al., 2024). Our previous study demonstrated that Arg-II plays a role in macrophage infiltration in different organ/tissues in high-fat diet and high-cholesterol diet-induced atherosclerosis and obesity mouse models (Ming et al., 2012). It remains to be investigated whether Arg-II regulates resident macrophage proliferation in the aging heart and what are the underlying mechanisms. Whether infiltrated macrophages can be switched to resident ones and regulated by Arg-II is also an interesting question to be investigated.

As shown above an increased macrophage accumulation in aging heart associates with elevated expression of numerous inflammatory cytokines in macrophages. This age-associated cardiac inflammation is inhibited in arg-ii^−/− animals. Among these inflammatory cytokines, an increase in IL-1β levels in cardiac tissue and macrophages or monocytes/macrophages derived from spleen of old mice are demonstrated accompanied with elevated Arg-II levels as compared to young animals. Importantly, we showed that Arg-II in the aged macrophages enhances IL-1β expression and release mediating cardiomyocyte apoptosis, since the cardiomyocyte apoptosis caused by the aged macrophages is inhibited in arg-ii^−/−mice and prevented by IL-1β receptor antagonist. This results further confirm our previous findings demonstrating a pro-inflammatory role of Arg-II in macrophages in vascular atherosclerosis and high-fat-diet-induced type 2 diabetes mouse models (Ming et al., 2012). This conclusion is also further supported by the findings with a mouse macrophage cell line RAW264.7. The question what are the underlying mechanisms that upregulate Arg-II levels in macrophages in aging remains to be investigated. Moreover, we also found increased apoptosis of other cell types in aging heart, which is also prevented in arg-ii^−/−mice. What are these cells, how this cell apoptosis is triggered, and what is the impact on age-associated cardiac vulnerability to stressors require further investigation. Moreover, research has provided evidence that cellular senescence including cardiomyocytes, endothelial cells, fibroblasts and immune cells all contributes to the cardiac aging and age-associated cardiac dysfunction (Luan et al., 2024). Future research shall investigate which cell types are protected from arg-ii deficiency from senescence and what are the underlying mechanisms.

One of the interesting questions is the relationship between Arg-II and iNOS in regulation of IL-1β production in macrophages. Both iNOS and Arg-II (not Arg-I) are highly induced in macrophages in response to several pro-inflammatory stimuli such as LPS, TNFα, and IFN-γ, etc. (Bogdan, 2015; Ming et al., 2012) and considered as pro-inflammatory M1-macrophage markers (Martinez et al., 2014; Ming et al., 2012). Since arginase and iNOS share the same metabolic substrate L-arginine, an increase in arginase levels is proposed to decrease intracellular L-arginine bioavailability for iNOS, and in turn to reduce pro-inflammatory responses of macrophages (Chang et al., 1998). Hence, arginase including the two isoforms Arg-I and Arg-II is regarded as anti-inflammatory (Yang et al., 2014). Arg-I is indeed associated with M2-anti-inflammatory macrophages (Yang et al., 2014), while Arg-II (but not Arg-I) along with iNOS is enhanced in pro-inflammatory macrophages upon LPS stimulation as shown in the previous study (Ming et al., 2012) and also confirmed in our present study. Importantly, knockout or knockdown of arg-ii gene reduces IL-1β production as shown in our present study and decreases pro-inflammatory cytokine levels in vitro and in vivo in various chronic inflammatory disease models and inflammaging (Atawia et al., 2019; Ming et al., 2012; You et al., 2013). These results demonstrate that Arg-II, in contrast to Arg-I, exerts pro-inflammatory functions in macrophages. Despite controversies published in the literature (Bogdan, 2015), iNOS is generally considered to exert pro-inflammatory effects in macrophages (Eslami et al., 2017; Yao et al., 2022). This is confirmed by our present study showing that LPS-stimulated IL-1β production in murine macrophages is inhibited when inos is ablated. Since Arg-II promotes IL-1β production and Arg-II levels are increased under inos-deficiency condition, one would expect an increased IL-1β production in inos^−/− macrophages. This is however not the case. A strong inhibition of IL-1β production in inos^−/− macrophages is observed. These results implicate that iNOS promotes IL-1β production independently of Arg-II and the inhibiting effect of IL-1β by inos deficiency is dominant and able to counteract Arg-II’s stimulating effect on IL-1β production. On the other hand, the question whether Arg-II promotes IL-1β production through iNOS could also be excluded due to the following reasons. First, in the human THP1 cells in which Arg-II but not iNOS is induced by LPS, ablation of arg-ii gene reduces IL-1β mRNA and protein levels; second, both Arg-I and Arg-II is believed to inhibits endothelial NOS-mediated NO synthesis, arg-ii deficiency would increase L-arginine bioavailability for NOS activity (Durante et al., 2007). Since iNOS induction by LPS is promoting IL-1β production as demonstrated in our current study, an increase in IL-1β production by arg-ii deficiency would be expected under this condition. We observe, however, a decreased IL-1β production in arg-ii^−/− cells. Hence, our results demonstrate that Arg-II promotes IL-1β production in macrophages independently of iNOS (This concept is illustrated in the Suppl. Fig. 6G). Future study shall further confirm the iNOS-independent stimulating effect on IL-1β production by Arg-II in the inos^−/− mouse model. It remains to be investigated what are the exact molecular mechanisms of IL-1β production promoted by Arg-II and iNOS in macrophages, respectively. Since IL-1β is produced and released via activation of inflammasome (Agostini et al., 2004; Cullen et al., 2015), whether Arg-II and iNOS promotes macrophage pro-inflammatory responses and IL-1β release through inflammasome activation remains to be studied.

Previous studies suggest that Arg-II exerts both L-arginine metabolizing-dependent and - independent (pleiotropic) effects depending on cell types. For example, in vascular smooth muscle cells in which no NOS existing, even inactive mutant of Arg-II is capable of inducing cell senescence (Xiong et al., 2013), while in endothelial cells, eNOS-uncoupling, i.e., decreased NO and increased superoxide anion production, is dependent on the L-arginine metabolizing activity of Arg-II (Yepuri et al., 2012). It is understandable that endothelial dysfunction caused by enhanced Arg-II levels would reduce coronary perfusion and in turn decrease cardiac contractile function as reported in the literature (Khan et al., 2012; Luo et al., 2014). Whether Arg-II promotes inflammatory responses in macrophages through the pleiotropic effects remains to be investigated.

It is of note that a study reported that Arg-II is required for IL-10 mediated-inhibition of IL-1β in mouse BMDM upon LPS stimulation (Dowling et al., 2021), which suggests an anti-inflammatory function of Arg-II. The results of our study, however, demonstrate that a pro-inflammatory effect of Arg-II in macrophages. Our findings are supported by the study from another group, which shows decreased pro-inflammatory cytokine production including IL-6 and IL-1β in arg-ii^−/−BMDM most likely through suppression of NFκB pathway, since arg-ii^−/− BMDM reveals decreased activation of NFκB and IL-1β levels upon LPS stimulation (Uchida et al., 2023). Most importantly, our previous study also showed that re-introducing arg-ii gene back to the arg-ii^−/−macrophages markedly enhances LPS-stimulated pro-inflammatory cytokine production (Ming et al., 2012), providing further evidence for a pro-inflammatory role of arg-ii under LPS stimulation. In support of this conclusion, chronic inflammatory diseases such as atherosclerosis and type 2 diabetes (Ming et al., 2012), inflammaging in lung (Zhu et al., 2023), kidney (Huang et al., 2021) and pancreas (Xiong, Yepuri, Necetin, et al., 2017) of aged animals or acute organ injury such as acute ischemic/reperfusion or cisplatin-induced renal injury are reduced in the arg-ii^−/− mice (Uchida et al., 2023). The discrepant findings between these studies and that with IL-10 may implicate dichotomous functions of Arg-II in macrophages, depending on the experimental context or conditions. Nevertheless, our results strongly implicate a pro-inflammatory role of Arg-II in macrophages in the inflammaging in aging heart.

One of the important features of cardiac aging is fibrosis which is due to chronic inflammation process (Lu et al., 2017). There are convincing evidences demonstrating that chronic inflammation facilitates development of myocardial fibrosis in heart failure which is associated with aging (Lillo et al., 2023). A crosstalk between macrophages and fibroblasts has been demonstrated to be important for cardiac fibrosis in diseased conditions (Hartupee et al., 2016). Studies show that monocyte-derived macrophages from bone marrow and spleen are recruited in advanced age, contributing to cardiac fibrosis and myocardial dysfunction (Hulsmans et al., 2018). In our present study using an in vitro cellular model, we show a pivotal role of Arg-II-expressing macrophages in activation of cardiac fibroblasts in aging heart. First, Arg-II protein is present in the cardiac macrophages and enhanced in monocyte-derived macrophages in aging mice; second, conditioned medium from aged arg-ii^−/− macrophages exhibits decreased potential of stimulating cardiac fibroblasts to produce components that are essentially involved in fibrosis such as hydroxyproline, fibronectin, coll-IIIa, and tgfb1. Interestingly, the enhanced hydroxyproline levels stimulated by old wt mouse macrophage-derived conditioned medium is inhibited by the IL-1β receptor blocker, demonstrating a paracrine effect of IL-1β release from aged macrophages on cardiac fibroblast activation. Moreover, the age-associated cardiac fibrosis and increase in fibroblast number in the heart is prevented in arg-ii^−/− mice, demonstrating a crosstalk between macrophage and fibroblasts in cardiac aging.

The fact that cardiac fibroblasts in aging heart express elevated Arg-II, indicates a contribution of a cell-autonomous effect of Arg-II in this cell type to cardiac fibrosis in aging. This concept is confirmed by the experiments showing that overexpression of arg-ii in cardiac fibroblasts enhances vimentin levels associated with increased expression of col-IIIa and mitochondrial ROS generation. Importantly, inhibition of mitochondrial ROS by TEMPO abolished the effect of arg-ii overexpression in the fibroblasts, demonstrating that Arg-II exerts its cell-autonomous stimulating effects in fibroblasts through mitochondrial ROS. The role of Arg-II in mitochondrial ROS generation is further confirmed by the results showing that silencing arg-ii abolished miROS generation in fibroblasts when Arg-II is upregulated under hypoxic condition. These findings are supported by the studies showing that ROS favors cardiac fibroblast proliferation and activation to produce collagen (Alili et al., 2014; Ohtsu et al., 2005). With these results, we demonstrate that cardiac fibroblasts are activated in aging by a paracrine effect of macrophages, that is mediated by Arg-II / IL-1β and also by a cell-autonomous effect of Arg-II through mtROS.

Endothelial cells play a key role in regulation of blood flow to organs through eNOS pathway, one of the most important regulatory mechanisms in cardiovascular system (Godo et al., 2017). Previous studies demonstrated a causal role of Arg-II in promotion of vascular endothelial aging through eNOS uncoupling (Yepuri et al., 2012). In line with this, our current study showed that Arg-II is expressed in the coronary vascular endothelium of old mice and also in human heart. Emerging evidence also suggests that endothelial cells have a significant capacity for plasticity and can change their phenotype, including endothelial-to-mesenchymal transition (EndMT), a process that endothelial cells undergoing extensive alterations, including changes in gene and protein expression, loss of the endothelial phenotype, and switch to mesenchymal phenotype such as increased cell migration, proliferation, and increased collagen production, contributing to cardiovascular disease and fibrosis (Murdoch et al., 2014; Zeisberg et al., 2007; Zhang et al., 2021). Although EndMT in cardiac fibrotic models, particularly in pro-atherosclerotic mouse models are at least partially evidenced, it is less known in cardiovascular aging (Evrard et al., 2016; Kidder et al., 2023; Xu et al., 2023). When compared to young animals, aged wt heart tissues show enhanced levels of SNAIL, the master regulator of End-MT process. Moreover, immunofluorescence reveals that CD31⁺ endothelial cells also co-express vimentin (mesenchymal marker). No changes of endothelial marker expression such as CD31 and vascular-endothelial cadherin (data not shown) are appreciated, suggesting that partial End-MT occurs during aging. Ablation of arg-ii blunts the age-related elevation of both vimentin and SNAIL, thereby reducing the number of CD31⁺/vimentin⁺ endothelial cells. This data hints that lower cardiac tissue fibrosis in arg-ii^−/− is at least partially contributed by dampened End-MT process. Indeed, conditioned medium from old wt macrophages is capable of enhancing mesenchymal markers N-cadherin and vimentin, as well as Arg-II protein levels in endothelial cells without changes in endothelial markers such as VE-cadherin, confirming a partial End-MT induced by aged macrophages. Interestingly, this effect of macrophages is prevented when arg-ii was ablated. Furthermore, the paracrine effects of the old macrophages on EndMT are prevented by IL-1 receptor antagonist ILRa, demonstrating a role of IL1β from old wt macrophages expressing Arg-II in EndMT process. Moreover, Arg-II is expressed and upregulated in endothelial cells with aging (Yepuri et al., 2012). Overexpression of arg-ii in the endothelial cells did not affect the End-MT markers (data not shown), suggesting that Arg-II alone is not sufficient to induce a cell-autonomous effect in End-MT. Finally, we demonstrate that arg-ii^−/− mice in aging exhibit improved cardiac function and are more resistant to ischemic/reperfusion stress and reveal a better recovery and less myocardial infarct area as compared to the old wt animals. This protection against age-related cardiac dysfunction and vulnerability to ischemic stress in arg-ii^−/− animals could be explained by the above described cardiac aging phenotype such as increase in apoptosis of cardiac cells, decrease in macrophage-mediated inflammation, endothelial dysfunction and EndMT, and fibroblast activation.

Previous studies provided sex-related difference in Arg-II levels at both mRNA (Xiong, Yepuri, Montani, et al., 2017) and protein levels in various organs (Huang et al., 2021; Xiong, Yepuri, Necetin, et al., 2017). We further confirmed the observation that in female mice, there is a greater age-associated upregulation of arg-ii levels in the heart, which is specifically found in non-cardiomyocytes but not in cardiomyocytes. A possible explanation for this sex-specific phenomenon may lie in differences in hormonal patterns between male and female animals throughout the aging process. Indeed, sex hormones, including estrogen and testosterone, have been shown to regulate arg-ii expression in various tissues. Estrogen seems to downregulate Arg-II (Hayashi et al., 2006). The age-related greater increase in arg-ii in the female mice might be due to loss or lack of estrogen in old female animals, which requires further investigation. However, testosterone has been reported to upregulate arg-ii expression (Levillain et al., 2005). This observation, however, does not seem to support the hypothesis that increased arg-ii expression in aged male mouse heart is related to sex hormones. Nevertheless, how age enhances Arg-II in sex-specific manner remains a topic of further investigation. Of note, previous studies showed that sex-related difference in pathological phenotypes in the aging kidney, pancreas, and lung are associated with higher levels of Arg-II in the females (Huang et al., 2021; Xiong, Yepuri, Necetin, et al., 2017; Zhu et al., 2023). The fact that aged females have higher Arg-II but are more resistant to I/R injury seems contradictory to the detrimental effect of Arg-II in I/R injury. It is presumable that cardiac vulnerability to injuries stressors depends on multiple factors/mechanisms in aging. Other factors/mechanisms associated with sex may prevail and determine the higher sensitivity of male heart to I/R injury, which requires further investigation. Nevertheless, the results of our study show that Arg-II plays a role in cardiac I/R injury also in males.

In conclusion, our study demonstrates that age-related upregulation of Arg-II in macrophage, fibroblasts, and endothelial cells contributes to the cardiac aging phenotype. This includes manifestations, such as cardiac inflammatory response, partial EndMT, fibroblast activation, resulting in cardiomyocyte apoptosis, cardiac fibrosis, and ultimately, myocardial dysfunction. Moreover, it enhances the vulnerability of aging heart to ischemic/reperfusion injury, primarily through a non-cell-autonomous paracrine release of IL-1β from macrophages acting on cardiomyocytes and endothelial cells. This effect is in addition to a cell-autonomous effect of Arg-II in cardiac fibroblasts (Fig. 11). Future research shall develop the macrophage-specific arg-ii^−/− mouse model to confirm this conclusion with aging animals. Since Arg-II is also expressed in fibroblasts and endothelial cells and exerts cell-autonomous and paracrine functions, aging mouse models with conditional arg-ii knockout in the specific cell types would be the next step to elucidate cell-specific function of Arg-II in cardiac aging. In this context, another interesting aspect is the cross-talk between macrophages and vascular SMC in the aging heart. In our present study, we could not detect Arg-II in vascular SMC of mouse heart but in that of human heart. This could be due to the difference in species-specific Arg-II expression in the heart or related to the disease conditions in human heart which is harvested from patients with cardiovascular diseases. Indeed, in the apoe^−/− mouse atherosclerosis model, aortic SMCs do express Arg-II (Xiong et al., 2013). It is interesting to note that rodents hardly develop atherosclerosis as compared to humans. Whether this could be partly contributed by the different expression of Arg-II in vascular SMC between rodents and humans requires further investigation. In our present study, the aspect of the cross-talk between macrophages and vascular SMC is not studied. Since the crosstalk between macrophages and vascular SMC has been implicated in the context of atherogenesis as reviewed (Gong et al., 2025), further work shall investigate whether Arg-II expressing macrophages could interact with vascular SMC in the coronary arteries in the heart and contribute to the development of coronary artery disease and/or vascular remodelling and the underlying mechanisms. Nevertheless, our study strongly suggests that targeting Arg-II has a wide spectrum of direct and indirect effects on multiple cell types in aging heart including macrophages, fibroblasts, and endothelial cells, and cardiomyocytes, resulting in improved cardiac resistance to ischemia/reperfusion injury, which is due to inhibition of inflammation, fibrosis, and cell apoptosis. Thus, Arg-II shall represent a promising therapeutic target in treatment of age-related cardiac dysfunction and protect heart under diseased conditions.

Fig. 11.

Materials and methods

Arg-II^−/− mouse and sample preparation

Wild type (wt) and arg-ii knockout (arg-ii^−/−) mice were kindly provided by Dr. William O’Brien (Shi et al., 2001) and back crossed to C57BL/6 J for more than 10 generations. Genotypes of mice were confirmed by polymerase chain reaction (PCR) as previously described (Ming et al., 2012). Offspring of wt and arg-ii^−/−mice were generated by interbred from hetero/hetero cross. Mice were housed at 23°C with a 12-h-light-dark cycle. Animals were fed a normal chow diet and had free access to water and food. The animals were selected and included in the study according to age, gender, and genotype. Only healthy animals, based on scoring of activity, posture, general appearance (coat, skin), locomotion, eyes/nose, body weight loss, were used. In this study, confounders such as the animal/cage location were controlled. All experimenters were aware of the group allocation at the different stages of the experiments. Male and female mice at the age of 3–4 months (young) or 20–22 months (old) were euthanized under deep anesthesia (i.p. injection of a mixture of ketamine/xylazine 50 mg/kg and 5 mg/kg, respectively) and death was confirmed by absence of all the reflexes and by exsanguination. The heart was then isolated and either snap frozen in liquid nitrogen and kept at −80 °C until use or fixed with 4% paraformaldehyde (pH 7.0), and then embedded in paraffin for immunofluorescence staining experiments. Experimental work with animals was approved by the Ethical Committee of the Veterinary Office of Fribourg Switzerland (2020-01-FR) and performed in compliance with guidelines on animal experimentation at our institution and in accordance with the updated ARRIVE guidelines (Percie du Sert et al., 2020). Experiments with human heart biopsies were approved by the Ethics Committee of Northwestern and Central Switzerland (Project-ID 2021-01445). Rat heart tissues were obtained from previous study (Borrego et al., 2022) approved by the Ethical Committee of the Veterinary Office of Fribourg Switzerland (2019-21-FR).

Langendorff-perfused mouse heart preparation

Ex vivo functional assessments were performed by using an isolated heart system (Langendorff apparatus – EMKA technologies, France). Briefly, after euthanasia as above described, the hearts of wt and arg-ii^−/− male and female mice (20–22 months old) were excised, cannulated and retrogradely perfused on the Langendorff system. 150 µL of heparin (2,000 units/ml) was injected 15 minutes before the heart removal to prevent blood coagulation. Hearts trimmed of extracardiac tissue were weighed immediately after surgical removal prior cannulation and heart weight to body weight ratio (HW/BW) was calculated as hypertrophy index. The aorta was cannulated on a shortened and blunted 21-gauge needle and perfusion initiated at a constant pressure of 80 mmHg. The hearts were perfused with a modified Krebs–Henseleit solution composed of (mmol/L): 118 NaCL, 4.7 KCl, 1.2 MgSO₄, 1.2 KH₂PO₄, 25 NaHCO₃, 11 Glucose, 2 CaCl₂. The fluid was bubbled with a mix of 95% O₂ and 5% CO₂ at 37°C to give a pH of 7.4 and was filtered through an in-line 0.45 μm Sterivex-HV filter before delivery to the heart. A fluid-filled balloon was introduced into the left ventricle through an incision in the left atrial appendage. The ventricular balloon was connected via fluid-filled tubing to a pressure transducer for continuous assessment of ventricular function. The balloon was inflated to yield a left ventricular end-diastolic pressure between 5 to 15 mmHg. Hearts were immersed in warmed perfusate in a jacketed bath maintained at 37°C, and perfusate delivered to the coronary circulation was maintained at the same temperature. After 20 minutes stabilization, hearts were subjected to 15 minutes (males), 20 minutes and 30 minutes (females) of no-flow ischemia followed by 30-45 minutes of reperfusion. Recovery of contraction and relaxation rates (dt/dp_min) as well as the recovery of the left ventricle developed pressure (LV-DP) and left ventricular end diastolic pressure (LV-EDP) were measured to investigate tolerance to ischemia/reperfusion (I/R)-injury. In addition to functional recovery, triphenyl tetrazolium chloride (TTC) staining was also performed to investigate a potential reduction in infarct size upon I/R-injury in arg-ii^−/− animals. TTC is a colorless water-soluble dye that is reduced by the mitochondrial enzyme succinate dehydrogenase of living cells into a water-insoluble, light sensitive compound (formazan) that turns healthy/normal tissue deep red. In contrast, damaged/dead tissue remains white showing the absence of living cells, and thereby indicating the infarct region (Bederson et al., 1986). After completing the I/R protocol, heart was carefully removed from the experimental set-up. After a freeze-thaw cycle (−20°C for 1 hour), the heart was cross-sectioned from the apex to the atrioventricular groove into four 2.5 mm thick slices. The slices were first incubated in TTC (20 minutes at 37°C), and then fixed in 10% formalin for 30 minutes. A cover glass was then placed over the tissues and images were acquired (Redfors et al., 2012). ImageJ software was used to analyze the images and assess the infarcted area.

Real-time quantitative RT-PCR

mRNA expression level of genes from mouse and human origin was measured by two-step quantitative Real Time-PCR as described previously (Ming et al., 2012). Ribosomal Protein S12 (rps12), and glyceraldehyde-3-phosphate dehydrogenase (gapdh) were used as reference genes. Total RNA from mouse tissue, splenic macrophages, and human cardiac fibroblasts was extracted with Trizol Reagent (TR-118, Molecular Research Center) following the manufacturer’s protocol. Real-time PCR reaction was performed with the GOTaq® qPCR Master Mix (A6001, Promega) and CFX96 Real-Time PCR Detection System (Bio-Rad). The mRNA expression level of all genes was normalized to the reference gene rps12 or gapdh. All the qRT-PCR primer sequences are shown in the Suppl. Table 2.

Confocal immunofluorescence microscope

Immunofluorescence staining was performed with heart tissues. Briefly, mouse hearts or human cardiac biopsies were fixed with 4% paraformaldehyde (pH 7.0) and embedded in paraffin. After deparaffinization in xylene (2 times of 10 minutes), the sections were treated in ethanol (twice in 100% ethanol for 3 minutes and twice in 95% ethanol and once in 80%, 75%, 50% ethanol for 1 minute sequentially) followed by antigen retrieval (EDTA buffer, pH 8.0) for Arg-II, troponin-T (TropT), CD31, p16, Mac-2, IL-1β, vimentin (Vim), CD-68, PDGF-Rα and α-smooth muscle actin (αSMA) in a pressure cooker. For co-immunofluorescence staining of Arg-II/TropT, Arg-II/CD31, Arg-II/Vim, Arg-II/CD-68, Arg-II/Mac-2, Arg-II/PDGF-Rα, Arg-II/αSMA, IL-1β/Mac-2, F4-80/CCR2, F4-80/LYVE1 (Suppl. Fig. 10 shows the validation of CCR2 and LYVE1 antibodies) and CD31/Vim, primary antibodies of different species were used. Transverse sections (5 μm) were blocked with mouse Ig blocking reagent (M.O.M, Vector laboratories) for 2 hours and then with PBS containing 1% BSA and 10% goat serum for 1 hour. For CD31 and PDGF-Rα (both polyclonal Goat IgGs), a blocking reagent containing 10% BSA was used. The sections were then incubated overnight at 4°C in a dark/humidified chamber with target primary antibodies, and subsequently incubated for 2 hours with the following secondary antibodies: Alexa Fluor 488–conjugated goat anti-rabbit IgG (H + L) and Alexa Fluor 568-conjugated goat anti-mouse IgG (H + L); Alexa Fluor 488–conjugated goat anti-mouse IgG (H + L) and Alexa Fluor 594-conjugated goat anti-rabbit IgG (H + L); Alexa Fluor 488–conjugated donkey anti-goat IgG (H + L) and Alexa Fluor 594-conjugated goat anti-rabbit IgG (H + L). All sections were finally counterstained with 300 nmol/L DAPI for 5 minutes. Immunofluorescence signals were visualized under Leica TCS SP5 confocal laser microscope. The antibodies used are shown in Suppl. Table 3.

Isolation and cultivation of ventricular cardiomyocytes

Ventricular myocytes were isolated from five-month old female mice according to an established protocol (Louch et al., 2011). Briefly, the hearts were excised, cannulated and retrogradely perfused on a Langendorff system as described above. 150 µl of heparin (2,000 units/ml) was injected 15 minutes before the heart removal to prevent blood coagulation. To isolate the cardiomyocytes, hearts were perfused at 37°C for 20 minutes with a Ca²⁺-free solution composed of (in mmol/ liter): 130 NaCl, 5 KCL, 0.5 NaH₂PO₄, 10 HEPES, 10 Glucose, 10 2,3 butanedione monoxime (BDM), 10 Taurine, 1 MgCl₂ (pH 7.4, adjusted with NaOH). Cells were enzymatically dissociated using a cocktail of collagenase type II (210 U/ml, Worthington), collagenase type IV (260 U/ml, Worthington) and protease type XIV (0.21 U/ml; Sigma-Aldrich). After isolation, cardiomyocytes were separated by gravity settling and Ca²⁺ was restored to a final concentration of 1 mmol/L. The cardiomyocytes were finally plated onto laminin-coated coverslips (5 μg/ml, Thermo Scientific) and cultured in the M-199 medium supplemented with bovine serum albumin (BSA; 0.1 % - A1470, Sigma-Aldrich), BDM (10 mmol/L), insulin, transferrin, selenium (ITS; I3146 - Sigma-Aldrich), CD lipid (11905-031 - Sigma-Aldrich) and penicillin / streptomycin mix (1 %).

Cell culture, adenoviral transduction, and inos^−/− and arg-ii^−/− macrophages

Human cardiac fibroblasts (HCF) from adult human heart tissue were purchased from Innoprot (P10452) and cultured in the proprietary fibroblast culture medium (P60108-2, Innoprot) composed as follow: 500 ml of fibroblast basal medium, 25 ml of FBS, 5 ml of fibroblast growth supplement-2, and 5 ml of penicillin/streptomycin in the Poly-L-Lysine (PLL) coated flasks and dishes. The cells were maintained at 37°C in a humidified incubator (5% CO₂ and 95% atmosphere air). To overexpress arg-ii, the cells were seeded on 6-cm dishes for 24 hours and transduced with rAd-CMV-Con/arg-ii for 48 hours as described previously (Ming et al., 2012).

Human umbilical vein endothelial cells (HUVEC) cells were cultured in RPMI-1640 (with 25 mmol/L HEPES and stable glutamine; Thermo Fisher Scientific) containing 5 % fetal calf serum (FCS; Gibco) and 1% streptomycin and penicillin in gelatin (1%) coated dishes.

Raw 264.7 cells (mouse macrophage cell line) were cultured in Dulbecco modified Eagle medium/F12 (DMEM/F12; Thermo Fisher Scientific) containing 10% fetal bovine serum (FBS; Gibco) and 1% streptomycin and penicillin. To silence arg-ii, the cells were seeded on 6-cm dishes for 24 hours and transduced first with the rAd at titers of 100 Multiplicity of Infection (MOI) and cultured in the complete medium for 2 days and then in serum-free medium for another 24 hours before experiments. The rAd expressing shRNA targeting arg-ii driven by the U6 promoter (rAd/U6-arg-ii-shRNA) and control rAd expressing shRNA targeting lacz (rAd/U6-lacz-shRNA) were generated as described previously (Ming et al., 2009).

Wt and inos gene deficient bone-marrow-derived macrophage cell lines (MØ^wt and MØ^inos-/-) were purchased from Kerafast (ENH166-FP and ENH176-FP) and cultured in Dulbecco modified Eagle medium (DMEM; Thermo Fisher Scientific) containing 10% fetal bovine serum (FBS; Gibco) and 1% streptomycin and penicillin. The cells were treated with LPS (100 ng/mL) for 24 hours before harvest. Gene deficiency of inos was verified by immunoblotting analysis.

Knockout of arg-ii gene in human THP-1 cells was generated by CRISPR-UTM-mediated genome engineering with gRNA targeting exon 1 (Guangzhou Ubigene Biosciences Co., Ltd.). The control THP1 cell line was generated with scramble gRNA and Cas9. Gene deficiency of arg-ii was verified by immunoblotting analysis.

TUNEL Staining

Mouse hearts were isolated and fixed with 4% paraformaldehyde (pH 7.0), embedded in paraffin and sliced at 5 μm of thickness. After deparaffinization (xylene, 2 times for 10 minutes) and rehydration (sequential washing in 100%, 95%, 80%, 75% and 50% ethanol), TUNEL staining was performed using a commercial kit (in situ Cell Death Detection Kit, TMR red – Roche, Switzerland). Briefly, 50 μl of TUNEL mixture, containing TdT and dUTP in reaction buffer were added onto the sections and incubated in a humidified chamber for 60 minutes at 37°C in darkness. Positive controls were generated by incubating samples at room temperature (RT) for 10 minutes with DNase in PBS (1 mg/ml) to induce strand breaks. Negative controls were generated by incubation with TUNEL mixture lacking TdT. DAPI counterstaining was finally performed (300 nmol/l for 5 minutes).

TUNEL staining was also performed on isolated ventricular cardiomyocytes. After tissue digestion, the cardiomyocytes were plated onto murine laminin-coated coverslips to promote cell adhesion. One hour after plating, several PBS washing steps were performed to remove damaged and partially adherent cells, ensuring that only well-shaped, viable, and strongly adherent cells remained as bioassay cells. These “healthy” cells were then selected for the experiments. After washing, cardiomyocytes were incubated for 24 hours with either splenic macrophage conditioned media (CM; see below) or with RAW cells CM, and the following protocol was applied: cells are fixed using 4% paraformaldehyde for 1 hour at RT, and then permeabilized via Triton X (0.1 %). As for tissues, 50 μl of TUNEL mixture, containing TdT and dUTP in reaction buffer were added onto each slide, and then incubated at 37°C for 60 minutes in darkness. DAPI counterstaining was then performed.

Upon plating onto murine laminin-coated coverslips, the cardiomyocytes were first incubated for 24 hours with either splenic macrophage conditioned media (CM; see below) or with RAW cells CM, and the following protocol was applied: cells are fixed using 4% paraformaldehyde for 1 hour at RT, and then permeabilized via Triton X (0.1 %). As for tissues, 50 μl of TUNEL mixture, containing TdT and dUTP in reaction buffer were added onto each slide, and then incubated at 37°C for 60 minutes in darkness. DAPI counterstaining was then performed.

For experiments aiming to define cell borders, TUNEL-protocol was followed by a staining with Wheat Germ Agglutinin (WGA)-Alexa Fluor 488 (10 μg/ml for 30 minutes). For heart tissue, five representative images of each section are taken using a Leica TCS SP5 confocal system, and WGA staining was performed to differentiate cardiomyocytes and non-cardiomyocytes according to cell size and presence or absence of striations (characteristics of cardiomyocytes). Data was reported as number of TUNEL-positive nuclei over the total number of nuclei in the images.

Masson’s trichrome staining

To analyze cardiac fibrosis, heart sections (5 µm) were subjected to Masson’s trichrome (ab150686, Abcam) staining according to the manufacturer’s instructions (de Guzman et al., 2023).

Hydroxyproline colorimetric assay

Collagen content was analyzed by determination of hydroxyproline levels in left ventricles or in cell homogenates using the Hydroxyproline Assay kit (MAK008, Sigma) according to the manufacturer’s instructions.

Isolation of splenic macrophages

Splenic cells were isolated as previously described (Wang et al., 2013). Briefly, young and old wt and arg-ii^−/− female mice were euthanized as described and spleens were dissected from abdominal cavity and filtered through a 70-μm nylon strainer. Red blood cell lysis buffer was used to remove red blood cells. The cell suspension obtained was plated onto gelatin-coated dishes at a density of 2 × 10⁶ cells/ml. After 3 days, cells were washed to ensure macrophage enrichment. Following washing, the macrophages were cultured for further 4 days in RPMI culture media with 20% of L929 cell line conditioned medium (source of the macrophage growth factor M-CSF) to allow cell maturation. Medium was changed every two days.

Crosstalk between macrophages and heart cells

For crosstalk experiments of splenic macrophages from wt and arg-ii^−/− mice with cardiac cells, conditioned medium from the isolated splenic macrophages was collected and transferred to cardiomyocytes, fibroblasts and endothelial cells in 1:1 dilution with the specific cell culture medium for the indicated time. To study effects of IL-1β in the conditioned medium of splenic macrophages (CM-SM), cardiac cells, fibroblasts or endothelial cells were pre-treated with IL-1 receptor antagonist IL-1ra (50 ng/ml) for 2 hours followed by the specific experimental protocols for analysis of cardiac apoptosis, fibroblast activation, and EndMT transition, respectively.

For the experiments with mouse macrophage cell line Raw 264.7, the cells were seeded on six-well plates with a density of 2 × 10⁵ cells per well. Upon transduction with rAd expressing arg-ii^shRNA (see above) and over-night serum starvation, the mouse macrophages were polarized toward a pro-inflammatory secretion profile by stimulation with lipopolysaccharide (LPS; 100 ng/mL) for 24 hours. The conditioned medium (CM) from non-treated control and LPS-activated RAW cells (referred to as CM-RAW) was then filtered and transferred onto cardiomyocytes for 24 hours to investigate cell apoptosis. To study effects of IL-1β in CM-RAW on cardiac cells apoptosis, myocytes were pre-treated with IL-1 receptor antagonist IL-1ra (280-RA, R&D systems) for 2 hours.

Hypoxia experiments with murine cardiac fibroblasts and cardiomyocytes

Cardiac cells were isolated from wt mice according to an established protocol (Louch et al., 2011). Briefly, after euthanasia of mice, hearts were excised, cannulated and retrogradely perfused on a Langendorff system. Cells were enzymatically dissociated using a cocktail of collagenase type II, collagenase type IV and protease type XIV. After isolation, cardiomyocytes were separated from the pull of cardiac cells by gravity settling and plated onto laminin-coated coverslips. After 2 hours, the cardiomyocytes were washed to remove non-adhering cells and incubated for 24 hours at 1% oxygen level in a Coy In Vitro Hypoxic Cabinet System (The Coy Laboratory Products, Grass Lake, MI USA). The rest of cardiac cells were plated on 12-well plates and cultured in DMEM (10% FBS) for 3 days, allowing cardiac fibroblasts to adhere. The fibroblasts were passaged once and subjected to hypoxia as for the cardiomyocytes.

Hypoxia experiments with human cardiac fibroblasts and mitochondrial superoxide detection (MitoSOX staining)

Human cardiac fibroblasts (HCF) from adult human heart tissue were incubated for 48 hours at 1% oxygen level in a Coy In Vitro Hypoxic Cabinet System (The Coy Laboratory Products, Grass Lake, MI USA). To silence arg-ii, HCFs were seeded on 6-cm dishes for 24 hours and transduced first with the rAd at titers of 100 Multiplicity of Infection (MOI) and cultured in the complete medium for 2 days and then in serum-free medium for another 24 hours before experiments. The rAd expressing shRNA targeting arg-ii driven by the U6 promoter (rAd/U6-arg-ii-shRNA) and control rAd expressing shRNA targeting lacz (rAd/U6-lacz-shRNA) were generated as described previously (Ming et al., 2009). Mitochondrial superoxide generation was monitored by using MitoSOX. Briefly, the cells were incubated with MitoSOX at the concentration of 1 μmol/L for 30 minutes. After washing, the cells were then fixed with 3.7% paraformaldehyde followed by counterstaining with DAPI and then subjected to imaging through 40× objectives with Leica TCS SP5 confocal laser microscope. To ensure the mitochondria as source of the ROS, cells were treated with Mito TEMPO (10 μmol/L, 1 hour) followed by MitoSOX staining.

ELISA for IL-1β detection in conditioned media

IL-1β concentrations in the conditioned medium from murine splenic cells were measured by ELISA kits (mouse IL-1β, 432604, BioLegend, San Diego, USA) according to the manufacturer’s instructions. IL-1β concentrations in the conditioned medium from THP1 human cells were measured by ELISA kits (human IL-1β, 437004, BioLegend, San Diego, USA) according to the manufacturer’s instructions.

Immunoblotting

Heart tissue or cell lysate preparation, SDS-PAGE and immunoblotting, antibody incubation, and signal detection were performed as described previously (Ming et al., 2012). To prepare heart homogenates, frozen cardiac tissues were crushed into the fine powder using a mortar and pestle in liquid nitrogen on ice. A portion of the fine powder was then homogenized (XENOX-Motorhandstück MHX homogenizer) on ice in 150 µl of ice-cold lysis buffer with the following composition: 10 mmol/L Tris-HCl (pH 7.4), 0.4 % Triton X-100, 10 μg/ml leupeptin, and 0.1 mmol/l phenylmethylsulfonyl fluoride (PMSF), protease inhibitor cocktail (B14002, Bio-Tool) and phosphatase inhibitor cocktail (B15002; Bio-Tool). Homogenates were centrifuged (Sorvall Legend Micro 17R) at 13,000 × g for 15 minutes at 4 °C. Protein concentrations of supernatants were determined by Lowry method (500-0116, Bio-Rad). Equal amount of protein from each sample was heated at 75°C for 15 minutes in loading buffer and separated by SDS-PAGE electrophoresis. Proteins in the SDS-PAGE gel were then transferred to PVDF membranes which are blocked with PBS-Tween-20 supplemented with 5% skimmed milk. The membranes were then incubated with the corresponding primary antibody overnight at 4°C with gentle agitation. After washing with blocking buffer, the membranes were then incubated with the corresponding anti-mouse or anti-rabbit secondary antibody. Signals were visualized using the Odyssey Infrared Imaging System (LI-COR Biosciences), or the FUSION FX Imaging system (Witec AG) for chemiluminescence, and quantified by Image Studio Lite (5.2, LI-COR Biosciences). The antibodies used in this study are listed in the Suppl. Table 3.

Statistical analysis

Data were presented as mean ± SD. Data distribution was determined by Kolmogorov-Smirnov test and statistical analysis for normally distributed values was performed with Student’s unpaired t-test or analysis of variance (ANOVA) with Bonferroni post hoc test. For non-normally distributed values, Mann–Whitney test or the Kruskal–Wallis test were used. Differences in mean values were considered statistically significant at a two tailed p-value ≤ 0.05.

Availability of data and materials

Data supporting the present study are available from the corresponding author upon reasonable request.

Supplementary tables and figures

Suppl. Table 1.

Suppl Table 2.

Suppl. Table 3.

Suppl. Fig. 1.

Suppl. Fig. 2.

Suppl. Fig. 3.

Suppl. Fig. 4.

Suppl. Fig. 5.

Suppl. Fig. 6.

Suppl. Fig. 7.

Suppl. Fig. 8.

Suppl. Fig. 9.

Suppl. Fig. 10.

Original blots

Figure 4 blots.

Figure 5 blots.

Figure 6 blots.

Figure 7 blots.

Figure 8F blots.

Figure 8M blots.

Figure 9A blots.

Figure 9F blots.

Supplemental Figure 5 blots.

Supplemental Figure 6A blots.

Supplemental Figure 6D blots.

Additional information

Author contributions

DMP, XC, GA, AB, M-NG, AF, and SC performed experiments, data acquisition, analysis, and interpretation, KDM provided human samples; DMP, ZY and X-FM designed the project and prepared figures and drafted the manuscript; ZY and X-FM received funding for the study; All the authors discussed the results and critically revised and approved the manuscript for important intellectual contents.

Funding

This work was supported by grants from the Swiss National Science Foundation (31003A_179261/1 to ZY) and Swiss Heart Foundation (FF19033 to X-FM).

Statements relating to relevant ethics

Experimental work with animals was approved by the Ethical Committee of the Veterinary Office of Fribourg Switzerland (2020-01-FR, and 2019-21-FR) and performed in compliance with guidelines on animal experimentation at our institution. The study was conducted according to the principles expressed in the Declaration of Helsinki. Experiments with human heart biopsies were approved by the Ethics Committee of Northwestern and Central Switzerland (Project-ID 2021-01445).